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
0839_frame_FM1 Page 5 Tuesday, May 22, 2001 2:42 PM
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.
0839_frame_FM1 Page 6 Tuesday, May 22, 2001 2:42 PM
0839_frame_FM1 Page 7 Tuesday, May 22, 2001 2:42 PM
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.
0839_frame_FM1 Page 8 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 9 Tuesday, May 22, 2001 2:42 PM
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.
0839_frame_FM1 Page 10 Tuesday, May 22, 2001 2:42 PM
0839_frame_FM1 Page 11 Wednesday, May 23, 2001 10:39 AM
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
0839_frame_FM1 Page 12 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 13 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 14 Tuesday, May 22, 2001 2:42 PM
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.
0839_frame_FM1 Page 15 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 16 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 17 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 18 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 19 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 20 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 21 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 22 Wednesday, April 10, 2002 8:17 AM
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
0839_frame_FM1 Page 23 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 24 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 25 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 26 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 27 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 28 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 29 Wednesday, May 23, 2001 10:40 AM
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
0839_frame_FM1 Page 30 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 31 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 32 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 33 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 34 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 35 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 36 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 37 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 38 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 39 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 40 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 41 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 42 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 43 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 44 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 45 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 46 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 47 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 48 Tuesday, May 22, 2001 2:42 PM
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
0839_frame_FM1 Page 49 Tuesday, May 22, 2001 2:42 PM
Zoonoses ...............................................................................................1003 Acknowledgments ................................................................................1003 References .............................................................................................1004
Appendices Appendix A Conversions ...............................................................1011 Appendix B Abbreviations ............................................................1015 Appendix C Characteristics of Common Disinfectants ....1017 Index ...........................................................................................................1019
0839_frame_FM1 Page 50 Tuesday, May 22, 2001 2:42 PM
0839_frame_C01 Page 1 Tuesday, May 22, 2001 10:44 AM
I Emerging Pathways in Marine Mammal Medicine
0839_frame_C01 Page 2 Tuesday, May 22, 2001 10:44 AM
0839_frame_C01 Page 3 Tuesday, May 22, 2001 10:44 AM
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
3
0839_frame_C01 Page 4 Tuesday, May 22, 2001 10:44 AM
4
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).
0839_frame_C01 Page 5 Tuesday, May 22, 2001 10:44 AM
Marine Mammals as Sentinels of Ocean Health
5
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
0839_frame_C01 Page 6 Tuesday, May 22, 2001 10:44 AM
6
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C01 Page 7 Tuesday, May 22, 2001 10:44 AM
Marine Mammals as Sentinels of Ocean Health
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
0839_frame_C01 Page 8 Tuesday, May 22, 2001 10:44 AM
8
CRC Handbook of Marine Mammal Medicine
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).
0839_frame_C01 Page 9 Tuesday, May 22, 2001 10:44 AM
Marine Mammals as Sentinels of Ocean Health
9
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.
References Addison, R.F., and Smith, T.G., 1998, Trends in organochlorine residue concentrations in ringed seal (Phoca hispida) from Holman, Northwest Territories, 1972–91, Arctic, 51: 253–261. Addison, R.F., Stobo, W.T., and Zinck, M.E., 1998, Organochlorine residue concentrations in blubber of grey seal (Halichoerus grypus) from Sable Island, N.S. 1974–1994: Compilation of data and analysis of trends, Can. Data Rep. Fish. Aquat. Sci., 1043. Aguilar, A., and Borrell, A., 1994, Assessment of organochlorine pollutants in cetaceans by means of skin and hypodermic biopsies. Chapter 11, in Nondestructive Biomarkers in Vertebrates, Fossi, M.C., and Leonzio, C. (Eds.), Lewis Publishers, Boca Raton, FL, 245–267. Aguilar, A., and Raga, J.A., 1993, The striped dolphin epizootic in the Mediterranean Sea, Ambio, 22: 524–528. Andrews, B.F., Duffield, D.A., and McBain, J.F., 1997, Marine mammal management: Aiming at year 2000, IBI Rep., 7: 125–130. Au, W.W.L., Nachtigall, P.E., and Pawloski, J.L., 1999, Temporary threshold shift in hearing induced by an octave band of continuous noise in the bottlenose dolphin, J. Acoust. Soc. Am., 106: 2251. Borrell, A., and Aguilar, A., 1990, Loss of organochlorine compounds in the tissues of a decomposing stranded dolphin, Bull. Environ. Contam. Toxicol., 45: 46–53. Bowen, W.D., 1997, Role of marine mammals in aquatic ecosystems, Mar. Ecol. Prog. Ser., 158: 267–274. Brouwer, A., Reijnders, P.J.H., and Koeman, J.H., 1989, PCB-contaminated fish induces vitamin A and thyroid hormone deficiency in the common seal (Phoca vitulina), Aquat. Toxicol., 15: 99–106. Buergelt, C.D., and Bonde, R.K., 1983, Toxoplasmic meningoencephalitis in a West-Indian Manatee, J. Am. Vet. Med. Assoc., 183: 1294–1296. Burns, W.C.G., 2000, From the Harpoon to the Heat: Climate Change and the International Whaling Commission in the 21st Century, Pacific Institute for Studies in Development, Environment, and Security, Oakland, CA, 26 pp.
0839_frame_C01 Page 10 Tuesday, May 22, 2001 10:44 AM
10
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C01 Page 11 Tuesday, May 22, 2001 10:44 AM
Marine Mammals as Sentinels of Ocean Health
11
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.
0839_frame_C01 Page 12 Tuesday, May 22, 2001 10:44 AM
12
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C01 Page 13 Tuesday, May 22, 2001 10:44 AM
Marine Mammals as Sentinels of Ocean Health
13
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.
0839_frame_C01 Page 14 Tuesday, May 22, 2001 10:44 AM
0839_frame_C02 Page 15 Tuesday, May 22, 2001 10:40 AM
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
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
15
0839_frame_C02 Page 16 Tuesday, May 22, 2001 10:40 AM
16
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C02 Page 17 Tuesday, May 22, 2001 10:40 AM
17
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)
0839_frame_C02 Page 18 Tuesday, May 22, 2001 10:40 AM
18
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C02 Page 19 Tuesday, May 22, 2001 10:40 AM
Emerging and Resurging Diseases
19
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
0839_frame_C02 Page 20 Tuesday, May 22, 2001 10:40 AM
20
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C02 Page 21 Tuesday, May 22, 2001 10:40 AM
21
Emerging and Resurging Diseases
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
0839_frame_C02 Page 22 Tuesday, May 22, 2001 10:40 AM
22
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C02 Page 23 Wednesday, May 23, 2001 10:41 AM
Emerging and Resurging Diseases
23
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
0839_frame_C02 Page 24 Tuesday, May 22, 2001 10:40 AM
24
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C02 Page 25 Tuesday, May 22, 2001 10:40 AM
Emerging and Resurging Diseases
25
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.
0839_frame_C02 Page 26 Tuesday, May 22, 2001 10:40 AM
26
CRC Handbook of Marine Mammal Medicine
Bossart, G.D., Decker, S.J., and Ewing, R.Y., in press, Cytopathology of cutaneous viral papillomatosis in the killer whale (Orcinus orca), in Molecular and Cell Biology of Marine Mammals, Pfeiffer, C.J. (Ed.), Krieger, Melbourne, FL. Bowie, W.R., King, A.S., Werker, D.H., Isaac-Renton, J.L., Bell, A., Eng, S.B., and Marion, S.A., 1997, Outbreak of toxoplasmosis associated with municipal drinking water, The BC Toxoplasma Investigation Team, Lancet, 350: 173–177. Bricker, B.J., Ewalt, D.R., MacMillan, A.P., Foster, G., and Brew, S., 2000, Molecular characterization of Brucella strains isolated from marine mammals, J. Clin. Microbiol., 38: 1258–1262. Buckles, E.L., Lowenstine, L.J., King, D.P., Stott, J.L., Garber, R., Spraker, T., Lipscomb, T., Haulena, M., and Gulland, F.M.D., 1999, Current investigations into the etiology and pathogenesis of neoplasms in California sea lions (Zalophus californianus), in Proceedings of the 30th International Association for Aquatic Animal Medicine Annual Conference, Boston, MA, 30: 99–101. Cassonnet, P., Van Bressem, M.F., Desaintes, C., Van Waerebeek, K., and Orth, G., 1999, Papillomavirus causes genital warts in small cetaceans from Peru, European Research on Cetaceans-12, European Cetacean Society, 12th Annual Conference Proceedings, Monaco, January 1998, 349. Chechowitz, M.A., Lowenstine, L.J., Gardner, I., Barr, B.C., Conrad, P.A., Gulland, F.M., and Jessup, D., 1999, Protozoal encephalitis in California sea otters and harbor seals: An update, in Proceedings of the 30th International Association of Aquatic Animal Medicine Annual Conference, Boston, MA, 30: 5. Clavareau, C., Wellemans, V., Walravens, K., Tryland, M., Verger, J.M., Grayon, M., Cloeckaert, A., Letesson, J.J., and Godfroid, J., 1998, Phenotypic and molecular characterization of a Brucella strain isolated from a minke whale (Balaenoptera acutorostrata), Microbiology, 144: 3267–3273. Cole, R., Lindsay, D.S., Howe, D.K., Roderick, C.L., Dubey, J.P., Thomas, N.J., and Baeten, L.A., 2000, Biological and molecular characterizations of Toxoplasma gondii strains obtained from southern sea otters (Enhydra lutris nereis), J. Parasitol., 86: 526–530. Cowan, D.F., and Turnbull, B.S., 1999, Renal neoplasms in the Atlantic bottlenose dolphin (Tursiops truncatus) from the western coast of the gulf of Mexico, presented at 13th Biennial Conference on the Biology of Marine Mammals, Wailea, Maui, HI, 39. 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. Dailey, M.D., and Brownell, R.L., 1972, A checklist of marine mammal parasites, in Mammals of the Sea, Biology and Medicine, Ridgway, S. (Ed.), Charles C Thomas, Springfield, IL, 528–589. Dailey, M.D., and Mayer, K., 1999, Parasitic helminth (Acanthocephalan) infection as a cause of mortality in the California sea otter (Enhydra lutris), in Proceedings of the 30th International Association for Aquatic Animal Medicine Annual Conference, Boston, MA, 30: 126–127. De Guise, S., Lagacé, A., and Béland, P., 1994, Gastric papillomas in eight St. Lawrence beluga whales (Delphinapterus leucas), J. Vet. Diagn. Invest., 6: 385–388. De Koeijer, A., Diekmann, O., and Reijnders, P., 1998, Modelling the spread of phocine distemper virus among harbour seals, Bull. Math. Biol., 60: 585–596. Deng, M.Q., Peterson, R.P., and Cliver, D.O., 2000, First findings of Cryptosporidium and Giardia in California sea lions (Zalophus californianus), J. Parasitol., 86: 490–494. Domingo, M., Ferrer, L., Pumarola, M., Marco, A., Plana, J., Kennedy, S., McAliskey, M., and Rima, B. K., 1990, Morbillivirus in dolphins, Nature, 348: 21. Duignan, P.J., Saliki, J.T., St. Aubin, D.J., House, J.A., and Geraci, J.R., 1994, Neutralizing antibodies to phocine distemper virus in Atlantic walruses (Odobenus rosmarus rosmarus) from Arctic Canada, J. Wildl. Dis., 30: 90–94. Duignan, P., House, C., Geraci, J.R., Duffy, N., Rima, B.K., Walsh, M.T., St. Aubin, D.J., Sadove, S., and Koopman, H., 1995a, Morbillivirus infection in cetaceans of the western Atlantic, Vet. Microbiol., 44: 241–249.
0839_frame_C02 Page 27 Tuesday, May 22, 2001 10:40 AM
Emerging and Resurging Diseases
27
Duignan, P., House, C., Geraci, J.R., Early, G., Copland, H.G., and Walsh, M.T., 1995b, Morbillivirus infection in two species of pilot whales (Globecephala sp.) from the western Atlantic, Mar. Mammal Sci., 11: 150–162. Duignan, P.J., Nielsen, O., House, C., Kovacs, K.M., Duffy, N., Early, G., Sadove, S., St. Aubin, D.J., Rima, B.K., and Geraci, J.R., 1997, Epizootiology of morbillivirus infection in harp, hooded, and ringed seals from the Canadian Arctic and western Atlantic, J. Wildl. Dis., 33: 7–19. Ewing, R., Bossart, G.D., and Lowe, M., 1997, Cutaneous viral papillomatosis in a West Indian Manatee (Trichechus manatus latirostris), presented at 46th Annual Wildlife Disease Association Conference, St. Petersburg, FL, 32. Ewing, R.Y., and Mignucci-Giannoni, A.A., in review, Stranded free-ranging offshore Atlantic bottlenose dolphin (Tursiops truncatus) with a poorly differentiated pulmonary squamous cell carcinoma, J. Vet. Diag. Invest. Fauquier, D., Gulland, F., Haulena, M., and Lowenstine, L., 1998, Northern fur seal (Callorhinus ursinus) strandings along the central California coast over twenty-two years, 1975–1997, in Proceedings of the 29th International Association for Aquatic Animal Medicine Annual Conference, San Diego, CA, 39. Fletcher, D., Gulland, F.M.D., Haulena, M., Lowenstine, L.J., and Dailey, M., 1998, Nematode-associated gastrointestinal perforations in stranded California sea lions (Zalophus californianus), in Proceedings of the 29th International Association for Aquatic Animal Medicine Annual Conference, San Diego, CA, 59. Follmann, E.H., Garner, G.W., Everman, J.F., and McKeirnan, A.J., 1996, Serological evidence of morbillivirus infection in polar bears (Ursus maritimus) from Alaska and Russia, Vet. Rec., 22: 615–618. Forbes, L.B., Nielsen, O., Measures, L., and Ewalt, D.R., 2000, Brucellosis in ringed seals and harp seals from Canada, J. Wildl. Dis., 36: 595–598. Forsyth, M.A., Kennedy, S., Wilson, S., Eybatov, T., and Barrett, T., 1998, Canine distemper virus in a Caspian seal, Vet. Rec., 143: 662–664. Foster, G., Jahans, K.L., Reid, R.J., and Ross, H.M., 1996, Isolation of Brucella species from cetaceans, seals and an otter, Vet. Rec., 138: 583–586. Fox, J.G., Harper, C.M.G., Dangler, C.A., Xu, S., Stamper, A., and Dewhirst, F.E., 2000, Isolation and characterization of Helicobacter sp. from the gastric mucosa of dolphins, in American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine Joint Conference Proceedings, New Orleans, LA, Sept. 17–24. Garner, M.M., Barr, B.C., Pockham, A.E., Marsh, A.E., Burek-Huntington, K.A., Wilson, R.K., and Dubey, J.P., 1997a, Fatal hepatic sarcocystosis in two polar bears (Ursus maritimus), J. Parasitol., 83: 523–526. 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., 1997b, Evidence of Brucella infection in Parafilaroides lungworms in a Pacific harbor seal (Phoca vitulina richardsi), J. Vet. Diagn. Invest., 9: 298–303. 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, National Marine Fisheries Service, U.S. Department of Commerce, Silver Spring, MD, 45 pp. Gulland, F.M., Lowenstine, L.J., Lapointe, J.M., Spraker, T., and King, D.P., 1997, Herpesvirus infection in stranded Pacific harbor seals of coastal California, J. Wildl. Dis., 33: 450–458. Gulland, F.M.D., Trupkiewictz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996, Metastatic carcinoma of probable transitional cell origin in 66 free-living California sea lions (Zalophus californianus), 1979 to 1994, J. Wildl. Dis., 32: 250–267. Gulland, F.M.D., Trupkiewicz, J.G., Spraker, T., Lowenstine, L.J., Stein, J., Tilbury, K.L., Reichert, W.L., and Hom, T., 1995, Metastatic carcinoma and exposure to chemical contaminants in California sea lions (Zalophus californianus) stranded along the central California coast, presented at 11th Biennial Conference on the Biology of Marine Mammals, Dec. 14–18, Orlando, FL, 48.
0839_frame_C02 Page 28 Tuesday, May 22, 2001 10:40 AM
28
CRC Handbook of Marine Mammal Medicine
Harder, T.C., Harder, M., Vos, H., Kulonen, K., Kennedy-Stoskopf, S., Liess, B., Appel, M.J.G., and Osterhaus, A.D.M.E., 1996, Characterization of phocid herpesvirus-1 and -2 as putative alpha and gamma herpesviruses of North American and European pinnipeds, J. Gen. Virol., 77: 27–35. Harvell, C.D., Kim, K., Burkholder, J.M., Colwell, R.R., Epstein, P.R., Grimes, 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. 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. Haubold, E.M., Aronson, J.F., Cowan, D.F., McGinnis, M.R., and Cooper, C.R., 1998, Isolation of fungal rDNA from bottlenose dolphin skin infected with Loboa loboi, Med. Mycol., 36: 263–267. Hernández, M., Robinson, I., Aguilara, A., González, L.M., López-Jurado, L.F., Reyero, M.I., Cacho, E., Franco, J., López-Rodas, V., and Costas, E., 1998, Did algal toxins cause monk seal mortality? Nature, 393: 28–29. Hoegh-Guldberg, O., 1999, Climate change, coral bleaching and the future of the world’s coral reefs, Mar. Freshwater Res., 50: 839–866. Howe, D.K., Honore, S., Derouin, F., and Sibley, L.D., 1997, Determination of genotypes of Toxoplasma gondii strains isolated from patients with toxoplasmosis, J. Clin. Microbiol., 35: 1411–1414. Hunter, J.E., Duignan, P.J., Dupont, C., Fray, L., Fenwick, S.G., and Murray, A., 1998, First report of potentially zoonotic tuberculosis in fur seals in New Zealand, N.Z. Med. J., 11: 130–131. Isaac-Renton, J.L., Bowie, W.R., King, A., Irwin, G.S., Ong, C.S., Fung, C.P., Shokeir, M.O., and Dubey, J.P., 1998, Detection of Toxoplasma gondii oocyst in drinking water, Appl. Environ. Microbiol., 64: 2278–2280. Jahans, K.L., Foster, G., and Broughton, E.S., 1997, The characterisation of Brucella strains isolated from marine mammals, Vet. Microbiol., 57: 373–382. Jauniaux, T., Charlier, G., Desmecht, M., and Coignoul, F., 1998, Lesions of morbillivirus infection in a fin whale (Balaenoptera physalus) stranded along the Belgian coast, Vet. Rec., 143: 423–424. Jensen, E.D., Lipscomb, T., Van Bonn, B., Miller, G., Fradkin, J.M., and Ridgway, S.H., 1998, Disseminated histoplasmosis in an Atlantic bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 456–460. Jepson, P.D., Brew, S., MacMillan, A.P., Baker, J.R., Barnett, J., Kirkwood, J.K., Kuiken, T., Robinson, I.R., and Simpson, V.R., 1997, Antibodies to Brucella in marine mammals around the coast of England and Wales, Vet. Rec., 141: 513–515. 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. Kennedy, S., 1990, A review of the 1988 European seal morbillivirus epizootic, Vet. Rec., 127: 563–567. Kennedy, S., 1998, Morbillivirus infection in aquatic mammals, J. Comp. Pathol., 119: 201–225. Kennedy, S., Kuiken, T., Jepson, P.D., Deaville, R., Forsyth, M., Barrett, T., van de Bildt, M.W.G., Osterhaus, A.D.M.E., Eybatov, T., Duck, C., Kydyrmanov, A., Mitrofanov, I., and Wilson, S., 2000, Mass die-off of Caspian seals caused by canine distemper virus, Emerging Infect. Dis., 6: 637–639. Lambertsen, R.H., Kohn, B.A., Sundberg, J.P., and Buergelt, C.D., 1987, Genital papillomatosis in sperm whale bulls, J. Wildl. Dis., 23: 361–367. La Pointe, J.M., Duignan, P.J., March, A.E., Gulland, F.M., Barr, B.C., Naydan, D.K., Kang, D.P., Farman, C.A., Burek, K.A., and Lowenstine, L.J., 1998, Meningoencephalitis due to a Sarcocystis neurona– like protozoan in Pacific harbor seals (Phoca vitulina richardsi), J. Parasitol., 84: 1184–1189. La Pointe, J.M., Gulland, F.M., Haines, D.M., Barr, B.C., and Duignan, P.J., 1999, Placentitis due to Coxiella burnetii in a Pacific harbor seal (Phoca vitulina richardsi), J. Vet. Diagn. Invest., 11: 541–543. Lefebvre, K.A., Powell, C.L., Busman, M., Doucette, G.J., Moeller, P.D., Silver, J.B., Miller, P.E., Hughes, M.P., Singaram, S., Silver, M.W., and Tjeerdema, R.S., 1999, Detection of domoic acid in northern anchovies and California sea lions associated with an unusual mortality event, Nat. Toxins, 7: 85–92.
0839_frame_C02 Page 29 Tuesday, May 22, 2001 10:40 AM
Emerging and Resurging Diseases
29
Lindsay, D.S., Thomas, N.J., and Dubey, J.P., 2000, Isolation and characterization of Sarcocystis neurona from a southern sea otter (Enhydra lutris nereis), Int. J. Parasitol., 30: 617–624. Lipscomb, T.P., Schulman, F.Y., Moffett, D., and Kennedy, S., 1994, Morbilliviral disease in Atlantic bottlenose dolphins (Tursiops truncatus) from 1987–1988 epizootic, J. Wildl. Dis., 30: 567–571. Lipscomb, T.P., Scott, D.P., Garber, R., Krafft, A.E., Tsai, M.M., Lichy, J.H., Taubenberger, J.K., Schulman, F.Y., and Gullen, F.M.D., 2000, Common metastatic carcinoma of California sea lions (Zalophus californianus): Evidence of genital origin and association with novel gammaherpesvirus, Vet. Pathol., 37: 609–617. Lyons, C., Welsh, M.J., Thorsen, J., Ronald, K., and Rima, B.K., 1993, Canine distemper virus isolated from a captive seal, Vet. Rec., 132: 487–488. Mamaev, L.V., Denikina, N.N., Belikov, S.I., Volchkov, V.E., Visser, I.K., Fleming, M., Kai, C., Harder, T.C., Liess, B., and Osterhaus, A.D., 1995, Characterisation of morbilliviruses isolated from Lake Baikal seals (Phoca sibirica), Vet. Microbiol., 44: 2–4, 251–259. Measures, L.N., and Olson, M., 1999, Giardiasis in pinnipeds from eastern Canada, J. Wildl. Dis., 35: 779–782. Miller, W.G., Adams, L.G., Ficht, T.A., Cheville, N., Payeur, J.P., Harley, D.R., House, C., and Ridgway, S.H., 1999, Brucella-induced abortions and infection in bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med., 30: 100–110. Mondragon, R., Howe, D.K., Dubey, J.P., and Silbey, L.D., 1998, Genotypic analysis of Toxoplasma gondii isolates from pigs, J. Parasitol., 84: 639–641. Olson, M.E., Roach, P.D., Stabler, M., and Chan, W., 1997, Giardiasis in ringed seals from the western Arctic, J. Wildl. Dis., 33: 5646–5648. O’Shea, T.J., Rathbun, G.B., Bonde, R.K., Buergelt, D.D., and Odell, D.K., 1991, An epizootic of Florida manatees associated with a dinoflagellate bloom, Mar. Mammal Sci., 7: 165–179. Osterhaus, A., Groen, J., Niesters, H., van de Bildt, M., Martina, B., Vedder, 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. Osterhaus, A.D.M.E., Groen, J., Uyt de Haag, F.G.C.M., Visser, I.K.G., van de Bildt, M.G.W., Bergman, A. et al., 1989, Distemper in Baikal seals, Nature, 338: 209–210. Osterhaus, A.D.M.E., Rimmelzwaan, G.F., Martina, B.E.E., Bestebroer, T.M., and Fouchier, R.A.M., 2000, Influenza B virus in seals, Science, 288: 1051–1053. Osterhaus, A., van de Bildt, M., Vedder, L., Martina, B., Niesters, H., Vos, J., Van Egmond, H., Liem, D., Baumann, R., Androukaki, E., Kotomatas, S., Komnenou, A., Sidi, B.A., Jiddou, A.B., and Barham, M.E.O., 1998, Monk seal mortality: Virus or toxin? Vaccine, 16: 979–981. Reidarson, T.H., Griner, L.A., Pappagianis, D., and McBain, J., 1998a, Coccidioidomycosis in a bottlenose dolphin, J. Wildl. Dis., 34: 629–631. Reidarson, T.H., McBain, J., House, C., King, D.P., Stott, J.L., Krafft, A., Taubenberger, J.K., Heyning, J., and Lipscomb, T.P., 1998b, Morbillivirus infection in stranded common dolphins from the Pacific Ocean, J. Wildl. Dis., 34: 771–776. Renner, M.S., Ewing, R., Bossart, G.D., and Harris, D., 1999, Sublingual squamous cell carcinoma in an Atlantic bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 30: 573–576. Rosonke, B.J., Brown, S.R., Tornquist, S.J., Snyder, S.P., Garner, M.M., and Blythe, L.L., 1999, Encephalomyelitis associated with a Sarcocystis neurona–like organism in a sea otter, J. Am. Vet. Med. Assoc., 215: 1839–1842. Ross, P.S., Visser, I.K.G., Broeders, H.W.J., van de Bildt, M.W.G., Bowen, W.D., and Osterhaus, A.D.M.E., 1992, Antibodies to phocine distemper virus in Canadian seals, Vet. Rec., 130: 514–516. Scholin, C.A., Gulland, F., Doucetts, G.J., Benson, S., Busman, M., Chavez, F.P., Cordaro, J., DeLong, R., deVogalaere, 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.
0839_frame_C02 Page 30 Tuesday, May 22, 2001 10:40 AM
30
CRC Handbook of Marine Mammal Medicine
Stamper, M.A., Gulland, F.M.D., and Spraker, T., 1998, Leptospirosis in rehabilitated Pacific harbor seals from California, J. Wildl. Dis., 34: 407–410. Taubenberger, J.K., Tsai, M., Krafft, A.E., Lichy, J.H., Reid, A.H., Schulman, F.Y., and Lipscomb, T.P., 1996, Two morbilliviruses implicated in bottlenose dolphin epizootics, Emerging Infect. Dis., 2: 213–261. Taubenberger, J.K., Tsai, M.M., Atkin, T.J., Fanning, T.G., Krafft, A.E., Moaller, R.B., Kodsi, S.E., Mense, M.G., and Lipscomb, T.P., 2000, Molecular genetic evidence of a novel morbillivirus in a longfinned pilot whale (Globicephalus melas), Emerging Infect. Dis., 6: 42–45. Thomas, N.J., and Cole, R.A., 1996, The risk of disease and threats to the wild population, Endangered Species Update, 13: 23–27. Thornton, S.M., Nolan, S., and Gulland, F.M., 1998, Bacterial isolates from California sea lions (Zalophus californianus), harbor seals (Phoca vitulina), and northern elephant seals (Mirounga angustirostris) admitted to a rehabilitation center along the central California coast, 1994–1995, J. Zoo Wildl. Med., 29: 171–176. Tryland, M., Kleivane, L., Alfredsson, A., Kjeld, M., Arnason, A., Stuen, S., and Godfroid, J., 1999, Evidence of Brucella infection in marine mammals in the North Atlantic Ocean, Vet. Rec., 144: 588–592. Turnbull, B.S., and Cowan, D.F., 1999, Angiomatosis, a newly recognized disease in Atlantic bottlenose dolphins (Tursiops truncatus) from the Gulf of Mexico, Vet. Pathol., 36: 28–34. Uchida, K., Murananka, M., Horii, Y., Murakami, N., Yamaguchi, R., and Tateyama, S., 1999, Nonpurulent meningoencephalomyelitis of a Pacific striped dolphin in the Pacific Ocean around Japan, J. Vet. Med. Sci., 61: 159–162. U.S. Marine Mammal Commission, 1996, Annual Report to Congress, U.S. Marine Mammal Commission, Bethesda, MD, 6–18. Van Bressem, M.F., Van Waerebeek, K., Piérard, G.E., and Desaintes, C., 1996, Genital and lingual warts in small cetaceans from coastal Peru, Dis. Aquat. Organisms, 26: 1–10. Van Bressem, M.F., Van Waerebeek, K., Fleming, M., and Barrett, T., 1998, Serological evidence of morbillivirus infection in small cetaceans from the Southeast Pacific, Vet. Microbiol., 59: 89–98. Van Bressem, M.F., Kastelein, R.A., Flamant, P., and Orth, G., 1999, Cutaneous papillomavirus infection in a harbour porpoise (Phocoena phocoena) from the North Sea, Vet. Rec., 144: 592–593. Visser, I.K.G., Kumarev, V.P., Orvell, C., De Vries, P., Broeders, H.W.J., van de Bildt, M.W.G., Groen, J., Teppema, J.S., Burger, M.C., Uyt de Haag, F.G.C.M., and Osterhaus, A.D.M.E., 1990, Comparison of two morbilliviruses isolated from seals during outbreaks of distemper in North West Europe and Siberia, Arch. Virol., 111: 149–164. Visser, I.K.G., Van der Heuden, R.W.J., van de Bildt, M.W.G., Kenter, M.J.H., Orvell, C., and Osterhaus, A.D.M.E., 1993, Antigenic and F gene nucleotide sequence similarities, and phylogenetic analysis suggest that phocid distemper virus-2 and canine distemper virus belong to the same virus entity, in Morbillivirus Infections in Seals, Dolphins and Porpoises, Visser, I.K.G. (Ed.), Seal Rehabilitation and Research Centre, Pieterburen, the Netherlands, 47–59. Walz, P.M., Garrison, D.L., Graham, W.M., Cattey, M.A., Tjeerdema, R.S., and Silver, M.W., 1994, Domoic acid-producing diatom blooms in Monterey Bay, California: 1991–1993, Nat. Toxins, 2: 271–279. Wilson, M.E., 1999, Emerging infections and disease emergence, Emerging Infect. Dis., 5: 308–309. Work, T.M., Barr, B., Beale, A.M., Fritz, L., Quilliam, L.A., and Wright., J.L.C., 1993, Epidemiology of domoic acid poisoning in brown pelicans (Pelecanus occidentalis) and Brandt’s cormorants (Phalacrocorax penicillatus) in California, J. Zoo Wildl. Med., 24: 54–62.
0839_frame_C03 Page 31 Tuesday, May 22, 2001 10:40 AM
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
31
D
B
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.)
C
A
0839_frame_C03 Page 32 Tuesday, May 22, 2001 10:40 AM
32
0839_frame_C03 Page 33 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
33
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
0839_frame_C03 Page 34 Tuesday, May 22, 2001 10:40 AM
34
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.)
0839_frame_C03 Page 35 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
35
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.
0839_frame_C03 Page 36 Tuesday, May 22, 2001 10:40 AM
36
Handbook of Marine Mammal Medicine
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
0839_frame_C03 Page 37 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
37
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
0839_frame_C03 Page 38 Tuesday, May 22, 2001 10:40 AM
38
Handbook of Marine Mammal Medicine
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
0839_frame_C03 Page 39 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
39
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
0839_frame_C03 Page 40 Tuesday, May 22, 2001 10:40 AM
40
Handbook of Marine Mammal Medicine
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
0839_frame_C03 Page 41 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
41
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.
0839_frame_C03 Page 42 Tuesday, May 22, 2001 10:40 AM
42
Handbook of Marine Mammal Medicine
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.
0839_frame_C03 Page 43 Tuesday, May 22, 2001 10:40 AM
Florida Manatees: Perspectives on Populations, Pain, and Protection
43
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.
0839_frame_C03 Page 44 Tuesday, May 22, 2001 10:40 AM
0839_frame_C04.fm Page 45 Tuesday, May 22, 2001 10:41 AM
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.,
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
45
0839_frame_C04.fm Page 46 Tuesday, May 22, 2001 10:41 AM
46
CRC Handbook of Marine Mammal Medicine
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]
0839_frame_C04.fm Page 47 Tuesday, May 22, 2001 10:41 AM
47
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 48 Tuesday, May 22, 2001 10:41 AM
48
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C04.fm Page 49 Tuesday, May 22, 2001 10:41 AM
49
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 50 Tuesday, May 22, 2001 10:41 AM
50
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 51 Tuesday, May 22, 2001 10:41 AM
51
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 52 Tuesday, May 22, 2001 10:41 AM
52
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 53 Tuesday, May 22, 2001 10:41 AM
53
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 54 Tuesday, May 22, 2001 10:41 AM
54
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 55 Tuesday, May 22, 2001 10:41 AM
55
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 56 Tuesday, May 22, 2001 10:41 AM
56
CRC Handbook of Marine Mammal Medicine
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]
0839_frame_C04.fm Page 57 Tuesday, May 22, 2001 10:41 AM
57
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 58 Tuesday, May 22, 2001 10:41 AM
58
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C04.fm Page 59 Tuesday, May 22, 2001 10:41 AM
59
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 60 Tuesday, May 22, 2001 10:41 AM
60
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 61 Tuesday, May 22, 2001 10:41 AM
61
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 62 Tuesday, May 22, 2001 10:41 AM
62
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 63 Tuesday, May 22, 2001 10:41 AM
63
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 64 Tuesday, May 22, 2001 10:41 AM
64
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 65 Tuesday, May 22, 2001 10:41 AM
65
Marine Mammal Stranding Networks
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)
0839_frame_C04.fm Page 66 Tuesday, May 22, 2001 10:41 AM
66
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C04.fm Page 67 Tuesday, May 22, 2001 10:41 AM
Marine Mammal Stranding Networks
67
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.
0839_frame_C04.fm Page 68 Tuesday, May 22, 2001 10:41 AM
0839_frame_C05.fm Page 69 Tuesday, May 22, 2001 10:41 AM
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
69
0839_frame_C05.fm Page 70 Tuesday, May 22, 2001 10:41 AM
70
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C05.fm Page 71 Tuesday, May 22, 2001 10:41 AM
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
72
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
0839_frame_C05.fm Page 72 Tuesday, May 22, 2001 10:41 AM
CRC Handbook of Marine Mammal Medicine
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)
0839_frame_C05.fm Page 73 Tuesday, May 22, 2001 10:41 AM
Marine Mammal Unusual Mortality Events 73
0839_frame_C05.fm Page 74 Tuesday, May 22, 2001 10:41 AM
74
CRC Handbook of Marine Mammal Medicine
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?
0839_frame_C05.fm Page 75 Tuesday, May 22, 2001 10:41 AM
75
Marine Mammal Unusual Mortality Events
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.
0839_frame_C05.fm Page 76 Tuesday, May 22, 2001 10:41 AM
76
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C05.fm Page 77 Tuesday, May 22, 2001 10:41 AM
Marine Mammal Unusual Mortality Events
77
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.
0839_frame_C05.fm Page 78 Tuesday, May 22, 2001 10:41 AM
78
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C05.fm Page 79 Tuesday, May 22, 2001 10:41 AM
Marine Mammal Unusual Mortality Events
79
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.
0839_frame_C05.fm Page 80 Tuesday, May 22, 2001 10:41 AM
80
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C05.fm Page 81 Tuesday, May 22, 2001 10:41 AM
Marine Mammal Unusual Mortality Events
81
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.
0839_frame_C05.fm Page 82 Tuesday, May 22, 2001 10:41 AM
0839-frame_C06 Page 83 Tuesday, May 22, 2001 10:42 AM
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
83
0839-frame_C06 Page 84 Tuesday, May 22, 2001 10:42 AM
84
CRC Handbook of Marine Mammal Medicine
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
0839-frame_C06 Page 85 Tuesday, May 22, 2001 10:42 AM
85
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.
0839-frame_C06 Page 86 Tuesday, May 22, 2001 10:42 AM
86
CRC Handbook of Marine Mammal Medicine
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.
0839-frame_C06 Page 87 Tuesday, May 22, 2001 10:42 AM
87
Mass Strandings of Cetaceans
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
0839-frame_C06 Page 88 Tuesday, May 22, 2001 10:42 AM
88
CRC Handbook of Marine Mammal Medicine
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
0839-frame_C06 Page 89 Tuesday, May 22, 2001 10:42 AM
89
Mass Strandings of Cetaceans
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
0839-frame_C06 Page 90 Tuesday, May 22, 2001 10:42 AM
90
CRC Handbook of Marine Mammal Medicine
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)
0839-frame_C06 Page 91 Tuesday, May 22, 2001 10:42 AM
Mass Strandings of Cetaceans 91
0839-frame_C06 Page 92 Tuesday, May 22, 2001 10:42 AM
92
CRC Handbook of Marine Mammal Medicine
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)
0839-frame_C06 Page 93 Tuesday, May 22, 2001 10:42 AM
Mass Strandings of Cetaceans 93
0839-frame_C06 Page 94 Tuesday, May 22, 2001 10:42 AM
94
CRC Handbook of Marine Mammal Medicine
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.
0839-frame_C06 Page 95 Tuesday, May 22, 2001 10:42 AM
Mass Strandings of Cetaceans
95
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.
0839-frame_C06 Page 96 Tuesday, May 22, 2001 10:42 AM
96
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C07 Page 97 Tuesday, May 22, 2001 10:42 AM
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
97
0839_frame_C07 Page 98 Tuesday, May 22, 2001 10:42 AM
98
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C07 Page 99 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
99
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.
0839_frame_C07 Page 100 Tuesday, May 22, 2001 10:42 AM
100
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C07 Page 101 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
101
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?”
0839_frame_C07 Page 102 Tuesday, May 22, 2001 10:42 AM
102
CRC Handbook of Marine Mammal Medicine
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,
0839_frame_C07 Page 103 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
103
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
0839_frame_C07 Page 104 Tuesday, May 22, 2001 10:42 AM
104
CRC Handbook of Marine Mammal Medicine
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?
0839_frame_C07 Page 105 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
105
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.
0839_frame_C07 Page 106 Tuesday, May 22, 2001 10:42 AM
106
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C07 Page 107 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
107
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
0839_frame_C07 Page 108 Tuesday, May 22, 2001 10:42 AM
108
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C07 Page 109 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
109
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
0839_frame_C07 Page 110 Tuesday, May 22, 2001 10:42 AM
110
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C07 Page 111 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
111
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
0839_frame_C07 Page 112 Tuesday, May 22, 2001 10:42 AM
112
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C07 Page 113 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
113
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.
0839_frame_C07 Page 114 Tuesday, May 22, 2001 10:42 AM
114
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C07 Page 115 Tuesday, May 22, 2001 10:42 AM
Careers in Marine Mammal Medicine
115
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.
0839_frame_C07 Page 116 Tuesday, May 22, 2001 10:42 AM
0839_frame_C08 Page 117 Tuesday, May 22, 2001 10:43 AM
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
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
117
0839_frame_C08 Page 118 Tuesday, May 22, 2001 10:43 AM
118
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C08 Page 119 Tuesday, May 22, 2001 10:43 AM
The Electronic Whale
119
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
0839_frame_C08 Page 120 Tuesday, May 22, 2001 10:43 AM
120
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C08 Page 121 Tuesday, May 22, 2001 10:43 AM
The Electronic Whale
121
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
0839_frame_C08 Page 122 Tuesday, May 22, 2001 10:43 AM
122
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C08 Page 123 Tuesday, May 22, 2001 10:43 AM
123
The Electronic Whale
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
0839_frame_C08 Page 124 Tuesday, May 22, 2001 10:43 AM
124
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C08 Page 125 Tuesday, May 22, 2001 10:43 AM
125
The Electronic Whale
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.
0839_frame_C08 Page 126 Tuesday, May 22, 2001 10:43 AM
126
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 127 Tuesday, May 22, 2001 10:43 AM
II Anatomy and Physiology of Marine Mammals
0839_frame_C09 Page 128 Tuesday, May 22, 2001 10:43 AM
0839_frame_C09 Page 129 Tuesday, May 22, 2001 10:43 AM
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.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
129
0839_frame_C09 Page 130 Tuesday, May 22, 2001 10:43 AM
130
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 131 Tuesday, May 22, 2001 10:43 AM
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
0839_frame_C09 Page 132 Tuesday, May 22, 2001 10:43 AM
132
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 133 Tuesday, May 22, 2001 10:43 AM
133
Gross and Microscopic Anatomy
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
0839_frame_C09 Page 134 Tuesday, May 22, 2001 10:43 AM
134
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 135 Tuesday, May 22, 2001 10:43 AM
135
Gross and Microscopic Anatomy
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
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 137 Tuesday, May 22, 2001 10:43 AM
137
Gross and Microscopic Anatomy
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
0839_frame_C09 Page 138 Tuesday, May 22, 2001 10:43 AM
138
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C09 Page 139 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
139
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
0839_frame_C09 Page 140 Tuesday, May 22, 2001 10:43 AM
140
CRC Handbook of Marine Mammal Medicine
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).
0839_frame_C09 Page 141 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
141
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).
0839_frame_C09 Page 142 Tuesday, May 22, 2001 10:43 AM
142
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 143 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
143
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).
0839_frame_C09 Page 144 Tuesday, May 22, 2001 10:43 AM
144
CRC Handbook of Marine Mammal Medicine
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).
0839_frame_C09 Page 145 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
145
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
0839_frame_C09 Page 146 Tuesday, May 22, 2001 10:43 AM
146
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C09 Page 147 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
147
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
0839_frame_C09 Page 148 Tuesday, May 22, 2001 10:43 AM
148
CRC Handbook of Marine Mammal Medicine
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).
0839_frame_C09 Page 149 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
149
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).
0839_frame_C09 Page 150 Tuesday, May 22, 2001 10:43 AM
150
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C09 Page 151 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
151
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
0839_frame_C09 Page 152 Tuesday, May 22, 2001 10:43 AM
152
CRC Handbook of Marine Mammal Medicine
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,
0839_frame_C09 Page 153 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
153
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
0839_frame_C09 Page 154 Tuesday, May 22, 2001 10:43 AM
154
CRC Handbook of Marine Mammal Medicine
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,
0839_frame_C09 Page 155 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
155
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
0839_frame_C09 Page 156 Tuesday, May 22, 2001 10:43 AM
156
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C09 Page 157 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
157
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).
0839_frame_C09 Page 158 Tuesday, May 22, 2001 10:43 AM
158
CRC Handbook of Marine Mammal Medicine
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).
References Arey, L.B., 1941, Throttling veins in the livers of certain mammals, Anat. Rec., 81: 21–33. Baker, J.R., 1989, Pollution-associated uterine lesions in grey seals from the Liverpool Bay area of the Irish Sea, Vet. Rec., 125: 303. Baker, M.A., 1982, Brain cooling in endotherms in heat and exercise, Annu. Rev. Physiol., 44: 85–96. Baker, M.A., and Chapman, L.W., 1977, Rapid cooling in exercising dogs, Science, 195: 781–783. Bedford, J.M., 1977, Evolution of the scrotum: The epididymis as the prime mover? in Reproduction and Evolution, Proceedings of the Fourth Symposium on Comparative Biology and Reproduction, Calaby, J.H., and Tyndale-Biscoe, C.H. (Eds.), Australian Academy of Sciences, Canberra City, 171–182. Bigg, M.A., and Fisher, H.D., 1974, The reproductive cycle of the female harbour seal off southeastern Vancouver Island, in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 329–347. Blumberg, M.S., and Moltz, H., 1988, How the nose cools the brain during copulation in the male rat, Physiol. Behav., 43: 173–176. 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), U.S. Department of the Interior, National Biological Survey, available from National Technical Information Service, Springfield, VA. Boshier, D.P., 1974, Proceedings: Structural features in the lung of the Weddell seal, Leptonychotes weddellii, which facilitate deep diving: a light and electron microscope study, J. Anat., 118: 400. Boshier, D.P., 1979, Electron microscopic studies on the endometrium of the grey seal (Halichoerus grypus) during its preparation for nidation, J. Anat., 128: 721–735. Boshier, D.P., 1981, Structural changes in the corpus luteum and endometrium of seals before implantation, J. Reprod. Fert., Suppl., 29: 143–149. Boyd, R.B., 1975, A gross and microscopic study of the respiratory anatomy of the Antarctic Weddell seal, Leptonychotes weddellii, J. Morphol., 147: 309–336. Boyd, I.L., Lockyer, C., and Marsh, H.D., 1999, Reproduction in marine mammals, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 218–286. Brecht, M., Preilowski, B., and Merzenich, M.M., 1997, Functional architecture of the mysticial vibrissae, Behav. Brain Res., 84: 81–97. Britt, J.O., and Howard, E.B., 1983, Anatomic variants of marine mammals, in Pathobiology of Marine Mammal Disease, Vol. 1, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 8–46. Brown, W.R., Geraci, J.R., Hicks, D.B., St. Aubin, D.J., and Schroeder, J.P., 1983, Epidermal proliferation in the bottlenose dolphin (Tursiops truncatus), Can. J. Zool., 61: 1587–1590.
0839_frame_C09 Page 159 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
159
Bryden, M.M., 1971, Myology of the southern elephant seal Mirounga leonina (L.), Antarctic Pinnipedia, American Geophysical Union, Antarct. Res. Ser., 18: 109–140. Bryden, M.M., and Tedman, R., 1974, Proceedings: The mammary gland of the elephant seal, Mirounga leonina, J. Anat., 118, 399–400. Bryden, M.M., Griffiths, D.J., Kennaway, D.J., and Ledingham, J., 1986, The pineal gland is very large and active in newborn Antarctic seals, Experientia, 42: 564–566. Clarke, J.A., de Burgh Daly, M., and Elsner, R., 1986, The carotid body of the harbour seal (Phoca vitulina richardsi), Anat. Embryol., 174: 391–400. Costa, D.P., and Williams, T.M., 1999, Marine Mammal Energetics, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 176–217. Cowan, D.F., and Brownell, R.L., 1974, Gut-associated lymphoepithelial organ (“anal tonsil”) in the gray whale, in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 321–327. Cowan, D.F., and Smith, T.L., 1999, Morphology of the lymphoid organs of the bottlenose dolphin, Tursiops truncatus, J. Anat., 194: 505–517. Cowles, R.B., 1958, The evolutionary significance of the scrotum, Evolution, 12: 417–418. Cowles, R.B., 1965, Hyperthermia, aspermia, mutation rates, and evolution, Q. Rev. Biol., 40: 341–367. Cranford, T.W., 1999, The sperm whale’s nose: Sexual selection on a grand scale? Mar. Mammal Sci., 15: 1133–1157. Cranford, T.W., Amundin, M., and Norris, K.W., 1996, Functional morphological and homology in the odontocete nasal complex: Implications for sound generation, J. Morphol., 228: 223–285. DeSmet, W.M.A., 1977, The position of the testes in cetaceans, in Functional Anatomy of Marine Mammals, Vol. 3, Harrison, R.J. (Ed.), Academic Press, London, 361–386. Domning, D.P., 1977, Observations on the myology of Dugong dugon (Muller), Smithsonian Contrib. Zool., 226. Domning, D.P., 1978, The myology of the Amazonian manatee, Trichechus inunguis (Natterer) (Mammalia: Sirenia), Acta Amazonica, 8(2), Supl. 1: 1–81. Donaldson, B.J., 1977, The tongue of the bottlenose dolphin (Tursiops truncatus), in Functional Anatomy of Marine Mammals, Vol. 3, Harrison, R.J. (Ed.), Academic Press, London, 175–197. Drabek, C.M., 1975, Some anatomical aspects of the cardiovascular system of Antarctic seals and their possible functional significance in diving, J. Morphol., 145: 85–105. Elsner, R., 1969, Cardiovascular adaptations to diving, in The Biology of Marine Mammals, Anderson, H.I. (Ed.), Academic Press, London, 117–145. Elsner, R.W., 1999, Living in water: solutions to physiological problems, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 73–116. Elsner, R., and Gooden, B., 1983, Diving and asphyxia: a comparative study of animals and man, Physiological Society Monographs, Cambridge University Press, 40: 1–175. Elsner, R., Pirie, J., Kenney, D.D., and Schemmer, S., 1974, Functional circulatory anatomy of cetacean appendages, in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 143–159. English, A.W., 1976a, Functional anatomy of the hands of fur seals and sea lions, Am. J. Anat., 147: 1–18. English, A.W., 1976b, Limb movements and locomotor function in the California sea lion (Zalophus californianus), J. Zool., 178: 341–364. English, A.W., 1977, Structural correlates of forelimb function in fur seals and seal lions, J. Morphol., 151: 325–352. Erasmus, C.P., and Van Aswegen, G., 1997, The endocrine pancreas of the Cape fur seal, Arctocephalus pusillus (Schreber, 1776): An immunocytochemical study, Onderstepoort J. Vet. Res., 64: 239–242. Evans, H.E., 1993, Miller’s Anatomy of the Dog, 3rd ed., W.B. Saunders, Philadelphia, 550 pp. Fagone, D.M., Rommel, S.A., and Bolen, M.E., 2000, Sexual dimorphism in vestigial pelvic bones of the Florida manatee, Fla. Sci., 63: 177–181.
0839_frame_C09 Page 160 Tuesday, May 22, 2001 10:43 AM
160
CRC Handbook of Marine Mammal Medicine
Fawcett, D.W., 1942, The amedullary bones of the Florida manatee (Trichechus latirostris), Am. J. Anat., 71: 271–309. Flanigan, N.J., 1972, The central nervous system, in Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 215–246. Flower, W.H., 1885, An Introduction to the Osteology of the Mammalia, 3rd ed., Macmillan, London (reprinted by A. Asher & Co., Amsterdam, 1966), 258 pp. Fraser, F.C., 1952, Handbook of R.H. Burne’s Cetacean Dissections, Trustees of the British Museum, London, 210 pp. Gaskin, D.E., 1978, Form and function of the digestive tract and associated organs in cetacea, with a consideration of metabolic rates and specific energy budgets, Oceanogr. Mar. Biol. Annu. Rev., 16: 313–345. Green, R.F., 1972, Observations on the anatomy of some cetaceans and pinnipeds, in Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 247–297. Greenwood, A.G., Harrison, R.J., and Whitting, H.W., 1974, Functional and pathological aspects of the skin of marine mammals, in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 73–110. Griffiths, D.J., and Bryden, M.M., 1986, Adenohypophysis of the elephant seal (Mirounga leonina): Morphology and seasonal histological changes, Am. J. Anat., 176: 483–495. Harrison, J.F., Johnson, F.R., and Young, B.A., 1977, The small intestine of dolphins (Tursiops, Delphinus, Stenella), in Functional Anatomy of Marine Mammals, Vol. 3, Harrison, R.J. (Ed.), Academic Press, London, 297–331. Harrison, R.J., 1969a, Reproduction and reproductive organs, in The Biology of Marine Mammals, Andersen, H.T. (Ed.), Academic Press, New York, 253–348. Harrison, R.J., 1969b, Endocrine organs: Hypophysis, thyroid and adrenal, in The Biology of Marine Mammals, Andersen, H.T. (Ed.), Academic Press, New York, 349–390. Harrison, R.J., Brownell, R.L., Jr., and Boice, R.G., 1972, Reproduction and gonadal appearances in some odontocetes, in Functional Anatomy of Marine Mammals, Vol. 1, Harrison, R.J. (Ed.), Academic Press, London, 362–429. Hayakawa, D., Chen, H., Emura, S., Tamada, A., Yamahira, T., Terasawa, K., Isono, H., and Shoumura, S., 1998, The parathyroid glands of two species of dolphin—Risso’s dolphin, Grampus griseus, and bottlenose dolphin, Tursiops truncatus, Gen. Comp. Endocrinol., 110: 58–66. Herbert, D., 1987, The Topographic Anatomy of the Sea Otter Enhydra lutris, M.S. thesis, Johns Hopkins University, Baltimore, MD, 42 pp. Hilton, J.W., and Gaskin, D.E., 1978, Comparative volumes and vascular microanatomy of the intrahepatic venous system of the harbour porpoise, Phocoena phocoena (L.), Can. J. Zool., 56: 2292– 2298. Howell, A.B., 1930, Aquatic Mammals: Their Adaptations to Life in the Water, Charles C Thomas, Springfield, IL, 338 pp. Huber, E., 1934, Anatomical notes on Pinnipedia and Cetacea, in Marine Mammals, Packard, E.L., Kellogg, R., and Huber, E. (Eds.), Carnegie Institute of Washington, Contrib. Paleontol., 447: 105–136. King, J.E., 1983, Seals of the World, 2nd ed., Comstock, Ithaca, NY, 240 pp. Klima, M., Oeleschlager, H.A., and Wunsch, D., 1980, Morphology of the pectoral girdle in the Amazon dolphin Inia geoffrensis with special reference to the shoulder joint and movements of the flippers, Sonderdruck Z. Saugetierk., 45: 288–309. Kooyman, G.L., 1985, Physiology without constraint in diving mammals, Mar. Mammal Sci., 1: 166–178. Kooyman, G.L., Castellini, M.A., and Davis, R.W., 1981, Physiology of diving in marine mammals, Annu. Rev. Physiol., 43: 343–356. Leatherland, J.F., and Roland, K., 1976, Structure of the adenohypophysis in juvenile harp seal, Pagophilus groenlandicus, Cell Tissue Res., 173: 367–382. Leatherland, J.F., and Ronald, K., 1978, Structure of the adenohypophysis in parturient female and neonate harp seals, Pagophilus groenlandicus, Cell Tissue Res., 192: 341–357.
0839_frame_C09 Page 161 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
161
Leatherland, J.F., and Ronald, K., 1979, Structure of the testis in neonate and adult harp seals, Pagophilus groenlandicus, Cell Tissue Res., 201: 45–49. Ling, J.K., 1974, The integument of marine mammals, in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 1–44. Little, G.J., 1991, Thyroid morphology and function and its role in thermoregulation in the newborn southern elephant seal (Mirounga leonina) at Macquarie Island, J. Anat., 176: 55–69. Lowenstine, L.J., and Osborne, K.G., 1990, Practical marine mammal microanatomy for pathologists, in Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 287–290. Maluf, N.S.R., 1989, Renal anatomy of the manatee, Trichechus manatus, Linnaeus, Am. J. Anat., 184: 269–286. Maluf, N.S.R., and Gassmann, J.J., 1998, Kidneys of the killer whale and significance of reniculism, Anat. Rec., 250: 34–44. Marsh, H., Heinsohn, G.E., and Spain, A.V., 1977, The stomach and duodenal diverticulae of the dugong (Dugong dugon), in Functional Anatomy of Marine Mammals, Vol. 3, Harrison, R.J. (Ed.), Academic Press, London, 271–295. Mauck, B., Eysel, U., and Dehnhardt, G., 2000, Selective heating of vibrissal follicles in seals (Phoca vitulina) and dolphins (Sotalia fluviatilis guianensis), J. Exp. Biol., 203(14): 2125–2131. McFarland, W.L., Jacobs, M.S., and Morgane, P.J., 1979, Blood supply to the brain of the dolphin, Tursiops truncatus, with comparative observations on special aspects of the cerebrovascular supply of other vertebrates, Neurosci. Biobehav. Rev. (Suppl. 1), 3L: 1–93. McGrath, C.J., Feeney, D., Crimi, A.J., and Ruff, J., 1981, Upper airway of the California sea lion: An anesthetist’s perspective, Vet. Med. Small Anim. Clin., 76: 548–549. Mead, J.G., 1975, Anatomy of the external nasal passages and facial complex in the Delphinidae (Mammalia: Cetacea), Smithsonian Contrib. Zool., 207: 1–72. Miller, E.H., Stewart, A.R.J., and Stenson, G.B., 1998, Bacular and testicular growth, allometry, and variation in the harp seal (Pagophilus groenlandicus), J. Mammal., 79: 502–513. Mills, R.P., and Christmas, H.E., 1990, Applied comparative anatomy of the nasal turbinates, Clin. Otolaryngol., 15: 553–558. Morgane, P.J., and Jacobs, M.S., 1972, Comparative anatomy of the cetacean nervous system, in Functional Anatomy of Marine Mammals, Vol. 1, Harrison, F.J. (Ed.), Academic Press, London, 117– 244. Murie, J., 1872, On the form and structure of the manatee, Trans. Zool. Soc. London, 8: 127–202. Murie, J., 1874, Researches upon the anatomy of the Pinnipedia, Part 3, Descriptive anatomy of the sealion (Otaria jubata), Trans. Zool. Soc. London, 8: 501–582. Nakakuki, S., 1993a, The bronchial tree, lobular division and blood vessels of the harbor seal (Phoca vitulina) lung, Kaibogaku Zasshi, 68: 497–503. Nakakuki, S., 1993b, The bronchial tree and lobular division of the lung of the California sea lion (Zalophus californianus), J. Vet. Med. Sci., 55: 669–671. Nill, E.K., Rommel, S.A., Pabst, D.A., and McLellan, W.A., 2000, Does the thick skin of the Florida manatee (Trichechus manatus latirostris) provide ballast? Am. Zool., 39: 114A. Pabst, D.A., 1990, Axial muscles and connective tissues of the bottlenose dolphin, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 51–67. Pabst, D.A., 1993, Intramuscular morphology and tendon geometry of the epaxial swimming muscles of dolphins, J. Zool., 230: 159–176. Pabst, D.A., Rommel, S.A., McLellan, W.A., Williams, T.M., and Rowles, T.K., 1994, Temperature regulation of the dolphin testis, J. Morphol., 220: 397. Pabst, D.A., Rommel, S.A., McLellan, W.A., Williams, T.M., and Rowles, T.K., 1995, Thermoregulation of the intra-abdominal testes of the bottlenose dolphin (Tursiops truncatus) during exercise, J. Exp. Biol., 198: 221–226.
0839_frame_C09 Page 162 Tuesday, May 22, 2001 10:43 AM
162
CRC Handbook of Marine Mammal Medicine
Pabst, D.A., Rommel, S.A., and McLellan, W.A., 1999, The functional morphology of marine mammals, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 15–72. Ponganis, P.J., Kooyman, G.L., Sartoris, D., and Jobsis, P., 1992, Pinniped splenic volumes, Am. J. Physiol., 262: R322–325. Reep, R.L., Stoll, M.S., and Marshall, C.D., 1999, Do sirenians have a mammalian version of the lateral line? Postcranial tactile hairs in Florida manatees, Soc. Neurosci., Abstr., 25: 102. Reep, R.L., Johnson, J.I., Switzer, and R.C., Welker, W.I., 1989, Manatee cerebral cortex: Cytoarchitecture of the frontal region in Trichechus manatus latirostris, Brain Behav. Evol., 34: 365–386. Reidenberg, J.S., and Laitman, J.T., 1987, Position of the larynx in Odontoceti (toothed whales), Anat. Rec., 218: 98–106. Reynolds, J.E., III, and Rommel, S.A., 1996, Structure and function of the gastrointestinal tract of the Florida manatee, Trichechus manatus latirostris, Anat. Rec., 245: 539–558. Reynolds, J.E., III, O’Dell, D.K., and Rommel, S.A., 1999, Marine mammals of the world, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 1–14. Reynolds, J.E., III, Rommel, S.A., and Bolen, M.E., in press, Gross visceral morphology of marine mammals, in Encyclopedia of Marine Mammals, Perrin, W.F., Wursig, B., and Thewissen, H.G.M. (Eds.), Academic Press, New York. Rhodahl, K., and Moore, T., 1943, Vitamin A content and toxicity of bear and seal liver, Biochem. J., 37: 166–168. Riedman, M., 1990, The Pinnipeds: Seals, Sea Lions and Walruses, University of California Press, Berkeley, 439 pp. Romano, T.A., Felten, S.Y., Olschowka, J.A., and Felten, D.L., 1993, A microscopic investigation of the lymphoid organs of the beluga whale, Delphinapterus leucas, J. Morphol., 215: 261–287. Romer, A.S., and Parsons, T.S., 1977, The Vertebrate Body, 5th ed., W.B. Saunders, Philadelphia, 624 pp. Rommel, S.A., 1990, The osteology of the bottlenose dolphin, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, New York, 29–49. Rommel, S.A., and Reynolds, J.E., III, 2000, Diaphragm structure and function in the Florida manatee (Trichechus manatus latirostris), Anat. Rec., 259: 41–51. Rommel, S.A. and Reynolds, J.E., III, in press, Postcranial skeletal morphology of marine mammals, in Encyclopedia of Marine Mammals, Perrin, W.F., Wursig, B., and Thewissen, H.G.M. (Eds.), Academic Press, New York. Rommel, S.A., Pabst, D.A., McLellan, W.A., Mead, J.G., and Potter, C.W., 1992, Anatomical evidence for a countercurrent heat exchanger associated with dolphin testes, Anat. Rec., 232: 150–156. Rommel, S.A., Pabst, D.A., and McLellan, W.A., 1993a, Functional morphology of the vascular plexus associated with the cetacean uterus, Anat. Rec., 237: 538–546. Rommel, S.A., Pabst, D., McLellan, W., Early, G., and Matassa, K., 1993b, Cooled abdominal and epidural blood in dolphins and seals: Two previously undescribed thermoregulatory sites, Abstr., Tenth Biennial Conference on the Biology of Marine Mammals, Nov., Galveston, TX. Rommel, S.A., Pabst, D.A., McLellan, W.A., Williams, T.M., and Friedel, W.A., 1994, Temperature regulation of the testes of the bottlenose dolphin (Tursiops truncatus): Evidence from colonic temperatures, J. Comp. Physiol. B, 164: 130–134. Rommel, S.A., Early, G.A., Matassa, K.A., Pabst, D.A., and McLellan, W.A., 1995, Venous structures associated with thermoregulation of phocid seal reproductive organs, Anat. Rec., 243: 390–402. Rommel, S.A., Pabst, D.A., and McLellan, W.A., 1998, Reproductive thermoregulation in marine mammals, Am. Sci., 86: 440–448. Rommel, S., Pabst, D., and McLellan, W., in press, Skull morphology of marine mammals, in Encyclopedia of Marine Mammals, Perrin, W.F., Wursig, B., and Thewissen, H.G.M. (Eds.), Academic Press, New York.
0839_frame_C09 Page 163 Tuesday, May 22, 2001 10:43 AM
Gross and Microscopic Anatomy
163
St. Pierre, H., 1974, The topographical splanchnology and the superficial vascular system of the harp seal Pagophilus groenlandicus (Erxleben 1777), in Functional Anatomy of Marine Mammals, Vol. 2, Harrison, R.J. (Ed.), Academic Press, London, 161–195. Saltin, B., Gagge, A.P., and Stolwijk, J.A.J., 1968, Muscle temperature during submaximal exercise in man, J. Appl. Physiol., 65: 460–468. Schaller, O., 1992, Illustrated Veterinary Anatomical Nomenclature, Ferdinand Enke Verlag, Stuttgart, 560 pp. Schulte, H. von W., 1916, Anatomy of a fetus of Balaenoptera borealis, Mem. Am. Mus. Nat. Hist., 1(VI): 389–502 + plates XLIII–XLII. Schumacher, U., Zahler, S., Horny, H.P., Heidemann, G., Skirnisson, K., and Welsch, U., 1993, Histological investigations on the thyroid glands of marine mammals (Phoca vitulina, Phocoena phocoena) and the possible implications of marine pollution, J. Wildl. Dis., 29: 103–108. Schumacher, U., Klein, P., Plotz, J., and Welsch, U., 1995, Histological, histochemical, and ultrastructural investigations on the gastrointestinal system of Antarctic seals: Weddell seal (Leptonychotes weddellii) and crabeater seal (Lobodon carcinophagus), J. Morphol., 225: 229–249. Shadwick, R.E., and Gosline, J.M., 1994, Arterial mechanics in the fin whale suggest a unique hemodynamic design, Am. J. Physiol., 267(36): R805–R818. Simpson, J.G., and Gardner, M.B., 1972, Comparative microanatomy of selected marine mammals, in Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 298–418. Sinha, A.A., and Erickson, A.W., 1972, Ultrastructure of the corpus luteum of Antarctic seals during pregnancy, Z. Zellforsch. Mikrosk. Anat., 133: 13–20. Sinha, A.A., Erickson, A.W., and Seal, U.S., 1977a, Fine structure of Leydig cells in crabeater, leopard and Ross seals, J. Reprod. Fertil., 49: 51–54. Sinha, A.A., Erickson, A.W., and Seal, U.S., 1977b, Fine structure of seminiferous tubules in Antarctic seals, Cell Tissue Res., 178: 183–188. Slijper, E.J., 1962, Whales, Hutchinson, London, 511 pp. Stewardson, C.L., Hemsley, S., Meyer, M.A., Canfield, P.J., and MacDonald, J.H., 1999, Gross and microscopic visceral anatomy of the male Cape fur seal, Arctocephalus pusillus pusillus (Pinnipedia: Otariidae), with reference to organ size and growth, J. Anat., 195: 235–255. Strickler, T.L., 1978, Myology of the shoulder of Pontoporia blainvillei, including a review of the literature on shoulder morphology in the cetacea, Am. J. Anat., 152: 419–431. Strickler, T.L., 1980, The axial musculature of Pontoporia blainvillei, with comments on the organization of this system and its effect on fluke-stroke dynamics, Am. J. Anat., 157: 49–59. Tarasoff, F.J., 1972, Comparative aspects of the hindlimbs of the river otter, sea otter and seal, in Functional Anatomy of Marine Mammals, Vol. 1, Harrison, R.J. (Ed.), Academic Press, London, 333–359. Tarasoff, F.J., and Kooyman, G.L., 1973, Observations on the anatomy of the respiratory system of the river otter, sea otter, and harp seal, II, The trachea and bronchial tree, Can. J. Zool., 51: 171–177. Tedman, R.A., and Bryden, M.M., 1981, The mammary gland of the Weddell seal, Leptonychotes weddellii (Pinnipedia), I. Gross and microscopic anatomy, Anat. Rec., 199: 519–529. VanDemark, N.L., and Free, M.J., 1970, Temperature effects, in The Testis, Vol. 3, Johnson, A.D., Gomes, W.R., and VanDemark, N.L. (Eds.), Academic Press, New York, 233–312. Vardy, P.H., and Bryden, M.M., 1981, The kidney of Leptonychotes weddellii (Pinnipedia: Phocidae) with some observations on the kidneys of two other southern phocid seals, J. Morphol., 167: 13–34. Welsch, U., Schwertfirm, S., Skirnisson, K., and Schumacher, U., 1997, Histological, histochemical, and fine structural observations on the lymph node of the common seal (Phoca vitulina) and the grey seal (Halichoerus grypus), Anat. Rec., 247: 225–242. Wessels, J.C., and Chase, C.C., 1998, Light and electron microscopical observations on the terminal airways and alveoli of the lung of the South African (Cape) fur seal Arctocephalus pusillus, Onderstepoort J. Vet. Res., 65: 253–262.
0839_frame_C09 Page 164 Tuesday, May 22, 2001 10:43 AM
164
CRC Handbook of Marine Mammal Medicine
Zapol, W.M., Liggins, G.C., Schneider, R.C., Qvist, J., Snider, M.T., Creasy, R.K., and Hochachka, P.W., 1979, Regional blood flow during simulated diving in the conscious Weddell seal, J. Appl. Physiol., 47: 986–973.
0839_frame_C10 Page 165 Tuesday, May 22, 2001 11:08 AM
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
165
0839_frame_C10 Page 166 Tuesday, May 22, 2001 11:08 AM
166
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C10 Page 167 Tuesday, May 22, 2001 11:08 AM
Endocrinology
167
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
0839_frame_C10 Page 168 Tuesday, May 22, 2001 11:08 AM
168
CRC Handbook of Marine Mammal Medicine
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 <93 23–93 14–23 23–93 23 29–68,904 1275–4172 239–927 10–110 12–60 26 0–9000 0–160
Reference Stokkan et al., 1995 Stokkan et al., 1995 Stokkan et al., 1995 Stokkan et al., 1995 Bryden et al., 1986 Bryden et al., 1986 Barrell and Montgomery, 1989 Barrell and Montgomery, 1989 Bryden, 1994; Bryden et al., 1994 Bryden, 1994; Bryden et al., 1994 Bryden, 1994; Bryden et al., 1994 Bryden et al., 1994 Bryden et al., 1994 Bryden et al., 1994 Bryden et al., 1994 Little and Bryden, 1990 Bryden, 1994 Bryden, 1994 Griffiths et al., 1979 Griffiths and Bryden, 1981; Griffiths, 1985 Bryden et al., 1986 Stokkan et al., 1995 Stokkan et al., 1995
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. pg/ml × 4.314 = pmol/l.
less than 1000 pg/ml over the ensuing month (Bryden et al., 1986). Harp (Pagophilus groenlandicus), hooded (Cystophora cristata), and gray (Halichoerus grypus) seals show a similar pattern, with peak values of roughly 6000 to 9000 pg/ml. Since all these species give birth under relatively harsh environmental conditions, at least in the areas where they were studied, it has been suggested that, as in some other mammals (Heldmaier et al., 1981; Puig-Domingo et al., 1988), the hormone acts to enhance the production of T3 to stimulate nonshivering thermogenesis (NST) (Little and Bryden, 1990; Bryden, 1994). However, since the commonly recognized mechanism for NST involves brown adipose tissue, which has yet to be demonstrated in species such as Weddell and hooded seals, Stokkan et al. (1995) have suggested an alternative, but as yet untested, explanation to account for the extraordinarily high circulating levels of melatonin in these animals. The potent antioxidant properties of the hormone might protect the fetus from the detrimental effects of hypoxia experienced in utero during diving. Melatonin levels in plasma vary seasonally in southern elephant seals, with high concentrations inhibiting gonadotropic hormones in winter; lower activity in summer allows gonadal recrudescence to occur (Griffiths et al., 1979; Griffiths and Bryden, 1981; Griffiths, 1985). Circadian rhythms typical of other mammals are evident in several species (Griffiths et al., 1979; Bryden et al., 1994; Stokkan et al., 1995), and are abolished as expected under conditions of continuous daylight in southern elephant seals and Weddell seals (Griffiths et al., 1979; Barrell and Montgomery, 1989).
0839_frame_C10 Page 169 Tuesday, May 22, 2001 11:08 AM
Endocrinology
169
Hypothalamus–Pituitary The endocrine connections linking higher centers of the central nervous system through to the pituitary gland have received little detailed study in marine mammals, and are presumed to function in a fashion similar to those in most other mammals. The organization of the pituitary itself is unremarkable, with distinguishable regions comparable to the pars distalis (adenohypophysis, anterior pituitary), pars nervosa (neurohypophysis, posterior pituitary), and pars intermedia (Harrison, 1969). The gland appears relatively immature in newborn elephant and harp seals (Leatherland, 1976; Leatherland and Ronald, 1978; Bryden, 1994), but well developed in the more precocious harbor seal (Amoroso et al., 1965). Immunohistochemical techniques have been used to identify the primary cell types typical of those for other mammals (Leatherland and Ronald, 1983; Bryden, 1994). Material derived from commercial whaling operations afforded the opportunity to isolate and characterize the adenohypophyseal hormones—adrenocorticotropic hormone (ACTH), thyroid-stimulating hormone (TSH), growth hormone (GH, somatotropin), lutenizing hormone (LH), and prolactin (PRL)—in a variety of mysticetes and also sperm whales (Physeter macrocephalus) (Kawauchi et al., 1978; Kawauchi and Tubokawa, 1979; Kawauchi, 1980). Considerable homology exists between the cetacean forms and those in other mammals. In fact, the amino acid sequence for ACTH from fin whales (Balaenoptera physalus) was found to be identical to that of humans (Kawauchi et al., 1978). Measurement of circulating levels of anterior pituitary hormones has seldom been reported. Most of the information available is for the gonadotropic hormones, considered elsewhere in this book (see Chapter 11, Reproduction). In view of the demonstrated homology between human and mysticete ACTH, analysis using conventional RIA systems would be expected to yield satisfactory results. Nevertheless, there are no published values for this hormone for any marine mammal. Commercially available reagents for measuring human TSH appear to be ineffective in detecting the hormone in belugas (Delphinapterus leucas) and bottlenose dolphins (Tursiops truncatus) (St. Aubin and Geraci, unpubl. data). John et al. (1980) used an RIA specific for ovine GH to monitor relative changes in GH-like protein in young harp seals. Since purified seal GH was unavailable to validate and calibrate the assay, no actual concentrations could be reported. The development and application of methodologies specific for the hormones in marine mammals would lead to greater insights into the regulation of these important endocrine pathways. Neurohypophyseal hormones principally include oxytocin (OT) and vasopressin. The latter will be reviewed in a subsequent section for its role in water balance. OT enhances smooth muscle contraction and plays a key role in parturition and milk flow during nursing. Injections of 15 to 50 IU of commercially available, synthetic hormone have been used to facilitate the collection of samples for studies on the energetic value and proximate content of milk from pinnipeds (Iverson et al., 1993; Lydersen et al., 1995; 1997). The effectiveness of the homologue suggests at least crude similarities in the role played by this hormone in pinnipeds and other mammals.
Thyroid Gland In contrast to the patchy information on most endocrine systems in marine mammals, reports on TH abound for these species. The integral role of TH in regulating metabolism has perhaps fueled a more extensive inquiry, given the long-standing, but more recently tempered, views suggesting extraordinarily elevated metabolism in marine mammals (Lavigne et al., 1986). THs are among the more broadly conserved and uniformly evident hormones in vertebrates. Thus, assays utilizing RIA or enzyme-linked immunosorbent assay (ELISA) developed for humans
0839_frame_C10 Page 170 Tuesday, May 22, 2001 11:08 AM
170
CRC Handbook of Marine Mammal Medicine
and other species have been extensively applied in the measurement of total thyroxine (tT4) and tT3, with apparently satisfactory results (Greenwood and Barlow, 1979). T4 is typically the form detected in highest concentration, but its ability to elicit cellular responses is generally less than for its principal metabolic derivative, T3. Activation of T4 to T3 is mediated by a suite of deiodinating enzymes occurring in, and sometimes specific to, various peripheral tissues. The enzymes vary in their kinetic properties and sensitivities to inhibitors such as propylthiouracil (PTU). The thyroid gland is the site of hormone synthesis and storage of both T4 and T3. It is the only endocrine gland that establishes a significant reserve of hormone that can be later metered into circulation to meet metabolic needs. The hormones are stored as part of a colloid matrix composed of thyroglobulin, which carries coupled iodinated tyrosine residues. The colloid is deposited extracellularly and contained within a follicle lined by thyrocytes, which can pinocytose the matrix and release the hormones as needed. Histological examination of marine mammal thyroid has revealed no important differences from this typical arrangement, but has shown marked variation in the apparent levels of activity of thyrocytes at various times during the development and life history of phocids (Harrison et al., 1962; Amoroso et al., 1965; Little, 1991) and cetaceans (Harrison, 1969; St. Aubin and Geraci, 1989). Early investigators were impressed by the size of the cetacean thyroid, particularly in its proportion to the weight of the animal. Belugas have three times more thyroid per unit body weight than a thoroughbred horse, and bottlenose dolphins have nearly twice as much as do humans (400 vs. 250 mg/kg) (Ridgway and Patton, 1971). This observation correlated well with assumptions that cetacean metabolic rate exceeded that predicted by Kleiber’s formula, presuming that the size of the gland reflected the amount of hormone released into circulation (Harrison and Young, 1970). Extensive measurements of both THs and reevaluation of assumptions about metabolic rate (see Chapter 36, Nutrition) have failed to support such a correlation. Nevertheless, the large reserve of hormone present in the beluga thyroid can sustain the marked elevation in circulating levels of TH that occurs during a brief period of thyroid hyperactivity in the summer period of estuarine occupation (St. Aubin and Geraci, 1989; St. Aubin et al., in press). An important consideration in the evaluation of thyroid status is the degree to which the hormones are bound by circulating proteins, principally thyroid binding globulin (TBG). Binding also occurs, but with lower affinity, to pre-albumin and albumin; efforts to demonstrate a pre-albumin binding protein in belugas and bottlenose dolphins have proved unsuccessful using methodologies established for other mammals (St. Aubin and Geraci, unpubl. data). It is presumed that the free, or unbound, hormone is responsible for regulating cellular processes, and that protein binding in circulation serves to deliver the hormone, maintain an available pool, and modulate the activity of metabolically potent substances such as TH (Ekins, 1986). The impact of TH can thus be regulated at a variety of levels, including rate of secretion from the thyroid gland, plasma binding capacity, rate of conversion to T3, and density of cellular receptors for the hormone. Although analysis of circulating levels represents the most readily obtained measure of thyroid status, it may yield misleading or confusing results if other elements are not taken into consideration. Blood concentrations of TH, both total and free hormone, have been reported for a number of marine mammal species (Tables 2 to 4). Variation in methodology, particularly with respect to the earlier literature, makes direct comparisons among species difficult. Nevertheless, certain patterns emerge. Levels of total T4 in cetaceans tend to be higher than for most other species, although concentrations in dolphins are roughly comparable to those in humans. Pinnipeds and polar bears (Ursus maritimus) show concentrations similar to those in most terrestrial mammals, but surprisingly are lower than in manatees (Trichechus manatus) which seems incongruous in light of the notoriously low metabolic rate of the latter (Gallivan and Best, 1980)
0839_frame_C10 Page 171 Tuesday, May 22, 2001 11:08 AM
171
Endocrinology
TABLE 2 Reported Circulating Concentrations (µg/dl) of Thyroxine in Marine Mammals Species
Specimens
Thyroxine (µg/dl)
Reference
Cetaceans Balaenoptera physalus (fin whale) Delphinapterus leucas (beluga) Globicephala macrorhyncus (short-finned pilot whale) Inia geoffrensis (Amazon River dolphin) Lagenorhynchus obliquidens (Pacific whitesided dolphin) Orcinus orca (killer whale) Phocoena phocoena (harbor porpoise) Tursiops truncatus (bottlenose dolphin)
Not specified
5.4
Kjeld and Olafsson, 1987
Various ages, both sexes Two males, age unspecified
8.0–19.2 4.3
St. Aubin and Geraci, 1988; 1989; 1992; St. Aubin et al., in press Ridgway et al., 1970
Various ages, both sexes
1.5
Ridgway et al., 1970
Five females, age unspecified
2.6–3.7
Ridgway et al., 1970
Two males, age unspecified Various ages, both sexes Various ages, both sexes
6.1
Ridgway et al., 1970
11.2
Koopman et al., 1995
7.4–13.6
Ridgway et al., 1970; Greenwood and Barlow, 1979; Orlov et al., 1988; St. Aubin et al., 1996
Pinnipeds Callorhinus ursinus (northern fur seal) Cystophora cristata (hooded seal) Halichoerus grypus (gray seal)
Leptonychotes weddellii (Weddell seal) Mirounga angustirostris (northern elephant seal)
Various ages, both sexes Neonates Neonates
2.8
St. Aubin, unpubl. data
4–9
Stokkan et al., 1995
3–10
Stokkan et al., 1995; Woldstad et al., 1999
Pups (4 d) Pups (10–14 d)
1–6 1.5–7.1
Juveniles (>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)
0839_frame_C10 Page 172 Tuesday, May 22, 2001 11:08 AM
172
CRC Handbook of Marine Mammal Medicine
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 (<10 d)
1.4–20
Pups (2–3 wk) Lactating
4.6–6.5 0.4–6.1
Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; Stokkan et al., 1995 Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; Stokkan et al., 1995 Engelhardt and Ferguson, 1980 Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980 John et al., 1987
Adults, molting and 1–4 wk postmolt Adults
Phoca largha (spotted seal)
Thyroxine (µg/dl)
5.9, 4.6, 5.9 0.6–3
Juveniles Adults Juveniles, molting Adults, molting Neonates Pups (2–4 wk) Juveniles
0.2–3 1.2–3 0.5–4 0.5–5 8.2 4.1–4.8 0.6–4
Adults
0.5–3
Lactating Juveniles, molting
1.9–3.1 1.8–5
Adults, molting
0.5–1
Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; John et al., 1987 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Haulena et al., 1998 Haulena et al., 1998 Riviere et al., 1977; Ashwell-Erickson et al., 1986 Ronald and Thomson, 1981; Ashwell-Erickson et al., 1986; Brouwer et al., 1989; Renouf and Brotea, 1991; Renouf and Noseworthy, 1991 Haulena et al., 1998 Riviere et al., 1977; Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986
Sirenians Trichechus manatus (West Indian and Florida manatees)
Captive Free-ranging
1.9–4.5 4.5–8.3
Ortiz et al., 2000 Ortiz et al., 2000
Sea Otter Enhydra lutris
Pups Juveniles Adults
3.75 2.7 2.45
Williams et al., 1992 Williams et al., 1992 Williams et al., 1992
Polar Bear Ursus maritimus
Adults
0.6–5.2
Leatherland and Ronald, 1981; Cattet, 2000
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. µg/dl × 12.87 = nmol/l.
0839_frame_C10 Page 173 Tuesday, May 22, 2001 11:08 AM
173
Endocrinology
TABLE 3 Reported Concentrations of Triiodothyronine (ng/dl) in Marine Mammals
Species
Triiodothyronine (ng/dl)
Specimens
Reference
Cetaceans Delphinapterus leucas (beluga) Tursiops truncatus (bottlenose dolphin)
Various ages, both sexes Adults, both sexes
59–177 83–165
St. Aubin and Geraci, 1988; 1989; St. Aubin et al., in press Greenwood and Barlow, 1979; Orlov et al., 1988; St. Aubin et al., 1996
Pinnipeds Callorhinus ursinus (northern fur seal) Cystophora cristata (hooded seal) Halichoerus grypus (gray seal)
Leptonychotes weddellii (Weddell seal) Mirounga leonina (southern elephant seal) Pagophilus groenlandicus (harp seal)
Various ages, both sexes Pups (1 d)
63 100–225
Neonates (1 d) Pups (4 d)
60–225 60–250
Pups (1–2 wk)
47–280
Post-weaned pups and juveniles Juveniles, molting Adults
44–130
Adults, molting Various ages, both sexes Neonates (6 h) Pups (14–20 d)
42 100 195 85
Little, 1991 Little, 1991
152 36–111
83–137
Pups (1–5 d)
60–360
Pups (7–10 d) Pups (2 wk)
130–226 60–170
Pups (3 wk) Adults
207–330 45–220
Adults, molting
Stokkan et al., 1995 Stokkan et al., 1995 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; Woldstad et al., 1999 Boily, 1996 Engelhardt and Ferguson, 1980; Boily, 1996; Hall et al., 1998 Boily, 1996 Schumacher et al., 1992
Neonates (9 h)
Lactating
St. Aubin, unpubl. data
45–120 227
Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; John et al., 1987; Stokkan et al., 1995 Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; Stokkan et al., 1995 Leatherland and Ronald, 1979 Engelhardt and Ferguson, 1980; Stokkan et al., 1995 Engelhardt and Ferguson, 1980 Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980; John et al., 1987 Leatherland and Ronald, 1979; Engelhardt and Ferguson, 1980 John et al., 1987 (Continued)
0839_frame_C10 Page 174 Tuesday, May 22, 2001 11:08 AM
174
CRC Handbook of Marine Mammal Medicine
TABLE 3 Reported Concentrations of Triiodothyronine (ng/dl) in Marine Mammals (continued)
Species Phoca largha (spotted seal)
Phoca vitulina (harbor seal)
Specimens Juveniles Adults Juveniles, molting Adults, molting Neonates (1 d) Pups (3–7 d) Pups (10–14 d) Pups (28 d) Juveniles Adults
Pregnant Postpartum Juveniles, molting Adults, molting
Triiodothyronine (ng/dl) 10–160 10–30 20–130 10–80 130 210 163 98 39–42 10–78
29 124 30–104 20–47
Reference Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Haulena et al., 1998 Haulena et al., 1998 Haulena et al., 1998 Haulena et al., 1998 Ashwell-Erickson et al., 1986; Renouf and Brotea, 1991 Ashwell-Erickson et al., 1986; Renouf and Brotea, 1991; Renouf and Noseworthy, 1991; Haulena et al., 1998 Brouwer et al., 1989 Ronald and Thomson, 1981 Ashwell-Erickson et al., 1986; Renouf and Brotea, 1991 Ashwell-Erickson et al., 1986; Renouf and Brotea, 1991
Sirenians Trichechus manatus (West Indian and Florida manatees)
140–160
Ortiz et al., 2000
Polar Bear Ursus maritimus
16–150
Leatherland and Ronald, 1983; Cattet, 2000
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. ng/dl × 0.01536 = nmol/l.
(see Chapter 36, Nutrition; Chapter 43, Manatees). Levels of T4 in sea otters (Enhydra lutris) reveal little of the very active metabolism of these animals (Williams et al., 1992). THs, particularly T4, appear to be cleared from circulation very rapidly in bottlenose dolphins, 15 times faster on average than in humans; a single study in a Pacific white-sided dolphin (Lagenorhynchus obliquidens) yielded results comparable to those in humans (Sterling et al., 1975). The authors suggested that low protein binding in bottlenose dolphins might account for the rapid loss from the circulation. However, free THs, expressed as a percentage of the total hormone concentration, are in fact lower in bottlenose dolphins than in humans and other mammals (St. Aubin et al., 1996). Removal of T4 through other metabolic pathways could play an important role in the dynamics of circulating TH in these species. A significant, but poorly understood, difference in THs in some marine mammals is their relatively high circulating levels of reverse T3 (rT3). The product of inner ring deiodination of T4 (outer ring deiodination of T4 yields T3), rT3 is considered to be an inactive metabolite found in blood in concentrations that are generally one third to one half those of T3. In cetaceans and harbor seals, however, rT3 concentrations are equivalent to or up to three times greater
0839_frame_C10 Page 175 Tuesday, May 22, 2001 11:08 AM
175
Endocrinology
TABLE 4 Reported Concentrations of f T4 (ng/dl), f T3 (pg/ml), and rT3 (ng/ml) in Marine Mammals Species
Specimens
Concentration
Reference
f T4 Delphinapterus leucas (beluga) Globicephala macrorhynchus (short-finned pilot whale) Lagenorhyncus obliquidens (Pacific white-sided dolphin) Orcinus orca (killer whale) Tursiops truncatus (bottlenose dolphin) Callorhinus ursinus (northern fur seal) Halichoerus grypus (gray seal)
Phoca largha (spotted seal)
Phoca vitulina (harbor seal)
Trichechus manatus (West Indian and Florida manatees)
Various ages, both sexes Males, age unspecified
1.52
St. Aubin et al., in press
3.99
Ridgway et al., 1970
Both sexes, age unspecified
1.7–2.3
Ridgway et al., 1970
Male, age unspecified Various ages, both sexes Various ages, both sexes Pups (<10 d) Pups (>2 wk) Juveniles Adults Juveniles, molting Adults, molting Juveniles
2.78
Ridgway et al., 1970
1.36–3.58 0.25
Ridgway et al., 1970; St. Aubin et al., 1996 St. Aubin, unpubl. data
2–2.57 2–2.23 1.07 1.1–1.44 2.35 1.22 1–4
Hall et al., 1998; Woldstad et al., 1999 Hall et al., 1998; Woldstad et al., 1999 Boily, 1996 Boily, 1996; Hall et al., 1998 Boily, 1996 Boily, 1996 Ashwell-Erickson et al., 1986
Adults Adults, molting Neonates Pups (1–4 wk) Juvenile
1.5–3 1.2–5.5 2.18 1.0 1.5–2
Adult
1.50–1.93
Pregnant Postpartum Free-ranging Captive
1.0 1.4 1.33–1.59 0.5–1.13
Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Haulena et al., 1998 Haulena et al., 1998 Ashwell-Erickson et al., 1986; Renouf and Brotea, 1991 Renouf and Brotea, 1991; Renouf and Noseworthy, 1991 Brouwer et al., 1989 Haulena et al., 1998 Ortiz et al., 2000 Ortiz et al., 2000
f T3 Delphinapterus leucas (beluga) Tursiops truncatus (bottlenose dolphin) Halichoerus grypus (gray seal)
Various ages, both sexes Adults, both sexes
1.68
St. Aubin et al., in press
1.29
St. Aubin et al., 1996
Preweaned pups Postweaned pups Adult female
0.87 0.84 0.90
Hall et al., 1998 Hall et al., 1998 Hall et al., 1998 (Continued)
0839_frame_C10 Page 176 Tuesday, May 22, 2001 11:08 AM
176
CRC Handbook of Marine Mammal Medicine
TABLE 4 Reported Concentrations of fT4 (ng/dl), fT3 (pg/ml), and rT3 (ng/ml) in Marine Mammals (continued) Species Phoca vitulina (harbor seal)
Specimens Neonates Pups (5–15 d) Pups (19–26 d) Postpartum
Concentration 1.79 1.63–2.28 1.11–1.63 0.2–0.39
Reference Haulena et al., 1998 Haulena et al., 1998 Haulena et al., 1998 Haulena et al., 1998
rT3 Delphinapterus leucas (beluga) Tursiops truncatus (bottlenose dolphin) Callorhinus ursinus (northern fur seal) Phoca vitulina (harbor seal)
Various ages, both sexes Adults, both sexes Various ages, both sexes Neonates Pups (5–25 d) Postpartum
4.0
St. Aubin et al., in press
1.81
St. Aubin et al., 1996
1.6
St. Aubin, unpubl. data
9.0 0.65–1.95 0.65–1.95
Haulena et al., 1998 Haulena et al., 1998 Haulena et al., 1998
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. ng/dl × 12.87 = pmol/l for f T4; pg/ml × 1.536 = pmol/l for f T3; ng/ml × 1.536 = nmol/l for rT3.
than T3 (St. Aubin et al. 1996; in press; Haulena et al., 1998). During the summer period of estuarine occupation in belugas, rT3 levels can reach 4.4 ng/ml, the highest reported for any adult mammal. The benefits of inactivating such a large proportion of T4 are unclear, but at the very least represent another option for managing the effects of circulating T4. Interpretation of TH levels in marine mammals, and particularly in pinnipeds, must take into account dynamic changes that occur in association with significant life-history events. Neonatal phocid seals typically show levels that are elevated above reference ranges for adults (Engelhardt and Ferguson, 1980; Stokkan et al., 1995; Haulena et al., 1998; Woldstad and Jenssen, 1999), a pattern similar to that in humans and domestic mammals. The elevations are consistent with histological evidence of hyperactivity in thyroid follicular cells, at least in harbor seals (Harrison et al., 1962; Amoroso et al., 1965) and elephant seals (Little, 1991); the harp seal shows no such correlation (Leatherland, 1976; Leatherland and Ronald, 1979). For many phocids, metabolically derived heat may be critical for survival until an insulative blubber layer is established, and the calorigenic effects of TH could readily explain the need for elevated levels at this time (Stokkan et al., 1995; Haulena et al., 1998). The levels decline during the first few weeks of life in most species, although a trend for T3 may not always be apparent. In southern elephant seals, high concentrations of melatonin are postulated to enhance the conversion of T4 to T3, thereby providing an additional stimulus to metabolism. Circulating levels of TH in marine mammals are also subject to considerable fluctuations throughout the year, particularly in phocid seals. Chief among the events associated with altered TH status is the molt (Riviere et al., 1977; Ashwell-Erickson et al., 1986; John et al., 1987; Boily, 1996). Among their many metabolic effects, THs are known to stimulate hair growth in terrestrial mammals, and presumably have the same effect in phocids. Elevated levels in TH are typically observed during the less obvious phase of follicular stimulation prior to the time of most extensive shedding, which may account for Renouf and Brotea’s (1991) inability to establish direct correlations with overt signs of molting. The profound changes in TH that have been documented in most phocids are indicative of the broad metabolic adjustments that occur
0839_frame_C10 Page 177 Tuesday, May 22, 2001 11:08 AM
Endocrinology
177
during the molt (Ashwell-Erickson et al., 1986; Boily, 1996), and often correlate with other diagnostic signs of less than optimal health (Riviere, 1978). Seasonal variation in TH in harbor seals was associated with changes in appetite, fat accumulation, and metabolism (Renouf and Noseworthy, 1991). The only cetacean yet to be shown to have a comparable cycle in TH is the beluga (St. Aubin and Geraci, 1989), in which a concurrent stimulation of epidermal cell growth takes on virtually all manifestations of a molt (St. Aubin et al., 1990). Captive belugas, held under relatively constant environmental conditions, show neither the marked seasonal variations in TH nor the intense variation in epidermal cell turnover, although episodic sloughing does occur. Altered TH status has been associated with capture and handling in belugas (St. Aubin and Geraci, 1988; 1992), initially as part of stress-mediated changes in hormone secretion and metabolism. In other mammals, cortisol inhibits TSH secretion and also the monoiodinase responsible for producing much of the T3 in circulation, and these pathways appear to be similarly affected in belugas. Acclimation to captivity results in somewhat lower TH levels than observed even in the less active spring and fall seasons in the wild (St. Aubin et al., in press; St. Aubin and Ridgway, unpubl. data). Free-ranging female bottlenose dolphins have higher levels of tT4, f T4, and fT3 than their counterparts in captivity, possibly reflecting differences in their reproductive status (St. Aubin et al., 1996). No diurnal cycle was noted in T4 or T3 in neonatal harp and gray seals (Stokkan et al., 1995). However, belugas sampled at various times of day over a 4-year period showed a nadir in T4 concentration at 2200 hours, and a peak in T3 at 1400 hours (St. Aubin and Ridgway, unpubl. data). Thyroid stimulation tests have been performed in belugas (St. Aubin, 1987; St. Aubin and Geraci, 1992). Marked differences in response to 10 IU of bovine TSH were observed as a function of the time after capture the hormone was administered, with apparently diminished sensitivity over time. In three individuals, three doses given over a 58-hour period had no apparent adverse effect, and resulted in substantial elevation of both T4 and T3; no attempt was made to establish the optimum dosage through the use of graded doses of TSH. Pathological changes in thyroid have been described, though principally in association with other clinical problems (Greenwood and Barlow, 1979). There is some evidence that environmental contaminants acting as endocrine disrupters can upset TH balance (Brouwer et al., 1989; Hall et al., 1998) and produce histologically detectable abnormalities (Schumacher et al., 1993). Belugas from the St. Lawrence estuary in Canada, which are known to accumulate substantial burdens of organochlorine contaminants, among others (see Chapter 22, Toxicology), also show evidence of thyroid pathology (De Guise et al., 1994), although the association with contaminants is likely to remain circumstantial in the absence of experimental data in these species.
Adrenal Gland The adrenal gland of marine mammals conforms to the same general architecture noted in terrestrial mammals, with a catecholamine-secreting medulla surrounded by a steroid-producing cortex. A prominent difference is the pseudolobulation of the cortex produced by septae of fibrous tissue arising from the capsule; these lobules are most extensively developed in cetaceans. The cortex is particularly well developed in fetal harbor seals, as a possible adaptation to precocious behavior and physiological accommodation in the neonate (Amoroso et al., 1965; Sucheston and Cannon, 1980). Within the cortex, the outermost layer, or zona glomerulosa, is most expansive, suggesting that the need to produce aldosterone for electrolyte homeostasis is critical at that time.
0839_frame_C10 Page 178 Tuesday, May 22, 2001 11:08 AM
178
CRC Handbook of Marine Mammal Medicine
Few studies have examined catecholamine function and physiology in marine mammals. Attention has focused principally on the role of epinephrine and norepinephrine in the dive response (Hance et al., 1982; Hochachka et al., 1995; Lohman et al., 1998). Catecholamineinduced splenic contraction can contribute substantially to the circulating pool of erythrocytes, extending the aerobic dive limits for these animals. The hormones increase during dives of more than a few minutes in Weddell seals, and rapidly return to resting levels following the dive. Efforts to extract and identify steroids from adrenal tissues have yielded conflicting, sometimes puzzling, results regarding the types of steroids utilized by cetaceans and pinnipeds (DeRoos and Bern, 1961; Borruel et al., 1974; Carballeira et al., 1987). Nevertheless, virtually all studies on circulating corticosteroids have established the prominence of cortisol over corticosterone as the principal glucocorticoid (Sangalang and Freeman, 1976; Thomson and Geraci, 1986; Ortiz and Worthy, 2000), and the presence of aldosterone as the mineralocorticoid hormone. Establishing baseline values for constituents known to be influenced by stressors such as chase, capture, and restraint is challenging for the researcher, and clinicians are tasked with interpreting whether the efforts to obtain a blood sample for diagnostic purposes have produced misleading information. Captive bottlenose dolphins conditioned to allow unrestrained blood collection and those calmly approached and sampled within minutes have yielded specimens as close to baseline as can reasonably be expected (Thomson and Geraci, 1986; St. Aubin et al., 1996). Pinnipeds resting on ice floes or shorelines, or held in exhibits or research facilities, can sometimes be captured and sampled before circulating hormones change appreciably. Studies on the dynamics of corticosteroid release following stimulation by exogenous ACTH suggest that cortisol levels are elevated by 30 min (St. Aubin and Geraci, 1986; 1990; Thomson and Geraci, 1986). Even taking into account the possible artifact of capture-related elevations, the Weddell seal is distinguished among marine mammals, and indeed among most vertebrate species, by its extraordinarily high circulating concentrations of cortisol (Table 5) (Liggins et al., 1979; Barrell and Montgomery, 1989; Bartsh et al., 1992). No clear explanation has emerged to account for this conspicuous difference; it is not an adaptation necessary to diving, given the much lower levels in other pinnipeds, including the deep-diving elephant seal. Cortisol secretion tends to show a circadian cycle in mammals, with increasing levels during the morning hours in diurnal species, but there is little information on this point for marine mammals. Harbor seals show the highest concentrations at night, and the lowest in the early afternoon (Gardiner and Hall, 1997). No periodicity was evident in samples collected from Weddell seals exposed to continuous daylight; however, the study used pooled data from different seals sampled at different times and did not strictly follow changes in individual animals (Barrell and Montgomery, 1989). Belugas showed lower levels of cortisol between noon and midnight than during the rest of the day (St. Aubin and Ridgway, unpubl. data); a similar pattern was evident in a captive killer whale (Orcinus orca) (Suzuki et al., 1998). Other factors contributing to alterations in circulating cortisol levels include reproduction and molt. High levels of cortisol have been noted in molting seals, generally in an inverse relationship with thyroid hormones (Riviere et al., 1977; Ashwell-Erickson et al., 1986), although Boily (1996) found lower levels in molting gray seals. Cortisol concentrations are elevated in neonatal harp seals, decline within 3 days, and then return to the higher range by 3 weeks, at the time of lanugo shedding (Engelhardt and Ferguson, 1980). Cortisol is known to promote hair loss in terrestrial mammals. During late pregnancy and the early postpartum period in harbor seals, total corticosteroids were high and ranged widely, up to nearly 40 µg/dl (Raeside and Ronald, 1981), although Gardiner and Hall (1997) found no significant difference in cortisol levels between a pregnant and a nonpregnant captive harbor seal. Harp seals have higher levels while lactating than during the postlactation period (Engelhardt and Ferguson, 1980).
0839_frame_C10 Page 179 Tuesday, May 22, 2001 11:08 AM
179
Endocrinology
TABLE 5 Reported Concentrations (µg/dl) of Cortisol in Marine Mammals Species
Specimens
Cortisol (µg/dl)
Reference
Cetaceans Balaenoptera acutorostrata (minke whale) Balaenoptera physalus (fin whale) Cephalorhynchus commersonii (Commerson’s dolphin) Delphinapterus leucas (beluga) Globicephala macrorhynchus (short-finned pilot whale) Grampus griseus (Risso’s dolphin) Inia geoffrensis (Amazon River dolphin) Kogia breviceps (pygmy sperm whale) Lagenorhynchus obliquidens (Pacific white-sided dolphin) Orcinus orca (killer whale) Phocoena phocoena (harbor porpoise) Phocoenoides dalli (Dall’s porpoise) Pseudorca crassidens (false killer whale) Stenella coeruleoalba (blue-white dolphin) Tursiops truncatus (bottlenose dolphin)
Various ages, both sexes Not specified
0.33
Suzuki et al., 1998
1.0–1.2
Not specified
0.5
Kjeld and Olafsson, 1987; Kjeld and Theodórsdóttir, 1991 Suzuki et al., 1998
Various ages, both sexes Not specified
0.7–3.2 0.4–0.7
Suzuki et al., 1998; St. Aubin et al., in press Suzuki et al., 1998
Not specified
0.9
Suzuki et al., 1998
Not specified
0.8
Suzuki et al., 1998
Not specified
0.2
Suzuki et al., 1998
Not specified
0.8
Suzuki et al., 1998
Not specified
0.4
Suzuki et al., 1998
Not specified Both sexes, age unspecified Not specified
0.4 8.8
Suzuki et al., 1998 Koopman et al., 1995
0.7
Suzuki et al., 1998
Not specified
0.7
Suzuki et al., 1998
Not specified
0.5
Suzuki et al., 1998
Various ages, both sexes
0.6–3.6
Thompson and Geraci, 1986; Orlov et al., 1988; St. Aubin et al., 1996; Suzuki et al., 1998; Ortiz and Worthy, 2000
Pinnipeds Halichoerus grypus (gray seal)
Leptonychotes weddellii (Weddell seal)
Pups (1–2 wk) Juveniles, molting and non-molting Juveniles, molting Adult
Adult male, breeding Adult
4.3 6.3–9.1
Engelhardt and Ferguson, 1980 Boily, 1996; Lohmann et al., 1998
4.5 3.6–5.9
Boily, 1996 Sangalang and Freeman, 1976; Engelhardt and Ferguson, 1980; Boily, 1996 Sangalang and Freeman, 1976; Engelhardt and Ferguson, 1980 Liggins et al., 1979; Barrell and Montgomery, 1989; Bartsh et al., 1992
21.2–35.4 69–153.9
(Continued)
0839_frame_C10 Page 180 Tuesday, May 22, 2001 11:08 AM
180
CRC Handbook of Marine Mammal Medicine
TABLE 5 Reported Concentrations (µg/dl) of Cortisol in Marine Mammals (continued) Species Pagophilus groenlandicus (harp seal)
Phoca hispida (ringed seal) Phoca largha (spotted seal) Phoca vitulina (harbor seal)
Specimens Neonate Pups (<1 wk) Pups (3 wk) Juvenile Adult female Lactating Adult male Juvenile Juveniles Adults
Cortisol (µg/dl) 5.3 1.8–2.5 5.5 11–15 3.6 8.2 11 12–20 4–16 7–24
Juveniles
3–8.6
Juveniles, molting
9–12
Adult female Prepartum (1–80 d) Postpartum (5 d)
8–16 6–16.4 39.2
Reference Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 St. Aubin and Geraci, 1986 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 St. Aubin and Geraci, 1986 Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Riviere et al., 1977; Ashwell-Erickson et al., 1986; Gulland et al., 1999 Riviere et al., 1997; Ashwell-Erickson et al., 1986 Ashwell-Erickson et al., 1986 Raeside and Ronald, 1981 Raeside and Ronald, 1981
Sirenians Trichechus manatus (West Indian and Florida manatees)
Age unspecified, both sexes
Enhydra lutris
Various ages, both sexes
Ursus maritimus
Adults, both sexes
0.15
Ortiz et al., 1998
Sea Otter 3.2–3.9
Williams et al., 1992
Polar Bear 6.9–54
Cattet, 2000
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. µg/dl × 27.59 = nmol/l.
Wild harbor seals show significant seasonal variation in cortisol levels, correlating with both breeding and molting; levels were lower during the breeding/molt season than at other times of the year (Gardiner and Hall, 1997). As for thyroid hormones, temporal associations between cortisol changes and overt signs of molt may be misleading, and asynchrony may simply reflect the slow development of follicular changes, either for hair loss or regrowth. The seasonal differences in cortisol were not evident in captive seals. Glucocorticoids have received the greatest attention in the literature for their role in the stress response; this subject is reviewed in more detail elsewhere in this volume (see Chapter 13, Stress). The basic physiology of glucocorticoid secretion has been investigated through the use of exogenous ACTH in cetaceans (Thomson and Geraci, 1986; St. Aubin and Geraci, 1990), phocids (St. Aubin and Geraci, 1986), and otariids (St. Aubin et al., unpubl. data). Gulland et al. (1999) used ACTH stimulation tests to assess adrenal function in harbor seals infected with an adrenotropic herpes virus. Dosages have ranged from 0.2 IU/kg in belugas, to 0.25 to
0839_frame_C10 Page 181 Tuesday, May 22, 2001 11:08 AM
Endocrinology
181
0.4 IU/kg in bottlenose dolphins, to 1 IU/kg in harbor seal pups. Various synthetic (Cortrosyn®, Organon Canada, Ltd., Toronto, Ontario, and Repository Corticotropin, Austin, Jolliette, Canada) and natural porcine (ACTHar®, Harris Laboratories, Toronto, Canada) preparations have been effective in elevating serum cortisol levels within 30 to 60 min of administration. Systemic consequences of rising cortisol concentrations, including stress leukograms (leukocytosis, lymphopenia, and eosinopenia) and hyperglycemia, indicate that the pituitary– adrenal axis functions for the most part in accordance with the relationships established for other mammals. A significant difference, however, lies in the relatively low circulating levels of cortisol in cetaceans, and the modest increases observed following stimulation, a condition that confounds the use of cortisol as a diagnostic indicator of stress in these animals. Yet, the characteristic changes in other circulating constituents normally sensitive to elevations in cortisol suggest that even small changes may be clinically important. A possible explanation for the difference between pinnipeds and cetaceans in this respect is the hormone-binding capacity of the plasma. Whereas more than 90% of cortisol is bound in the Weddell seal (Liggins et al., 1979), studies on belugas and bottlenose dolphins indicate that the bound fraction represents 50% or less of the total hormone (St. Aubin and Geraci, unpubl. data). Small increments might therefore translate into relatively more free hormone and greater availability to exert effects on the organism. Although the importance of aldosterone is suggested by the prominence of the zona glomerulosa cells that produce it, particularly in young seals, its value in marine mammals is enigmatic. There would appear to be little advantage to conserving sodium in an environment in which the greater need would be to excrete it. Nevertheless, aldosterone is detectable in most samples drawn from marine mammals (Table 6). Levels tend to be higher in young phocids (Engelhardt and Ferguson, 1980), as might be expected from the histological appearance of the gland (Amoroso et al., 1965). Manatees, whether in the wild or in captivity, have higher aldosterone concentrations when in fresh than in salt water (Ortiz et al., 1998). By contrast, belugas sampled in fresh and marine waters showed wide ranges for plasma sodium and aldosterone concentrations (St. Aubin et al., in press), with no significant correlation between the two constituents and the environment in which they were sampled. This apparently casual approach to electrolyte regulation in belugas contrasts with significant clinical problems associated with both hyper- and hyponatremia in other species, particularly some phocid seals. Chronic salt deprivation produced widely fluctuating, but generally elevated, plasma levels of aldosterone in a ringed seal (Phoca hispida) that was able to maintain normonatremia (St. Aubin and Geraci, 1986). Salt deprivation in a second ringed seal resulted in mild hyponatremia (Na: 142 to 145 mEq/l), and slightly reduced but variable plasma aldosterone levels; a spontaneously hyponatremic harp seal with sodium concentrations of 115 to 130 mEq/l had low but still detectable aldosterone. Thus, while hyponatremia can occur in the presence of seemingly adequate levels of the hormone, it appears that sodium conservation in these phocids is achieved by increasing aldosterone. Elevated aldosterone during the postweaning fast in northern elephant seals appears to be a strategy to help conserve water, which is resorbed along with sodium (Ortiz et al., 2000). Electrolyte imbalance is not the sole stimulus for aldosterone secretion; angiotensin II (AII), which will be addressed later, and ACTH both play a role. It is the particular sensitivity of the zona glomerulosa to the latter that distinguishes marine from terrestrial mammals. In contrast to the modest increases found in humans and other mammals, elevations in aldosterone as high as sevenfold have been noted in ringed (St. Aubin and Geraci, 1986) and harbor (Gulland et al., 1999) seals, northern fur seals (St. Aubin et al., unpubl. data), bottlenose dolphins (Thomson and Geraci, 1986), and belugas (St. Aubin and Geraci, 1990). Sodium conservation during times of stress apparently is an important requirement shared by a variety of species adapted to the
0839_frame_C10 Page 182 Wednesday, May 23, 2001 10:43 AM
182
CRC Handbook of Marine Mammal Medicine
TABLE 6 Reported Concentrations of Aldosterone (pg/ml) in Marine Mammals Species
Aldosterone (pg/ml)
Specimens
Reference
Cetaceans Balaenoptera physalus (fin whale) Delphinapterus leucas (beluga) Tursiops truncatus (bottlenose dolphin)
Not specified
17–168
Various ages, both sexes Adults, both sexes
203–450 3–677
Kjeld and Olafsson, 1987; Kjeld and Theodórsdóttir, 1991 St. Aubin and Geraci, 1989; St. Aubin et al., 2001 Malvin et al., 1978; Thomson and Geraci, 1986; St. Aubin et al., 1996
Pinnipeds Mirounga angustirostris (northern elephant seal) Pagophilus groenlandicus (harp seal)
Phoca hispida (ringed seal) Phoca vitulina (harbor seal) Halichoerus grypus (gray seal) Zalophus californianus (California sea lion)
Pups, at weaning Pups, fasting 5–7 wk Neonates Pups (<2 wk) Juveniles Adults Adults Juveniles
220 1000 2250 600–1180 300 400–1200 140–1040 750–1110
Adults
1400–3200
Juveniles
140–310
Ortiz et al., 2000 Ortiz et al., 2000 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 Engelhardt and Ferguson, 1980 St. Aubin and Geraci, 1986 Gulland et al., 1999 Sangalang and Freeman, 1976 Malvin et al., 1978
Sirenians Trichechus manatus (West Indian and Florida manatees)
Both sexes (fresh water) Both sexes (brackish and salt water)
660 37–95
Ortiz et al., 1998 Ortiz et al., 1998
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source. Some of the data were estimated from figures. pg/ml × 2.775 = pmol/l.
marine environment. In ringed seals stressed by salt restriction, the aldosterone response to ACTH stimulation is exaggerated until the zona glomerulosa is exhausted (Figure 1). Pinniped hyponatremia, which can occur under conditions other than Na deprivation, may thus be a consequence of adrenal failure precipitated by chronic stress (Geraci, 1972; St. Aubin and Geraci, 1986).
Osmoregulatory Hormones The classification of a subset of mammals as “marine” might suggest the presence of hormonemediated physiological adaptations to cope with a substantially hypertonic environment. In fact, with the exception of the large size of the kidney in the sea otter, renal tubular morphology and function in marine mammals are unremarkable, and render unnecessary the requirement for unusual endocrine pathways to manage water and electrolytes. Nevertheless, other aspects of marine mammal life histories, such as prolonged fasting in pinnipeds, place particular demands on these systems and have been the subject of numerous investigations. It is these adaptations that represent the more important concerns for the clinician.
0839_frame_C10 Page 183 Tuesday, May 22, 2001 11:08 AM
Endocrinology
183
FIGURE 1 Plasma aldosterone concentration in five seals following intramuscular injection of ACTH. Two saltsupplemented ringed seals maintained in salt water served as normal controls ( ------ and ■ ------■). One ringed seal remained normonatremic when salt deprived (———), while another became hyponatremic (------). A harp seal spontaneously developed hyponatremia while being held in salt water and receiving dietary salt supplements (------). (Redrawn from St. Aubin and Geraci, 1986.)
Vasopressin An antidiuretic hormone (ADH) was demonstrated in pituitaries from commercially harvested whales by researchers in the 1930s. Specific RIAs on seal pituitary extracts (Dogterom et al., 1980) and plasma from various pinnipeds, cetaceans, and sirenians (Table 7) suggest that the form elaborated by marine mammals is arginine vasopressin (AVP). In general, the range of reported AVP concentrations noted in pinnipeds and cetaceans is higher than that in manatees. None of the reported values is unusual relative to most other mammals. The dynamics of AVP during various physiological stresses challenge conventional expectations based on the role of this hormone in other mammals. During their prolonged postweaning fast, northern elephant seal pups showed declining levels of both AVP and urinary output (Ortiz et al., 1996); a subsequent study on the same species found no change in AVP during the fast (Ortiz et al., 2000). The hormone thus appears to be inconsequential in water conservation at this time. In fasted gray seals, AVP levels increased as much as threefold, an expected response that is likely tied to the concurrently increasing urinary osmolality (Skog and Folkow, 1994). However, water loading in gray seals failed to suppress AVP, and the excess fluid was cleared in a large volume of dilute urine despite the persistence of elevated AVP levels in circulation. A significant role for AVP could not be demonstrated in bottlenose dolphins in an early study monitoring urinary flow and osmolality (Malvin et al., 1971). Other physiological actions of AVP have been explored. Ortiz and Worthy (2000) considered the relationship between AVP and adrenal corticosteroids during capture stress in bottlenose dolphins; the lack of correlation suggested that AVP did not induce changes in ACTH, as it does in other mammals. Bradycardia during resting apnea in Weddell and northern elephant
0839_frame_C10 Page 184 Tuesday, May 22, 2001 11:08 AM
184
CRC Handbook of Marine Mammal Medicine
TABLE 7 Reported Concentrations of Arginine Vasopressin (AVP) (pg/ml), Angiotensin II (AII) (pg/ml), and Atrial Natriuretic Peptide (ANP) (pg/ml) in Marine Mammals Species Tursiops truncatus (bottlenose dolphin) Eumetopias jubatus (Steller sea lion)
Specimens
AVP
AII
ANP
Reference
Both sexes, ages unspecified Pups
3.3
—
—
Ortiz and Worthy, 2000
7.2
46.9
Yearling, subadult Adults
6.2–6.5
20.5–24.6
Leptonychotes weddellii (Weddell seal) Mirounga angustirostris (northern elephant seal) Phoca hispida (ringed seal) Phoca vitulina (harbor seal)
Various ages
3.2–7.2
12.2–39.6
12.5–30.6
Pups
1.5–28
16.5–33.2
20.9–26.3
Adults
9.3
14.0
Various ages
7.2–13.3
29.0
Trichechus manatus (West Indian and Florida manatees)
Fresh water Salt water and brackish water (wild) Salt water (captive)
0.6–1.1 2.1–2.5
— —
— —
Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998a,b Zenteno-Savin and Castellini, 1998a,b; Ortiz et al., 1996; 2000 Zenteno-Savin and Castellini, 1998b Zenteno-Savin and Castellini, 1998b; Ellsworth et al., 1999 Ortiz et al., 1998 Ortiz et al., 1998
0.5
—
—
Ortiz et al., 1998
Zalophus californianus (California sea lion)
88.3 6.5–32
14.2
55.8
139.3
Pups
4.7
7.6
26.9
Adult
10.2
8.4
31.7
126.8 12.2–66.8
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source.
seals is associated with rapid decreases in AVP, a response that develops with age in the latter species (Zenteno-Savin and Castellini, 1998a) The closely related Baikal (Phoca sibirica) and ringed seals were studied for evidence of hormonal differences associated with specific needs for water conservation in their respective environments (Hong et al., 1982). Urinary ADH (AVP) concentration, expressed relative to that of creatinine, was similar in both species, and increased during water deprivation and fasting. After water loading, the hormone was undetectable. Thus, the Baikal seal exhibited no obvious adaptations in this mode of water management after an estimated half-million years of isolation in fresh water. Manatees naturally occur in habitats of varying salinity and, in the wild, show differences in blood AVP consistent with the expected need to conserve or eliminate water (see Table 7) (Ortiz et al., 1998). Paradoxically, captive manatees in salt water have lower AVP concentrations than those in fresh water, although the differences are small and insignificant. Perhaps the low salt content of the lettuce diet offered to the captive animals, compared with that in the natural marine vegetation, can account for the reduced need for water conservation in the former environment (see Chapter 36, Nutrition). Overall, plasma AVP and osmolality were significantly correlated.
0839_frame_C10 Page 185 Tuesday, May 22, 2001 11:08 AM
Endocrinology
185
Renin–Angiotensin System The scant information on the renin–angiotensin system (RAS) in marine mammals is surprising in light of the profound changes in blood pressure and flow associated with at least some stages of the dive response. Secreted from juxtaglomerular cells of the kidney in response to hypotension in afferent arterioles, renin converts angiotensin I released from lung cells to AII, a potent vasoconstrictor. Renin activity is measured according to the rate of production of AII, while AII levels are determined directly by RIA. A recent survey of AII concentrations in pinnipeds found concentrations similar to those in most other mammals (see Table 7) (Zenteno-Savin and Castellini, 1998b). Early studies by Malvin and co-workers (Malvin and Vander, 1967; Malvin et al., 1978) focused on the RAS from the perspective of its role in osmoregulation in cetaceans and pinnipeds. Renin and aldosterone levels were significantly correlated in bottlenose dolphins, California sea lions (Zalophus californianus), and northern elephant seals, suggesting that this arm of aldosterone control is functional in at least some marine mammals (Malvin et al., 1978; Ortiz et al., 2000). The only insight into the dynamics of AII during diving comes from observations during apnea in Weddell and northern elephant seals (Zenteno-Savin and Castellini, 1998a). The observed decrease in AII levels appears to be inconsistent with a presumed rise in renin resulting from reduced blood flow to the kidney. The authors speculated that a response in AII might be delayed until circulation to the kidney is reestablished and renin is delivered systemically.
Atrial Natriuretic Peptide First described in the literature as atrial natriuretic “factor” in the mid-1980s, this substance received growing attention in the human medical literature, including its characterization as a peptide and subsequent renaming as atrial natriuretic peptide (ANP). Consideration of its presence in marine mammals began with the demonstration of characteristic secretory granules in cardiomyocytes of ringed, harp, and northern elephant seals (Pfeiffer and Viers, 1995; Tagoe et al., 1998). The significance of the osmoregulatory function of this hormone was questionable in elephant seals, particularly, because of the sparseness of the structure. Investigation of the activity of the hormone in marine mammals is limited to a survey of circulating levels in some pinnipeds (Zenteno-Savin and Castellini, 1998b) and two functional studies. “Resting” levels are comparable to those measured in other mammals (see Table 7). Concentrations increase during apnea in Weddell seals, but not northern elephant seals (ZentenoSavin and Castellini, 1998a). The only experiment to examine the osmoregulatory action of this hormone yielded inconclusive results (Ellsworth et al., 1999). Named for its action, ANP responds more consistently to volumetric expansion, and the resultant atrial stretch, than to sodium loading, although levels do increase with sodium burden. The consequence of natriuresis serves to rectify hypervolemia rather than correct hypernatremia. Nevertheless, intravenous administration of up to 2 l of normal saline in a 1-hour period in adult harbor seals failed to consistently produce the expected increase in circulating ANP. It was postulated that the large, distensible vascular reservoirs in these animals dampened the intended stimulus, which had produced consistent changes in comparably sized humans. The functional significance of this hormone in phocids, at least, remains a question for further research.
Endocrine Pancreas Insulin extracted from mysticete and sperm whale pancreas was found to have the same amino acid sequence as the porcine hormone, and to differ from the human form by only a single amino acid (Hama et al., 1964). Although the structure of glucagon has not been reported, it
0839_frame_C10 Page 186 Tuesday, May 22, 2001 11:08 AM
186
CRC Handbook of Marine Mammal Medicine
TABLE 8 Reported Concentrations of Insulin (µU/ml) and Glucagon (pg/ml) in Marine Mammals Species Tursiops truncatus (bottlenose dolphin) Mirounga angustirostris (northern elephant seal)
Phoca vitulina (harbor seal) Ursus maritimus (polar bear)
Specimens
Insulin
Glucagon
Reference
12–70 h fast Postprandial Not specified Pup (<3 wk) Weaned (2– 4 wk) molting, fasting Weaned (4–11 wk) fasting Lactating (1–4 wk) Adult, molting Pre-dive
11.2 12.3 10 9.9–11.3 9.2
94 117 — 195–844 153–346
Patton et al., 1977 Patton et al., 1977 Orlov et al., 1988 Kirby, 1990 Kirby, 1990
7.2–8.1 8.9–11.9 4.3 4–12
179–363 — 145–379 30–75
Kirby, 1990 Kirby, 1990 Kirby, 1990 Robin et al., 1981
3–96
18–637
Adults, feeding and fasting
Cattet, 2000
Note: Values given as a range of means from multiple publications, or either the mean or range (when available) from a single source.
is similar enough to that in other mammals to be measured using methods developed for other species. Both hormones are presumed to function in marine mammals as they do in other mammals. With a diet typically very low in carbohydrates, marine mammals sustain their glucose requirements principally through gluconeogenesis. As such, the hormones responsible for glucose homeostasis, insulin and glucagon, are balanced to deliver glucose into circulation rather than promote its uptake. The ratio of insulin to glucagon is consequently very low in virtually all groups studied (Table 8). The exception is the polar bear, in which insulin concentrations invariably exceed those of glucagon. Given that the polar bear’s diet for much of the year is also devoid of carbohydrate, the reversed relationship probably reflects a physiology more reminiscent of that of a terrestrial mammal. Insulin levels in harbor seals, elephant seals, and bottlenose dolphins were mostly unaffected during glucose tolerance tests, but were increased in the latter following protein meals and oral arginine (Ridgway et al., 1970; Patton, 1977; Patton et al., 1977; Kirby, 1990). The blunted response in these animals undermines the utility of conventional approaches to assess pancreatic function. Such information might be particularly useful in species such as harbor porpoises, which commonly show extensive pancreatic fibrosis as a result of trematode infections. The more important role for insulin and glucagon in marine mammals is maintaining circulating levels of glucose for delivery to the brain during dives. The ratio of insulin with respect to glucagon falls during voluntary dives in Weddell seals and contributes to hyperglycemia at the end of the dive (Hochachka et al., 1995).
Future Studies Although the basic framework of marine mammal endocrinology has essentially been described, intriguing questions remain regarding the dynamics of some of these systems during physiologically challenging conditions such as diving and fasting. The measurement of circulating levels is only one index of hormone activity, and can sometimes be misleading. Binding proteins, metabolic clearance rate, and cell receptor density all play a role in modulating the actions of hormones, but the information on these points for marine mammals is sparse or nonexistent. The development of specific assays for peptide hormones will lead to a better
0839_frame_C10 Page 187 Tuesday, May 22, 2001 11:08 AM
Endocrinology
187
understanding of factors, such as GH and tropic hormones, that may show important changes during the life history of these animals. Advances in the fundamental endocrinology of marine mammals will also improve our ability to recognize the effects of environmental contaminants that can disrupt endocrine systems.
Acknowledgments The author is grateful to Pauline Schwalm for her assistance in tabulating the reported hormone data and to Shannon Atkinson and Ailsa Hall for their helpful reviews. Data on thyroid and adrenal hormones in northern fur seals at Mystic Aquarium were collected in collaboration with Thom Lembo and Larry Dunn, with support from the staff of the Departments of Husbandry and Research and Veterinary Services. These studies were funded by Mystic Aquarium and the Bernice Barbour Foundation. The Office of Naval Research and the Naval Ocean Systems Center (now NCCOSC RDTE), through Sam Ridgway, supported the research on hormone cycles in captive belugas. Joseph Geraci is thanked for fostering the author’s early interest in marine mammal endocrinology, and for providing the guidance, resources and encouragement to pursue those inquiries. This is contribution number 120 from the Sea Research Foundation.
References Amoroso, E.C., Bourne, G.H., Harrison, R.J., Matthews, L.H., Rowlands, I.W., and Sloper, J.C., 1965, Reproductive and endocrine organs of foetal, newborn and adult seals, J. Zool., 147: 430–486. Arvy, L., 1970, Endocrine glands and hormonal secretion in cetaceans, in Investigations on Cetacea, Pilleri, G. (Ed.), Vol. 3, Part 2, Waldau, Berne, Switzerland, 230–251. Ashwell-Erickson, S., Fay, F.H., Elsner, R., and Wartzok, D., 1986, Metabolic and hormonal correlates of molting and regeneration of pelage in Alaskan harbor and spotted seals (Phoca vitulina and Phoca largha), Can. J. Zool., 64: 1086–1094. Barrell, G.K., and Montgomery, G.W., 1989, Absence of circadian patterns of secretion of melatonin or cortisol in Weddell seals under continuous natural daylight, J. Endocrinol., 122: 445–449. Bartsh, S.S., Johnson, S.D., and Siniff, D.B., 1992, Territorial behavior and breeding frequency of male Weddell seals (Leptonychotes weddellii) in relation to age, size, and concentrations of serum testosterone and cortisol, Can. J. Zool., 70: 680–692. Behrmann, G., 1990, The pineal organ (epiphysis cerebri) of the harbor porpoise, Phocoena phocoena, Aquat. Mammals, 16: 96–100. Boily, P., 1996, Metabolic and hormonal changes during the molt of captive gray seals (Halichoerus grypus), Am. J. Physiol., 270 (5 Pt 2): R1051–1058. Borruel, M., Borruel, P., Damasco, M.C., and Lantos, C.P., 1974, The in vitro investigation of four radioactive corticosteroids in the Antarctic seal (Leptonychotes weddellii), Gen. Comp. Endocrinol., 22: 1–12. Brouwer, A., Reijnders, P.J.H., and Koeman, J.H., 1989, Polychlorinated biphenyl (PCB) induces vitamin A and thyroid hormone deficiency in the common seal (Phoca vitulina), Aquat. Toxicol., 15: 99–106. Brown, J.L., Wasser, S.K., Wildt, D.E., and Graham, L.H., 1994, Comparative aspects of steroid hormone metabolism and ovarian activity in felids, measured noninvasively in feces, Biol. Reprod., 51: 776–786. Bryden, M.M., 1994, Endocrine changes in newborn southern elephant seals, in Elephant Seals: Population Ecology, Behavior and Physiology, LeBoeuf, B.J., and Laws, R.M. (Eds.), University of California Press, Berkeley, 387–397.
0839_frame_C10 Page 188 Tuesday, May 22, 2001 11:08 AM
188
CRC Handbook of Marine Mammal Medicine
Bryden, M.M., Griffiths, D.J., Kennaway, D.J., and Ledingham, J., 1986, The pineal gland is very large and active in newborn Antarctic seals, Experientia, 42: 564–566. Bryden, M.M., Buckendahl, P., Sanders, J., Ortiz, C.L., and Kennaway, D.J., 1994, Plasma melatonin concentration in neonatal northern elephant seals, Mirounga angustirostris, Comp. Biochem. Physiol. A, 109: 895–904. Carballeira, A., Brown, J.W., Fishman, L.M., Trujillo, D., and Odell, D.K., 1987, The adrenal gland of stranded whales (Kogia breviceps and Mesoplodon europaeus): Morphology, hormonal contents, and biosynthesis of corticosteroids, Gen. Comp. Endocrinol., 68: 293–303. Cattet, M.R.L., 2000, Biochemical and Physiological Aspects of Obesity, High Fat Diet, and Prolonged Fasting in Free-Ranging Polar Bears, Ph.D. thesis, University of Saskatchewan, Saskatoon, Saskatchewan, Canada. Cuello, A.C., and Tramezzani, J.H., 1969, The epiphysis cerebri of the Weddell seal: Its remarkable size and glandular pattern, Gen. Comp. Endocrinol., 12: 154–164. Dawbin, W.H., 1966, The seasonal migratory cycle of humpback whales, in Whales, Dolphins and Porpoises, Norris, K.S. (Ed.), University of California Press, Berkeley, 145–170. DeGuise, S., Lagacé, A., and Béland, P., 1994, Tumors in St. Lawrence beluga whales (Delphinapterus leucas), Vet. Pathol., 31: 444–449. DeRoos, C.C., and Bern, H.A., 1961, The corticoids of the adrenal of the California sea lion Zalophus californianus, Gen. Comp. Endocrinol., 1: 275–285. Dogterom, J.F., Snijdewint, G.M., Pevet, P., and Swaab, D.F., 1980, Studies on the presence of vasopressin, oxytocin and vasotocin in the pineal gland, subcommissural organ and fetal pituitary gland: Failure to demonstrate vasotocin in mammals, J. Endocrinol., 84: 115–123. Ekins, R., 1986, The free hormone concept, in Thyroid Hormone Metabolism, Hennemann, G. (Ed.), Marcel Dekker, New York, 77–106. Elden, C.A., Keyes, M.C., and Marshall, C.E., 1971, Pineal body of the northern fur seal (Callorhinus ursinus): A model for studying the probable function of the mammalian pineal body, Am. J. Vet. Res., 32: 639–647. Ellsworth, L.B., St. Aubin, D.J., Dunn, J.L., and Zenteno-Savin, T., 1999, Effects of saline infusions on circulating levels of plasma atrial natriuretic peptide in harbor seals (Phoca vitulina), in Proceedings of the 13th Biennial Conference on the Biology of Marine Mammals, Maui, HI, Nov. 28–Dec. 3, p. 52. Engelhardt, F.R., and Ferguson, J.M., 1980, Adaptive changes in harp seals, Phoca groenlandicus, and gray seals, Halichoerus grypus, during the post-natal period, Gen. Comp. Endocrinol., 40: 434–445. Flanigan, N., 1972, The central nervous system, in Mammals of the Sea: Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 215–246. Gallivan, G.J., and Best, R.C., 1980, Metabolism and respiration of the Amazonian manatee (Trichechus inunguis), Physiol. Zool., 53: 245–253. Gardiner, K.J., and Hall, A.J., 1997, Diel and annual variation in plasma cortisol concentrations among wild and captive harbor seals (Phoca vitulina), Can. J. Zool., 75: 1773–1780. Geraci, J.R., 1972, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618–623. Goldman, B.D., 1983, The physiology of melatonin in mammals, Pineal Res. Rev., 1: 145–182. Greenwood, A.G., and Barlow, C.E., 1979, Thyroid function in dolphins: Radioimmunoassay measurement of thyroid hormones, Br. Vet. J., 135: 96–102. Griffiths, D.J., 1985, Endocrine regulation of seasonal breeding in the male southern elephant seal (Mirounga leonina) at Macquarie Island, in Studies of Sea Mammals in South Latitudes, Ling, J.K., and Bryden, M.M. (Eds.), South Australia Museum, Graphic Services Pty. Ltd., Northfield, S.A., 31–40. Griffiths, D.J., and Bryden, M.M., 1981, The annual cycle of the pineal gland of the elephant seal (Mirounga leonina), in Pineal Function, Matthews, C.D., and Seamark, R.F. (Eds.), Elsevier NorthHolland Biomedical Press, Amsterdam, 57–66.
0839_frame_C10 Page 189 Tuesday, May 22, 2001 11:08 AM
Endocrinology
189
Griffiths, D., Seamark, R.F., and Bryden, M.M., 1979, Summer and winter cycles in plasma melatonin levels in the elephant seal (Mirounga leonina), Aust. J. Biol. Sci., 32: 581–586. Gulland, F.M.D., Haulena, M., Lowenstine, L.J., Munro, C., Graham, P.A., Bauman, J., and Harvey, J., 1999, Adrenal function in wild and rehabilitated Pacific harbor seals (Phoca vitulina richardii) and in seals with phocine herpesvirus-associated adrenal necrosis, Mar. Mammal Sci., 15: 810–827. Hall, A.J., Green, N.J.L., Jones, K.C., Pomeroy, P.P., and Harwood, J., 1998, Thyroid hormones as biomarkers in grey seals, Mar. Pollut. Bull., 36: 424–428. Hama, H., Titani, K., Sakaki, S., and Narita, K., 1964, The amino acid sequence in fin-whale insulin, J. Biochem., 56: 285–293. Hance, A.J., Robin, E.D., Halter, J.B., Lewiston, N., Robin, D.A., Cornell, L., Caligiuri, M., and Theodore, J., 1982, Hormonal changes and enforced diving in the harbor seal Phoca vitulina, II, Plasma catecholamines, Am. J. Physiol., 242: R528–R532. Harrison, R.J., 1969, Endocrine organs: Hypophysis, thyroid, and adrenal, in The Biology of Marine Mammals, Anderson, H.T. (Ed.), Academic Press, London, 349–390. Harrison, R.J., and Young, B.A., 1970, The thyroid gland of the common (Pacific) dolphin, Delphinus delphis bairdi, J. Anat., 106: 243–254. Harrison, R.J., Rowlands, I.W., Whitting, H.W., and Young, B.A., 1962, Growth and structure of the thyroid gland in the common seal (Phoca vitulina), J. Anat., 96: 3–15. Haulena, M., St. Aubin, D.J., and Duignan, P.J., 1998, Thyroid hormone dynamics during the nursing period in harbour seals, Phoca vitulina, Can. J. Zool., 76: 48–55. Heldmaier, G., Steinlaechner, S., Rafael, J., and Vsiansky, P., 1981, Photoperiodic control and effects of melatonin on nonshivering thermogenesis and brown adipose tissue, Science, 212: 917–919. Hochachka, P.W., Liggins, G.C., Guyton, G.P., Schneider, R.C., Staneck, K.S., Hurford, W.E., Creasy, R.K., Zapol, D.G., and Zapol, W.M., 1995, Hormonal regulatory adjustments during voluntary diving in Weddell seals, Comp. Biochem. Physiol. B, 112: 361–375. Hong, S.K., Elsner, R., Claybaugh, J.R., and Ronald, K., 1982, Renal functions of the Baikal seal Pusa sibirica and ringed seal Pusa hispida, Physiol. Zool., 55: 289–299. Iverson, S.J., Bowen, W.D., Boness, D.J., and Oftedal, O.T., 1993, The effect of maternal size and milk energy output on pup growth in grey seals (Halichoerus grypus), Physiol. Zool., 66: 61–88. John, T.M., McKeown, B.A., Geaorge, J.C., and Ronald, K., 1980, Plasma levels of growth hormone and free fatty acids in the harp seal, Comp. Biochem. Physiol. B, 66: 159–162. John, T.M., Ronald, K., and George, J.C., 1987, Blood levels of thyroid hormones and certain metabolites in relation to moult in the harp seal (Phoca groenlandicus), Comp. Biochem. Physiol. A, 88: 655–657. Kawauchi, H., 1980, Isolation and primary structure of whale and fish pituitary hormones, Canadian Translation, Fisheries Aquatic Science No. 4642, Department of Fisheries and Oceans, Nanaimo, BC, 24. Kawauchi, H., and Tubokawa, M., 1979, Isolation and characterization of fin whale prolactin, Int. J. Peptide Protein Res., 13: 229–234. Kawauchi, H., Muramoto, K., and Ramachandran, J., 1978, Isolation and primary structure of adrenocorticotropin from several species of whales, Int. J. Peptide Protein Res., 12: 318–324. Kirby, V.M., 1990, Endocrinology of marine mammals, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 303–351. Kjeld, M., and Olafsson, I., 1987, Some biochemical parameters in blood and urine of fin whales (Balaenoptera physalus), Isr. J. Vet. Med., 43: 117–121. Kjeld, M., and Theodórsdóttir, A., 1991, Some electrolytes, hormones and other substances in the blood of fin whales of the coast of Iceland, Náttúrufrœdingurinn, 60: 147–154. Koopman, H.N., Westgate, A.J., Read, A.J., and Gaskin, D.E., 1995, Blood chemistry of wild harbor porpoises, Phocoena phocoena (L.), Mar. Mammal Sci., 11: 123–135. Lavigne, D.M., Innes, S., Worthy, G.A.J., Kovacs, K.M., Schmitz, O.J., and Hickie, J.P., 1986, Metabolic rates of seals and whales, Can. J. Zool., 64: 279–284.
0839_frame_C10 Page 190 Tuesday, May 22, 2001 11:08 AM
190
CRC Handbook of Marine Mammal Medicine
Leatherland, J.F., 1976, Structure of the adenohypophysis in juvenile harp seal, Pagophilus groenlandicus, Cell Tissue Res., 173: 367–382. Leatherland, J.F., and Ronald, K., 1978, The structure of the hypophysis in parturient and neonatal harp seals, Pagophilus groenlandicus, Cell Tissue Res., 192: 341–357. Leatherland, J.F., and Ronald, K., 1979, Thyroid activity in adult and neonate harp seals Pagophilus groenlandicus, J. Zool., 189: 399–405. Leatherland, J.F., and Ronald, K., 1981, Plasma concentrations of thyroid hormones in a captive and feral polar bear, Comp. Biochem. Physiol. A, 70: 575–578. Leatherland, J.F., and Ronald, K., 1983, Immunohistochemical identification of cell types in the pars distalis of harp seals, Phoca groenlandicus, Acta Zool. (Stockholm), 64: 97–106. Liggins, G.C., France, J.T., Knox, B.S., and Zapol, W.M., 1979, High corticosteroid levels in plasma of adult and foetal Weddell seals (Leptonychotes weddellii), Acta Endocrinol., 90: 718–726. Little, G.J., 1991, Thyroid morphology and function and its role in thermoregulation in the newborn southern elephant seal (Mirounga leonina) at Macquarie Island, J. Anat., 176: 55–69. Little, G.J., and Bryden, M.M., 1990, The pineal gland in newborn southern elephant seals, Mirounga leonina, J. Pineal Res., 9: 139–148. Lohman, S., Folkow, L.P., Osterud, B., and Sager, G., 1998, Changes in fibrinolytic activity in diving grey seals, Comp. Biochem. Physiol. A, 120: 693–698. Lydersen, C., Hammill, M.O., and Kovacs, K.M., 1995, Milk intake, growth and energy consumption in pups of ice-breeding grey seals (Halichoerus grypus), J. Comp. Physiol. B, 164: 585–592. Lydersen, C., Kovacs, K.M., and Hammill, M.O., 1997, Energetics during nursing and early postweaning fasting in hooded seal (Cystophora cristata) pups from the Gulf of St. Lawrence, Canada, J. Comp. Physiol. B, 167: 81–88. Malvin, R.L., and Vander, A.J., 1967, Plasma renin activity in marine teleosts and cetacea, Am. J. Physiol., 213: 1582–1584. Malvin, R.L., Bonjour, J.P., and Ridgway, S.H., 1971, Antidiuretic hormone levels in some cetaceans, Proc. Soc. Exp. Biol. Med., 136: 1203–1205. Malvin, R.L., Ridgway, S.H., and Cornell, L., 1978, Renin and aldosterone levels in dolphins and sea lions (40117), Proc. Soc. Exp. Biol. Med., 157: 665–668. Orlov, M.V., Mukhlya, A.M., and Kulikov, N.A., 1988, Hormonal indices in the bottle-nosed dolphin Tursiops truncatus in the norm and in the dynamics of experimental stress, Sov. J. Evol. Biochem. Physiol., 24: 431–436. Ortiz, R.M., and Worthy, G.A.J., 2000, Effects of capture on plasma adrenal steroids and vasopressin levels in free-ranging bottlenose dolphins (Tursiops truncatus), Comp. Biochem. Physiol. A, 125: 317–324. Ortiz, R.M., Adams, S.H., Costa, D.P., and Ortiz, C. L., 1996, Plasma vasopressin levels and water conservation in fasting, postweaned northern elephant seals pups (Mirounga angustirostris), Mar. Mammal Sci., 12: 99–106. Ortiz, R.M., Worthy, G.A.J., and MacKenzie, D.S., 1998, Osmoregulation in wild and captive West Indian manatees (Trichechus manatus), Physiol. Zool., 71: 449–457. Ortiz, R.M., Wade, C.E., and Ortiz, C.L., 2000, Prolonged fasting increases the response of the reninangiotensin-aldosterone system, but not vasopressin levels, in postweaned northern elephant seal pups, Gen. Comp. Endocrinol., 119: 217–223. Ortiz, R.M., MacKenzie, D.S., and Worthy, G.A.J., 2000, Thyroid hormone concentrations in captive and free-ranging West Indian manatees (Trichechus manatus), J. Exp. Biol., 203: 3631–3637. Palmer, J.F., and Atkinson, S.A., 1998, Corticosteroid detection in cetacean blood and biopsy samples, Abstr., World Marine Mammal Science Conference, Monaco, 103. Patton, G.S., 1977, Aminogenic stimulation of insulin and glucagon release in Atlantic bottlenosed dolphins, Fed. Proc. Abstr., 36: 298. Patton, G.S., Orr, J.M., Gulli, R., and Searle, G.L., 1977, Glucose regulation in Atlantic bottlenosed dolphins, Diabetes, 26 (Suppl. 1): 411.
0839_frame_C10 Page 191 Tuesday, May 22, 2001 11:08 AM
Endocrinology
191
Pfeiffer, C.J., and Viers, V.S., 1995, Cardiac ultrastructure in the ringed seal, Phoca hispida, and harp seal, Phoca groenlandicus, Aquat. Mammals, 21: 109–119. Puig-Domingo, M., Guerrero, J.M., Reiter, R.J., Tannenbaum, M.J., Hurlbut, E.C., Gonzales-Brito, A., and Santana, C., 1988, Thyroxine 5′-deiodination in brown adipose tissue and pineal gland: Implications for thermogenic regulation and role of melatonin, Endocrinology, 123: 677–680. Raeside, J.I., and Ronald, K., 1981, Plasma concentrations of oestrone, progesterone and corticosteroids during late pregnancy and after parturition in the harbour seal, J. Reprod. Fertil., 61: 135–139. Renouf, D., and Brotea, G., 1991, Thyroid hormone concentrations in harbour seals (Phoca vitulina): No evidence of involvement in the moult, Comp. Biochem. Physiol. A, 99: 185–194. Renouf, D., and Noseworthy, E., 1991, Changes in food intake, mass, and fat accumulation in association with variations in thyroid hormone levels of harbour seals (Phoca vitulina), Can. J. Zool., 69: 2470–2479. Ridgway, S.H., and Patton, G.S., 1971, Dolphin thyroid: Some anatomical and physiological findings, Z. Vgl. Physiol., 71: 129–141. Ridgway, S.H., Simpson, J.G., Patton, G.S., and Gilmartin, W.G., 1970, Hematologic findings in certain small cetaceans, J. Am. Vet. Med. Assoc., 157: 566–575. Riviere, J.E., 1978, Molting in the harbor seal (Phoca vitulina) and its possible significance to exotic animal medicine, J. Zoo Anim. Med., 9: 46–52. Riviere, J.E., Engelhardt, F.R., and Solomon, J., 1977, The relationship of thyroxine and cortisol to the moult of the harbor seal Phoca vitulina, Gen. Comp. Endocrinol., 31: 398–401. Robin, E.D., Ensinck, J., Hance, A.J., Newman, A., Lewiston, N., Cornell, L., Davis, R.W., and Theodore, J., 1981, Glucoregulation and simulated diving in the harbor seal Phoca vitulina, Am. J. Physiol., 241: R293–R300. Ronald, K., and Thomson, C.A., 1981, Parturition and postpartum behaviour of a captive harbour seal, Phoca vitulina, Aquat. Mammals, 8: 79–90. St. Aubin, D.J., 1987, Stimulation of thyroid hormone secretion by thyrotropin in beluga whales, Delphinapterus leucas, Can. J. Vet. Res., 51: 409–412. St. Aubin, D.J., and Geraci, J.R., 1986, Adrenocortical function in pinniped hyponatremia, Mar. Mammal Sci., 2: 243–250. St. Aubin, D.J., and Geraci, J.R., 1988, Capture and handling stress suppresses circulating levels of thyroxine and triiodothyronine in beluga whales, Delphinapterus leucas, Physiol. Zool., 61: 170–175. St. Aubin, D.J., and Geraci, J.R., 1989, Seasonal variation in thyroid morphology and secretion in the white whale, Delphinapterus leucas, Can. J. Zool., 67: 263–267. St. Aubin, D.J., and Geraci, J.R., 1990, Adrenal responsiveness to stimulation by adrenocorticotropic hormone (ACTH) in captive beluga whales, Delphinapterus leucas, in Advances in Research on the Beluga Whale, Delphinapterus leucas, Smith, T.G., St. Aubin, D.J., and Geraci, J.R. (Eds.), Can. Bull. Fish. Aquat. Sci., 224: 149–157. St. Aubin, D.J., and Geraci, J.R., 1992, Thyroid hormone balance in beluga whales, Delphinapterus leucas: Dynamics after capture and influence of thyrotropin, Can. J. Vet. Res., 56: 1–5. St. Aubin, D.J., Smith, T.G., and Geraci, J.R., 1990, Seasonal epidermal molt in beluga whales, Delphinapterus leucas, Can. J. Zool., 68: 359–367. St. Aubin, D.J., Ridgway, S.H., Wells, R.S., and Rhinehart, H., 1996, Dolphin thyroid and adrenal hormones: Circulating levels in wild and semi-domesticated Tursiops truncatus, and influence of sex, age, and season, Mar. Mammal Sci., 12: 1–13. St. Aubin, D.J., De Guise, S., Richard, P., Smith, T.G., and Geraci, J.R., in press, Hematology and plasma chemistry as indicators of health and ecological status in beluga whales, Delphinapterus leucas, in Belugas and Narwhals: Application of New Technologies to Whale Science in the Arctic, Reeves, R.R., and St. Aubin, D.J. (Eds.), Arctic, 54. Sangalang, G.B., and Freeman, H.C., 1976, Steroids in the plasma of the gray seal, Halichoerus grypus, Gen. Comp. Endocrinol., 29: 419–422.
0839_frame_C10 Page 192 Tuesday, May 22, 2001 11:08 AM
192
CRC Handbook of Marine Mammal Medicine
Schumacher, U., Raugh, G., Plötz, J., and Welsch, U., 1992, Basic biochemical data on blood from antarctic Weddell seals (Leptonychotes weddellii): Ions, lipids, enzymes, serum proteins and thyroid hormones, Comp. Biochem. Physiol. A, 102: 449–451. Schumacher, U., Zahler, S., Heidemann, G., Skirnisson, K., and Welsch, U., 1993, Histological investigations on the thyroid glands of marine mammals and the possible implications of marine pollution, J. Wildl. Dis., 29: 103–108. Skog, E.B., and Folkow, L.P., 1994, Nasal heat and water exchange is not an effector mechanism for water balance regulation in grey seals, Acta Physiol. Scand., 151: 233–240. Sterling, K., Milch, P.O., and Ridgway, S.H., 1975, The day of the dolphin: Thyroid hormone metabolism in marine mammals, in Thyroid Hormone Metabolism, Harland, W.A., and Orr, J.S. (Eds.), Academic Press, London, 241–248. Stokkan, K.A., Vaughan, M.K., Reiter, R.J., Folkow, L.P., Martensson, P.E., Sager, G., Lydersen, C., and Blix, A.S., 1995, Pineal and thyroid functions in newborn seals, Gen. Comp. Endocrinol., 98: 321–331. Sucheston, M.E., and Cannon, M.S., 1980, Cortex of the suprarenal (adrenal) gland of Phoca vitulina richardi, Ohio J. Sci., 80: 140–144. Suzuki, M., Tobayama, T., Katsuma, E., Fujise, Y., Yoshioka, M., and Aida, K., 1998, Serum cortisol levels in 14 cetacean species, Abstr., World Marine Mammal Science Conference, Monaco, 131. Tagoe, C., Ayettey, S., Yates, R., and Gulland, F., 1998, Myocyte ultrastructural morphometry of the northern elephant seal (Mirounga angustirostris) and the harbor seal (Phoca vitulina), Abstr., Proceedings of the 29th Annual International Association for Aquatic Animal Medicine, 29: 106–107. Theodorou, J., and Atkinson, S., 1998, Monitoring total androgen concentrations in saliva from captive Hawaiian monk seals (Monachus schauinslandi), Mar. Mammal Sci., 14: 304–310. Thomson, C.A., and Geraci, J.R., 1986, Cortisol, aldosterone, and leucocytes in the stress response of bottlenose dolphins, Tursiops truncatus, Can. J. Fish. Aquat. Sci., 43: 1010–1016. Vivien-Roels, B., and Pévet, P., 1983, The pineal gland and the synchronization of reproductive cycles with variations of the environmental climatic conditions, with special reference to temperature, Pineal Res. Rev., 1: 91–143. Williams, T.D., Rebar, A.H., Teclaw, R.F., and Yoos, P.E., 1992, Influence of age, sex, capture technique, and restraint on hematologic measurements and serum chemistries of wild California sea otters, Vet. Clin. Pathol., 21: 106–110. Wilson, J.D., Foster, D.W., Kronenberg, H.M., and Larsen, P.R. (Eds.), 1998, Williams Textbook of Endocrinology, W.B. Saunders, Philadelphia, 1819. Woldstad, S., and Jenssen, B.M., 1999, Thyroid hormones in grey seal pups (Halichoerus grypus), Comp. Biochem. Physiol. A, 122: 157–162. Young, B.A., and Harrison, R.J., 1970, Ultrastructure of the dolphin adenohypophysis, Z. Zellforsch., 103: 475–482. Zenteno-Savin, T., and Castellini, M.A., 1998a, Changes in the plasma levels of vasoactive hormones during apnea in seals, Comp. Biochem. Physiol. C, 119: 7–12. Zenteno-Savin, T., and Castellini, M.A., 1998b, Plasma angiotensin II, arginine vasopressin and atrial natriuretic peptide in free ranging and captive seals and sea lions, Comp. Biochem. Physiol. C, 119: 1–6.
0839_frame_C11 Page 193 Tuesday, May 22, 2001 11:09 AM
11 Reproduction Todd R. Robeck, Shannon K. C. Atkinson, and Fiona Brook
Introduction The reproductive physiology of marine mammals is an extremely diverse topic; yet the small amount of information that has been collected has come from only a few species of cetaceans and pinnipeds. As a result, generalizations are made concerning the reproductive function of entire families based on information obtained from these few species. These generalizations must be interpreted with caution, as important differences exist among species within each family. In addition, this chapter focuses on reproductive aspects of species most likely to be encountered by veterinarians working with animals in captivity. This chapter assumes the reader has a basic knowledge of the physiology of mammalian reproduction. Reviews by Harrison and Ridgway (1971), Richkind and Ridgway (1975), Hill and Gilmartin (1977), Kirby (1982), Sawyer-Steffan et al. (1983), Kirby and Ridgway (1984), Schroeder and Keller (1989; 1990), and Schroeder (1990a,b), documented work with bottlenose dolphins (Tursiops truncatus). Perrin et al. (1984) reviewed cetacean reproduction. For pinnipeds, Riedman (1990) provided useful tables on reproductive timing and maternal care, and a review of reproduction by Atkinson (1997) focused primarily on phocids. Most recently, Boyd et al. (1999) reviewed reproductive physiology, timing of reproduction, and different lifehistory strategies for pinnipeds, sirenians, and cetaceans.
Physiology of Reproduction Although the reproductive function of mammals varies among species, the hormones involved and their general functions tend to be conserved across the mammalian class. A general review of the control of reproduction, with emphasis on the estrous cycle, will give the reader a foundation on which other reproductive processes can be discussed in both the male and female. If a more detailed understanding of the physiology of these processes is desired, there are a number of good reference books available (Knobil and Neill, 1988; Cupps, 1991; Youngquist, 1997). Mammalian reproduction is regulated by a series of neurological and hormonal feedback mechanisms involving the hypothalamus, pituitary, and gonads. These three organs are commonly referred to as the hypothalamic–pituitary–gonadal axis (see Chapter 10, Endocrinology). The effects that photoperiod and other environmental stimuli have on reproductive events
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
193
0839_frame_C11 Page 194 Tuesday, May 22, 2001 11:09 AM
194
CRC Handbook of Marine Mammal Medicine
provide evidence that neurological transduction of these stimuli in the brain leads to control of reproductive events. Most of this transduction appears to occur in the hypothalamus and associated nuclei where neurons originate that secrete hypophysiotropic hormones into the hypophyseal portal system. These hormones control the anterior pituitary gland. Gonadotropin-releasing hormone (GnRH) is one of these hormones and is of primary importance in regulating reproductive endocrinological events. GnRH receptor binding in the anterior pituitary causes luteinizing hormone (LH) and follicle-stimulating hormone (FSH) to be released into circulation. GnRH secretion is important for reproductive control and is pulsatile in nature. Secretion of GnRH is mediated by a pulse generator located in the mediobasal hypothalamus. The episodic generation of GnRH translates into a subsequent pulsatile release of LH and FSH from the anterior pituitary. The significance of the episodic secretion is apparent when comparing the effects of exogenous GnRH delivered as a constant infusion or as a pulse infusion (Ganong, 1991). GnRH receptors in the anterior pituitary rapidly downregulate in both numbers and sensitivity when exposed to continual GnRH input and upregulate when GnRH concentration is low. Thus, constant GnRH infusion first stimulates LH release; then, as receptor sensitivity decreases, GnRH will inhibit LH release (Nett et al., 1981; Conn et al., 1988; Blue et al., 1991). This response is the basis for the use of GnRH agonist as contraception agents and will be discussed further below. Control of GnRH release is mediated by neurological input and feedback from gonadal hormones. Feedback appears to have direct effects on the pulse generator by causing changes in the amplitude and frequency of GnRH release. The basic model for this control is based on primate research, but the control appears to be similar in most mammalian species. During the early follicular phase of the estrous cycle, FSH production is slightly elevated. This increase in FSH production results in follicular recruitment and growth and causes an increase in LH receptor concentrations in the follicle(s) (Brown et al., 1986). Estrogen has also been positively correlated with numbers of LH receptors in the preovulatory follicle. As the follicles continue to expand or grow, estrogen is produced through paracrine interactions between thecal and granulosa cells that line the follicle. Increased estrogen production initially inhibits both FSH and LH secretion from the pituitary. As the follicle(s) approaches preovulatory stage, estrogens reaching maximal production (the preovulatory estrogen surge) exert a positive effect on frequency and amplitude of GnRH secretion resulting in the preovulatory LH surge. LH causes the follicle to produce a small two-subunit glycoprotein, called inhibin. Inhibin not only suppresses FSH production, but increases thecal cell sensitivity to LH in the preovulatory follicle (Baird and Smith, 1993). This combination of increased LH receptors and increased sensitivity to LH ensures an adequate response to the LH surge and ovulation. Once ovulation occurs, granulosa and thecal cells are converted to progesterone-secreting large and small luteal cells, respectively (Hendricks, 1991). These morphologically different luteal cells appear to have different functions in the corpus luteum (CL) and have been shown to have different secretory capacities. The luteal cells of the recently ruptured follicle rapidly organize into the CL. Progesterone, and to a smaller extent estrogen, produced by the CL inhibit LH and FSH secretion by decreasing the frequency of GnRH release from the hypothalamus. If the cycle is nonfertile, the uterus releases a series of five to eight pulses of prostaglandin F2α , which, in turn, result in luteal regression. The release of prostaglandin, at least in ruminants, appears to be initiated by pulsatile oxytocin release from the neurohypophysis, encouraged by release of oxytocin from the CL, and a concomitant decrease in circulating progesterone and estrogen (Silvia et al., 1991). The decrease in progesterone and estrogen allows the GnRH pulse generator once again to increase in frequency and amplitude, resulting in FSH and LH secretion and initiating folliculogenesis of the next cycle.
0839_frame_C11 Page 195 Tuesday, May 22, 2001 11:09 AM
Reproduction
195
The pineal gland influences reproductive function by transducing photoperiodic messages to chemical messages through innervation in the superior ganglia (Lindsay, 1991). In response to changes in photoperiod, the pineal gland releases melatonin. The increase in melatonin appears to inhibit reproduction by affecting the pulsatile release of LH. Melatonin is synthesized only during dark hours, and its production can be inhibited by nocturnal exposure to artificial light. Prolactin may play an important role in regulating seasonality, but this remains to be determined.
Pinniped Reproduction Pinniped reproduction has recently been reviewed by Atkinson (1997) and Boyd et al. (1999). This chapter summarizes basic reproductive physiology and focuses on clinically significant parameters. The tremendous variability that exists among the three pinniped families (phocidae, otariidae, and odobenidae) and the lack of information on their reproductive physiology preclude any detailed discussion concerning any one species. Instead, gross generalizations have often been made out of necessity. Research with harbor seals (Phoca vitulina) will often be used as an example of normal phocid reproductive parameters. As this approach may be misleading, any reader who truly wants a deeper appreciation of a particular species is advised to use this chapter as a beginning, or foundation, for further inquiry.
Female Pinniped Reproduction Reproductive Cycle
For this chapter, the reproductive cycle is defined as the period during which all major components of reproduction are experienced. These components arbitrarily begin with a fertile estrous period (which includes estrus, ovulation, and conception), followed by gestation, lactation, anestrus, and back to a fertile estrous cycle. As most marine mammals have some seasonal component to their reproductive events, and as seasonality has a direct impact on when a fertile estrus can occur, seasonality of reproductive events is included in this discussion. The reproductive cycle of pinnipeds is dominated by three basic phases: estrus, embryonic diapause, and fetal growth and development (Boyd et al., 1999). Embryonic diapause, or delayed implantation, was recognized in pinnipeds as early as 1940 (Harrison, 1968). Pinnipeds are classified as having obligate embryonic diapause (Renfree and Calaby, 1981). The time when the embryo resumes cellular divisions is a critical point during embryonic development of the fetus and, in nonpregnant females, is a period of reactivation of sexual activity. Understanding this phenomenon is important when attempting to diagnose pregnancy in these species. It appears that most if not all pinnipeds have either postpartum or postlactational estrus periods. Otariids generally have a postpartum estrus 6 to 12 days after birth. California sea lions (Zalophus californianus), however, appear to be exceptions among otariids in that their estrous period is approximately 1 month after birth (Heath, 1985). In phocids, estrus begins toward the end of lactation, or after weaning (Riedman, 1990; Atkinson, 1997). Harbor seal lactation can last 21 to 42 days, with estrus occurring after that time (Bigg, 1969; 1973). Estrus can last from 1 to 9 weeks, with some animals being induced ovulators. In walrus (Odobenus rosmarus), an approximate 4-month postpartum estrus occurs in late summer; however, conception cannot occur because males are infertile at this time. The females have a second midlactational estrus approximately 6 months later, around February, during peak male fertility (Fay et al., 1981; Riedman, 1990). Thus, walrus are polyestrous, but functionally monoestrous. The potential fertility of the late-summer postpartum estrus is unknown. For a summary of reproductive events in pinnipeds, see Table 1.
0839_frame_C11 Page 196 Tuesday, May 22, 2001 11:09 AM
196
CRC Handbook of Marine Mammal Medicine
TABLE 1 Reproductive Characteristics of Three Species of Pinnipeds Species Reproductive Characteristics
Harbor Seal a (Phoca vitulina) d
California Sea Lion b (Zalophus californianus)
Walrus c (Odobenus rosmarus) Mid-April to mid-June, peak May Midlactation
Pupping period
Early May
Late May, early June
Timing of ovulation
End of lactation
Conception Duration of lactation Delayed implantation period Delayed implantation Postimplantation gestation Total gestation interval
June 21–42 days July–Aug.
Approx. 28 days postpartum Late June, early July 6–12 months July–Sept.
1.5–3 months Sept.–May
3 months Oct.–May
4–5 months Aug.–May
11 months
11 months
15 months
Jan.–March (peak Feb.) 24+ months March–July
a
Sources: Bigg, 1969; Bigg, 1973; Gardiner et al., 1999; Odell, pers. comm. Sources: Odell, 1981; Odell, pers. comm. c Sources: Fay et al., 1981; Fay, 1981. d These pupping data are based on U.S. captive animal observations. Time intervals are consistent, but timing of the reproductive cycle varies with latitude. For example, postimplantation gestation lasts 8.5 months for all harbor seals, but on the West Coast of North America, pups are born in February in Mexico and in July in Alaska. b
Estrous Cycle
The onset of the estrous cycle of pinnipeds is closely tied to the annual reproductive cycle (or biennual in the case of the walrus). Available data suggest that otariids and phocids are monestrous, spontaneous ovulators, and if pregnancy does not occur, they do not have a second estrous period until the following year. The known exception to this generality is the Hawaiian monk seal (Monachus schauinslandi), which has been shown to exhibit polyestrous activity (Iwasa et al., 1997; Iwasa and Atkinson, 1997). This may be a result of the animal’s subtropical environment and lesser dependence on a well-defined annual reproductive cycle than other species, or a reflection of true reproductive potential of the phocids (Atkinson and Gilmartin, 1992; Pietraszek and Atkinson, 1994). It can be assumed that the onset of parturition and the subsequent diminishing levels of circulating progesterone are the triggers that cause the pinniped hypothalamus to begin increased GnRH secretion, LH and FSH release, and to initiate follicular recruitment and development. In northern fur seals (Callorhinus ursinus), follicular recruitment begins in February in the nongravid ovary, before parturition and ovulation in July (Craig, 1964). In phocids, follicular recruitment starts in close proximity to parturition, resulting in mid- to late-lactational ovulation. This difference may reflect differences between phocids and otariids in the functional life span of the CL. In otariids, the CL begins regressing, and blood progesterone levels begin to fall in February, coincident with follicular recrudescence in the opposite ovary (Kiyota et al., 1999). In seals, the CL regresses and circulating progesterone declines rapidly after parturition (Boyd, 1983; Iwasa et al., 1997). In most phocids and otariids, follicular growth and ovulation occur on alternating ovaries during subsequent pregnancies. It appears that the presence of a CL inhibits follicular activity on the ipsilateral ovary to a greater degree than the contralateral ovary due to some local or paracrine effect (Craig, 1964; Amoroso et al., 1965; Boyd, 1983).
0839_frame_C11 Page 197 Tuesday, May 22, 2001 11:09 AM
Reproduction
197
Follicular maturation continues either during late pregnancy or lactation, and results in a rapid rise in estrogen production, presumptive LH surge, and ovulation. This rise in estrogen that has been observed in northern fur seals lasts less than 5 days and reaches circulating concentrations greater than 30 pg/ml (Kiyota et al., 1999). Estrus in Hawaiian monk seals lasts 2 to 6 days (Atkinson et al., 1994; Pietraszek and Atkinson, 1994). In the northern fur seal, multiple follicular recruitment results in approximately four Graffian follicles greater than 10 mm in diameter around parturition. From this group of follicles, one is selected and ovulates 3 to 5 days after parturition (Craig, 1964). Pregnancy and Pseudopregnancy
Pregnancy in pinnipeds can be divided into five distinctly important events: (1) conception, (2) embryonic diapause, (3) embryo reactivation and implantation, (4) fetal development, and (5) parturition. In otariids, it appears that an obligate pseudopregnancy ensues after ovulation, regardless of the presence of a normal blastocyst (Boyd, 1991; Atkinson, 1997). However, after the 4 months physiologically allotted for embryonic diapause, uterine development and placental formation can only occur if a functional blastocyst is present. The specific period during embryonic diapause or gestation when maternal recognition of pregnancy (MRP) occurs is unknown. After implantation, and during the latter part of gestation, it is believed that placental gonadotropin, acting via fetal gonads, results in placental production of estrogens and progesterone. This fetal–placental unit is then believed to be responsible for the maintenance of pregnancy, and for triggering parturition. Fetal production of adrenal or gonadal hormones results in hypertrophy of these organs, which are similar in size at birth to adult organs, but rapidly regress in size until puberty. Despite circumstantial evidence for the importance of placental steroid production, some conflicting data recently have been obtained. A complete monitoring of pinniped serum progesterone and estrogen was done by Kiyota et al. (1999) on four northern fur seals during 2 consecutive years. They observed an initial rise in progesterone to 20 to 30 ng/ml in July, indicating ovulation. Progesterone concentrations dropped to 5 to 10 ng/ml during embryonic diapause from August through October, and increased again in November to 25 to 35 ng/ml (similar to observations in wild fur seals; Daniel, 1974). This pattern was observed in seven cycles, but only two resulted in pregnancy. One of the cycles had an initial progesterone spike of around 8 ng/ml that rapidly dropped to slightly over 1 ng/ml. The five hormonal profiles in nonpregnant fur seals studied by Kiyota et al. (1999) that appeared similar to the two hormonal profiles of pregnant animals provide evidence that otariids exhibit an obligatory pseudopregnancy beyond the period of normal implantation. That is, maintenance of the CL is not dependent upon maternal recognition of pregnancy or an embryonic product. The presence of circulating progesterone also contradicts Laws’ (1955) assumption that the CL was nonfunctional in late gestation. In contrast, endocrine data from the harbor seal show evidence for pseudopregnancy that only lasts through diapause, with blood progesterone levels declining rapidly after the window of implantation has occurred in nonpregnant seals (Reijnders, 1991; Atkinson, 1997). High circulating levels of progesterone (greater than 3 ng/ml) have been observed for long periods of time in nonpregnant captive walrus. However, no serial sampling has been done to define the duration of this pseudopregnancy. Placental gonadotrophins isolated by Hobson and Boyd (1984) also appear to be required for CL function in some species. Since pinnipeds are an extremely diverse group of animals, it would be safe to assume that extreme species variations can occur, and fetal–placental or maternal control of luteal function should be addressed on a species-by-species basis.
0839_frame_C11 Page 198 Tuesday, May 22, 2001 11:09 AM
198
CRC Handbook of Marine Mammal Medicine
Embryonic Diapause and Reactivation
Embryonic diapause in pinnipeds appears to be regulated maternally. During early postconception, the embryo divides at a normal (compared with mammals without diapause) rate until the blastocyst stage around day 5 to 8. At this point, cellular divisions, as determined by a mitotic index, decline rapidly to a point where the embryo doubles in cell numbers every 50 to 60 days (Daniel, 1971). The embryo remains in this slow period of growth for 2 to 4 months (species dependent) until it is reactivated by maternal physiology. During this slow-growth period, the embryo remains in its zona pellucida and does not hatch until after reactivation (Harrison, 1968). Reactivation of the blastocyst appears to be controlled by photoperiod, with most animals implanting during a decreasing photoperiod. Water temperature and nutritional availability may also be important factors regulating pinniped reproductive cycles (Atkinson, 1997). Research into photoperiodic control of reproduction in other mammals has found that an animal does not have to be exposed to a continual light/dark cycle, but has windows of receptivity when exposure to light or dark can define the endocrine response. Thus, exposure to a 1-hour “pulse” of light during the receptive period approximately 9.5 hours after the onset of darkness can be enough to induce early reactivation of reproductive activity in the mare (Sharp et al., 1997). In the same manner, it has been postulated that pinniped parturition and blastocyst reactivation are controlled by the date they are exposed to a particular length of day, generally during a decrease in day length (Temte, 1991). This time appears to vary slightly with each species, but generally occurs around the autumn equinox, when the day length is 12 hours. Recent research in harbor seals demonstrated a significant decrease in pituitary sensitivity to LH during winter and spring (Gardiner et al., 1999). This decreased sensititivity to LH is consistent with animals whose reproduction is under photoperiodic control. During reactivation of the blastocyst, the quantity and molecular weight of uterine protein secretions increase. The increase in uterine protein secretion corresponds to an eight-to ninefold increase in blastocyst mitotic activity, with cell number doubling every 12 hours (Harrison, 1968). A protein, possibly related to blastokinin, is believed to be responsible for blastocyst reactivation. Concurrent with uterine protein secretion, and possibly regulated by photoperiod, progesterone and estrogen increase dramatically. The estrogen increase has been described as a “surge,” and may represent follicular activity on the ovaries prior to reactivation (Temte, 1985). These follicles quickly become atretic after implantation, but the estrogen surge may prime the pituitary to secrete more LH, causing the luteotrophic effects required for CL stimulation and the resulting increase in progesterone secretion. The estrogen increase may also be required to increase uterine progesterone receptors, thus increasing sensitivity to progesterone and ensuring the proper endometrial response to progesterone. Progesterone causes the uterus to prepare for approaching implantation. In harbor seals in the United Kingdom, implantation occurs in November and, by December, placentation has been established. The exact timing varies with latitude. Implantation
Although the preimplantation estrogen surge was observed in harbor seals by earlier researchers, until recently, it had not been observed in other pinniped species. This lack of duplicative research left many questioning its existence. However, Kiyota et al. (1999) observed an estrogen “surge” in northern fur seals associated with implantation in November. Unfortunately, they did not clarify if the surge was documented in animals that were pregnant, or nonpregnant, or both. Since the surge was not observed in all of the animals sampled, they felt that their sampling frequency of every 5 days was insufficient to determine whether it was inconsistently observed, or whether they just missed it.
0839_frame_C11 Page 199 Tuesday, May 22, 2001 11:09 AM
Reproduction
199
FIGURE 1 An approximately 8-month-old Odobenus rosmarus fetus. The dotted line represents a biparital measurement of 6.1 cm. Fetus is estimated between 90 and 120 days post-reactivation. (From T. Robeck, unpubl. data.)
As fetal development proceeds, and in support of placental–fetal maintenance of pregnancy, progesterone secretion declines slightly until parturition occurs. Fetal gonads hypertrophy and are believed to be responsible for secretion of important steroid precursors that are converted to estrogens by the placenta. In addition, placental chorionic gonadotropin (CG) production is believed to be essential for CL maintenance. It was hypothesized that nonpregnant pinnipeds have an obligate pseudopregnancy interval equal to the period during gestation prior to placental gonadotropin production. CLs in pregnant animals will have a third surge (the second surge occurs at implantation) of luteal activity in response to placental CG production, resulting in continued production of progesterone until parturition. However, CG concentrations in the placenta are extremely low when compared with other species that rely on CG for luteal maintenance (Hobson and Boyd, 1984; Hobson and Wide, 1986). Recent research in northern fur seals demonstrates similar hormonal profiles in pregnant and pseudopregnant animals (Kiyota et al., 1999), while research on harbor seals continues to provide support for this theory of extra-hypophyseal or placental support of the CL (Hobson and Boyd, 1984; Reijnders, 1991; Gardiner et al., 1999). These differences demonstrate the lack of understanding of mechanisms involved in maintenance of pregnancy in pinnipeds and the differences between phocids and otariids. Pregnancy Diagnosis
Ultrasound diagnosis of pregnancy has been used successfully in mid- to late-gestation in a variety of pinnipeds, but will not detect the fetus during embryonic diapause. Elevated progesterone levels are a useful indication of pregnancy, although values have only been published for a limited number of species. Levels also may be elevated during pseudopregnancy and in nonpregnant animals (levels greater than 3 ng/ml were observed in a nonpregnant captive walrus). The practitioner is advised to use a combination of high progesterone and ultrasound to detect pregnancy (i.e., after embryonic reactivation) (Figure 1). Induction of Parturition
Cloprostenol, a synthetic prostaglandin F2α (Estrumate®, Mobay, Shawnee, KS), has been used on California sea lions with full-term fetuses to induce abortion (Gulland, 2000). Five animals that had been exposed to domoic acid (with dead fetuses) were given 500 mg cloprostenol intramuscularly (IM) and delivered fetuses 36 to 40 hours later.
0839_frame_C11 Page 200 Tuesday, May 22, 2001 11:09 AM
200
CRC Handbook of Marine Mammal Medicine
Lactation
As with most eutherian mammals, prolactin and oxytocin appear to be crucial hormones for regulating lactation. No single prolactin-releasing hormone has been identified, but a number of neuropeptides in the hypothalamus, including vasoactive intestinal polypeptide, thyrotropinreleasing hormone (TRH), and prolactin-releasing factor (perhaps identical to TRH), may all function in this capacity (Norman and Litwack, 1987; Ganong, 1991). Prolactin secretion is increased by neurogenic stimulation via suckling. Prolactin appears essential for mammary gland secretory cell development, and increases in otariids 1 to 2 days prior to parturition, peaking 0 to 3 days postpartum (Boyd, 1991). However, unlike in mink (Mustela vison), in which a preimplantation rise in prolactin is believed to be involved with reactivation, prolactin concentrations decreased to undetectable levels toward the end of lactation and embryonic diapause (Boyd, 1991). In carnivores, prolactin has been identified as playing a role in the development of the CL. In otariids, it appears that prolactin may play a role in both ovulation and CL formation; however, additional data are needed to determine this. Oxytocin (synergistically with prolactin or somatotropin and cortisol) is believed to be essential for the maintenance of lactation. This hormone is secreted via the neurohypophysis in response to suckling stimuli. Once released, oxytocin is important for milk letdown. During this process, myoepithelial cells surrounding the alveoli contract, forcing milk out of the glands. In addition, oxytocin causes relaxation of smooth muscles surrounding the ducts and teat cisterns, resulting in space for milk ejected from the alveoli. Thus, suckling animals only have to overcome the teat sphincter resistance to nurse effectively (Baldwin and Miller, 1991). Generally, continued suckling stimulation, and subsequent oxytocin release, is required to maintain milk production. Indeed, phocid females, whose lactations last from 4 to 60 days, will spend almost the entire time with the pup during this period, with short or no intervals for feeding. However, otariids whose lactation period lasts 4 to 12 months will often leave the pup for feeding from 1 to 8 days. Thus, continual suckling is not required for maintenance of lactation in otariids. During periods of low or no stimuli, milk production slows down or stops; however, under the influence of prolactin, mammary glands do not involute. Once suckling recurs, milk is let down, most likely via oxytocin secretion, and milk production increases or is reinitiated (Boyd, 1991). Milk Collection
Oxytocin has been used on a variety of pinnipeds to enhance collection of milk samples for research purposes. Intramuscular injection of 20 Posterior Pituitary Units (USP) of oxytocin will facilitate collection of milk by stimulating milk let down from the teat. Unlike many species of cetacea, pinnipeds do not have a lactational or suckling suppression of estrus. In fact, all pinniped species undergo estrus toward the end of, or during, lactation.
Male Pinniped Reproduction Anatomy
The reproductive anatomy of male pinnipeds varies with the family. Phocids and odobenids have para-abdominal testes that lie below the blubber layer adjacent to the abdominal musculature, while the otariids have scrotal testes. Some otariids are seasonally scrotal; that is, their testes descend into the scrotum only during the breeding season. Testis size in all marine mammals is proportional to body mass and, in most cases, body length (Kenagy and Trombulak, 1986). Testis size is also related to the mating system and/or the length of the breeding season, with relatively large testes in species that have high rates of copulatory activity and associated high rates of spermatogenesis.
0839_frame_C11 Page 201 Tuesday, May 22, 2001 11:09 AM
Reproduction
201
Animals with scrotal testes are able to lower and raise the testes using the cremaster and dartos muscles. This scrotal agility protects the sperm from cold shock of the surrounding aquatic environment as well as physically protecting the testes when the animal is moving on land. In ascrotal species, protection of the testes and developing spermatocytes from hyperthermia is accomplished through a direct vascular heat-exchange mechanism using arteriovenous anastomoses. The anatomy of the arteriovenous anastomoses allows cool blood from the skin and flippers to flow directly to the testicular artery, preventing hyperthermic insult to the developing sperm (Rommel et al., 1995) (see Chapter 9, Anatomy). All of the pinnipeds, polar bears (Ursus maritimus), and sea otters (Enhydra lutris) have bacula, or penis bones. The distal end of the baculum is morphologically variable and differs substantially among species (Morejohn, 1975). Most of the phocids are aquatic copulators with relatively large bacula, which may function either to prevent water damage to sperm cells after ejaculation or to increase sperm competition in species where the female mates with more than one male (Miller et al., 1999). Most otariids are of large body size and are terrestrial copulators; bacula in otariids are relatively small; bacular fractures have been reported in otariids. Most of the growth in bacular length is achieved by puberty; however, bacular mass and density continue to increase for another decade. Sexual Maturity
Sexual maturity in male pinnipeds tends to occur at 2 to 7 years of age (Atkinson, 1997; Boyd et al., 1999). Diagnostic measures of puberty are the relative weight of the testes, an increase in the circulating concentrations of testosterone, and active spermatogenesis. Bacular mass and length also increase during puberty. Testosterone concentrations have been measured in many pinnipeds (Noonan et al., 1991; Atkinson and Gilmartin, 1992), and in all species the concentrations increase around the time of sexual maturity. Histological evidence of sexual maturity can be measured in the diameter of the seminiferous tubules, proportion of the tubules to interstitium, and the presence, abundance, and maturation of spermatocytes in the tubules. Although the age of puberty may occur early in life, many pinnipeds are not behaviorally capable of breeding until 8 to 10 years of age (Atkinson, 1997). In sexually dimorphic species, the secondary sexual characteristics generally become obvious during and after puberty. Examples in pinnipeds include increased body size, a developed sagittal crest, elongated proboscis or hood, calloused chest shield, development of a musky odor, and/or more or elongated guard hairs on the neck and shoulders. In some species, the secondary sexual characteristics are only fully developed in males that are both physiologically and behaviorally mature. Seasonality
Pinnipeds are seasonally fertile, with the length of the fertile season greatest in tropical animals and shortest in temperate animals (Atkinson and Gilmartin, 1992). Seasonality is associated with increased size of the testes and accessory reproductive glands, increased testicular and circulating testosterone concentrations, and spermatogenesis during the breeding season (Griffiths, 1984a,b). Increased size and mass of the testes are due to increased diameter of the seminiferous tubules and the epididymis. Decreases in testicular tissue and associated glands during the nonbreeding season are thought to be due to the shrinkage of the anterior pituitary cells that produce gonadotropins. This theory has recently been supported by the significant seasonal decrease in pituitary release of LH in response to an exogenous GnRH challenge (Gardiner et al., 1999). The lack of gonadotrophic support via decreased LH and FSH release from the pituitary leads to seasonal atrophy of the testes, and testosterone concentrations decline to baseline during the nonbreeding season (Frick et al., 1977; Gardiner et al., 1999).
0839_frame_C11 Page 202 Tuesday, May 22, 2001 11:09 AM
202
CRC Handbook of Marine Mammal Medicine
Mature sperm in both the seminiferous tubules and epididymis and elevated testosterone concentrations are apparent preceding the breeding period in several species of pinnipeds. Spermatogenesis usually lags behind testosterone production by 1 to 3 months, as production of testosterone by testicular Leydig cells is necessary for germ-cell differentiation in the seminiferous tubules. During seasonal quiescence, spermatogenesis ceases. In addition, at least in gray seals (Halichoerus grypus), the seminiferous tubules undergo involution, resulting in a decrease in both testicular dimension and mass (Griffiths, 1984a).
Contraception and Control of Aggression A common concern in facilities housing marine mammals is the control of fertility in captive animals. Three particular species for which fertility control has become a concern are bottlenose dolphin, California sea lion, and harbor seal. All of these species can be prolific breeders in the captive setting. The most common methods of reducing fertility have been physical separation, castration of males, and contraception of female animals. Sexual behaviors are often associated with territorial or aggressive behaviors. The need to control behavior is obvious in the captive setting. It also is important in the management of declining species in which male aggression inhibits the recovery of the species. Sexual behaviors may be as obvious as approaching, chasing, and nudging of females, vocalizations, and agonistic threats to neighboring males. Many intraspecific acts of aggression indicate a form of dominance. In several pinniped species, territorial and/or aggressive behaviors occur when testosterone concentrations are increasing, suggesting a behavioral role for the elevated hormone concentrations (Atkinson and Gilmartin, 1992; Theodorou and Atkinson, 1998). Increased testosterone concentrations usually coincide with the seasonal approach of the breeding season. In many species, the ability of an adult male to maintain rank and access to estrus females correlates with age and territorial behavior. Females
For female pinnipeds, the majority of research, sparse as it may be, has been conducted on phocids. Research has focused on the use of porcine zona pellucida vaccine (PCP). PCP vaccine uses sperm-binding sites on the porcine zona pellucida as a source of antigen. Thus, the vaccine causes an autoimmune antibody response directed against recently ovulated ova that blocks sperm binding. Without sperm binding, the degranulation reaction cannot occur and sperm are unable to penetrate the zona to fertilize the ova. This vaccine has been effectively applied to a number of captive and wild hoofstock (Kirkpatrick et al., 1982; 1990; 1996). Its practical application to wild pinniped populations was hindered because of the requirement for up to four booster vaccinations. Recent improvements to the delivery system, however, have resulted in effective contraception after single-dose administration in wild seals (Brown et al., 1997a,b). Although this vaccine may have its use in captive populations, and a large body of evidence suggests that in some species it may not be reversible, and it has been associated with negative ovarian and systemic inflammatory side effects in canids and felids (Mahi-Brown et al., 1988; Asa, 2000). Males
Castration has been used routinely to prevent breeding of captive harbor seals and captive California sea lions. Recently, GnRH agonists have been applied to male marine mammals in an effort to reduce fertility and control aggression (Atkinson et al., 1993; 1998; Briggs, 2000). In the male, episodic pulses of GnRH occur at regular, species-specific frequencies (Sisk and Desjardins, 1986) concurrent with cyclic changes in GnRH secretion frequency and amplitude observed in females (Ganong, 1991; Mariana et al., 1991). The periodicity of the pulse rate of GnRH secretion is
0839_frame_C11 Page 203 Tuesday, May 22, 2001 11:09 AM
Reproduction
203
important for normal reproductive function. This is evident when comparisons are made between steady infusions or pulsatile infusions of GnRH. Since GnRH regulates its own receptor production at the pituitary, receptor production is high when ligand is low and low when ligand is high, and receptor changes can occur rapidly. Constant infusions of GnRH result in constant downregulation of receptors (Conn et al., 1987). Thus, when GnRH is administered in a constant fashion, there is an initial dramatic increase in LH secretion from the pituitary, and subsequent LH secretion becomes refractory to GnRH as the pituitary receptors for GnRH are reduced (Sundaram et al., 1982; Mann et al., 1984; Schurmayer et al., 1984). In addition to the initial post-GnRH agonist administration surge of LH, a temporally associated testosterone increase is also observed (Belanger Anclair et al., 1980). After 3 to 4 days of constant infusion of GnRH agonist, basal levels of testosterone can double, declining to baseline, or less than baseline, as the pituitary becomes desensitized to GnRH around day 10 (Chrisp and Goa, 1990). Depression of testosterone synthesis and secretion beyond day 10 requires continued, steady administration of the agonist. When administered to Hawaiian monk seals, GnRH agonists (D-Trp-6-LHRH and D-Ala-6LHRH) have reduced circulating testosterone concentrations to castrate levels by approximately 2 weeks after injection, with results lasting approximately 2 months (Atkinson et al., 1993; 1998). As predicted, reduction in circulating testosterone concentrations was preceded by a dramatic elevation in testosterone concentrations; however, LH concentrations have never been measured to evaluate exactly when the pituitary becomes refractive (Atkinson, unpubl. data). Doses of 2.5 to 11.25 mg of the GnRH agonist incorporated into microlatex beads were administered to Hawaiian monk seals, with similar results after all doses. Harbor seals and northern elephant seals (Mirounga angustirostris) exhibited similar responses; however, the northern elephant seals required 40 mg to produce a discernible effect on testosterone concentrations (Atkinson, Yochem, and Stewart, unpubl. data). The effects of GnRH agonists on fertility have been demonstrated in two facilities that house harbor seals. After annual treatment of males, no offspring have occurred.
Reproductive Abnormalities in Pinnipeds Very little information is available concerning pathological conditions of reproductive events. Reijnders (1986) showed reduced reproductive rates in harbor seals fed fish from polluted waters, and Gilmartin et al. (1976) demonstrated an association between maternal and fetal concentrations of pesticides and premature births in California sea lions (see Chapter 22, Toxicology). High tissue concentrations of polychlorinated biphenyls and reproductive tract abnormalities including uterine stenosis have been described in gray, harbor, and ringed (Phoca hispida) seals (see Chapter 22, Toxicology). The mechanisms for these changes are unknown, but pregnancy rates of seals in the Gulf of Bothnia decreased from a normal of 60 to 90%, to as low as 25% (Boyd et al., 1999). Rates of dystocia in captive-bred animals have not been determined; however, they appear to be low since no cases have been reported. Stillbirths occur infrequently, with no data available on causes or incidence of occurrence. In a few species of pinnipeds, mobbing behavior is observed, in which groups of males attempt a mass mating, typically with an adult female or juvenile seal of either sex. In Hawaiian monk seals, the behavior is primarily targeted at female seals that are periovulatory, and is concurrent with a seasonal rise in testosterone concentrations (Atkinson et al., 1994). In northern elephant seals, the females are thought to submit to the mobbing behavior as they leave the territory of the dominant male, returning to the sea. These behaviors have yet to be documented in captive animals; however, the species in which the behaviors have been demonstrated are not commonly maintained in captivity.
0839_frame_C11 Page 204 Tuesday, May 22, 2001 11:09 AM
204
CRC Handbook of Marine Mammal Medicine
Cetacean Reproduction The majority of cetaceans housed in zoological settings can be divided into two different taxonomic families: Delphinidae and Monodontidae. The most commonly displayed Delphinidae include the bottlenose dolphin, the killer whale (Orcinus orca), the white-sided dolphins (Lagenorhynchus obliquidens and L. acutus), and the false killer whale (Pseudorca crassidens). The only Monodontidae displayed is the white whale, or beluga (Delphinapterus leucus). The diverse reproductive strategies and physiology among the Delphinidae alone demonstrate the importance of learning basic reproductive physiology for each species. Inefficiency and inaccuracy can occur when using one species as a model for reproductive patterns in another. As with pinnipeds, the amount of information available for each species varies tremendously, which reflects the lack of systematic research that has been conducted with most cetacean species. Before advances in manipulation and control of reproduction can occur, these systematic studies must be conducted.
Female Cetacean Reproduction Reproductive Maturity Bottlenose Dolphin
The age of sexual maturity of the Tursiops truncatus aduncus subspecies of bottlenose dolphins in the wild was estimated at over 10 years for females (Ross, 1977). Brook (1997) documented first ovulation in two captive T. t. aduncas at 6 to 7 years of age. The youngest captive bottlenose dolphin to give birth was 4 years of age; however, the majority first gave birth at 7 to 10 years (Duffield et al., 2000). In wild animals, the youngest female observed to calve was 6 years old, and the majority of females gave birth at 8 years of age (Wells, 2000). White-Sided Dolphin
Sergeant et al. (1980) and Rogan et al. (1997) estimated sexual maturity for Atlantic whitesided dolphins (L. acutus) at around 218 cm in length and 6 to 8 years of age. The authors observed a captive dolphin conceive at 3 years of age and deliver a healthy calf 1 year later (Dalton and Robeck, unpubl. data). The lack of data from wild animals precludes one from determining whether reproductive capabilities of this animal were accelerated by an increased plane of nutrition or if normal reproductive potential is as early as 3 years. Killer Whale
In the wild, sexual maturity was estimated at 8 to 10 years of age and greater than 3 m in length (Christensen, 1984). The average age at which captive killer whales first exhibited luteal activity was 9.06 ± 2.1 years (range 5.8 to 12 years, n = 9) and first conception was observed at 11.7 ± 2.9 years (range 6 to 14 years, n = 9). The average age of first calving in wild killer whales off the northwest coast of the United States was 14.9 years (Olesiuk et al., 1990). During analysis of urinary endocrine data in captive killer whales, Walker et al. (1988) and Robeck (1996) observed short transient elevations in estrogen conjugates (EC) without luteal phases, or with short luteal phases in young animals, which may have represented normal endocrine activity during reproductive maturation (Robeck, 1996). The short spikes of EC without subsequent immunoreactive pregnanediol-3-glucuronide (iPdG) appear similar to reproductive endocrine characteristics exhibited by primates during sexual maturation (Plant, 1988). Low progesterone levels and irregular short luteal phase lengths during sexual maturation also have been observed in the ovine and primate (Goodman, 1988; Plant, 1988).
0839_frame_C11 Page 205 Tuesday, May 22, 2001 11:09 AM
Reproduction
205
False Killer Whale
False killer whales were thought to attain sexual maturity at 3.7 to 4.3 m in length, and 8 to 14 years of age (Purves and Pilleri, 1978). In agreement with these data, Atkinson et al. (1999) did not observe any evidence of ovarian activity in a 6-year-old, 3.15-m female. However, another facility has recently had a 5-year-old, 347-kg, 3.4-m false killer whale conceive, although the outcome of this pregnancy is still pending (Walsh, pers. comm.). For captive false killer whales, body length at first conception is close to lengths observed in mature wild females. Beluga
Sexual maturity in white whales has been estimated at 6 to 7 years in both captive and wild populations (Braham, 1984; Calle et al., 1996). Females in captivity have conceived up to 20 years of age. This correlates with the estimated age of senescence for wild populations of 21 years (Brodie, 1971a). Reproductive Cycle
Most Monodontidae or Delphinidae exhibit seasonal reproductive activity or show seasonal trends that may reflect adaptations to food sources or climate. Photoperiod is thought to provide an environmental cue to seasonal breeders. For a species to be considered a seasonal breeder regulated by photoperiod, it must have repeatable patterns of reproductive quiescence that correlate with increasing or decreasing changes in light. In addition, physiological evidence of changes in pituitary sensitivity to gonadotropic hormones must exist. As shall be seen, two species pass the criteria for seasonal quiescence, the Pacific white-sided dolphin and the beluga; however, no data exist on seasonal pituitary down regulation. Bottlenose Dolphin
The bottlenose dolphin can be defined loosely as seasonally polyestrous (Kirby and Ridgway, 1984; Robeck et al., 1994a; Robeck, 2000). Most estrous cycling activity occurs spring through fall, but births have occurred in every month of the year. When cycling, individual animals can cycle one or more times during the year. If animals are in a breeding colony, the majority will get pregnant on the first or second estrus. Gestation for bottlenose dolphins is estimated at 12 months, and lactation can last up to 2 years or more for wild animals. Lactational suppression of estrus does occur; however, there appears to be a threshold level. When daily suckling decreases below a certain time period, usually after 1 year, ovarian activity will resume (West et al., 2000). Thus, the entire reproductive cycle or calving interval may last 3 to 4 years. Wells (2000) describes a calving interval for wild populations that varies with age class and ranges from 3 to 6 years. Females in their twenties produce calves most frequently, while younger and older females have longer calving intervals. This age-related change in fecundity is also described for captive populations (Duffield et al., 2000). In wild animals, age-associated fecundity rates may be a reflection of social status in younger animals, and reduced fertility in older animals. These factors may also play a role with captive populations; however, controlled access to females of certain age classes by males often biases captive breeding results. Managers of breeding colonies should be aware of bottlenose dolphin reproductive potential, and should try to maintain colonies that mimic natural social groupings (Wells, 2000). These natural social groups contain three basic units: (1) female/nursery groups consisting of mothers with their most recent calves; (2) juveniles in mixed-gender groups forming temporary relationships; and (3) adult males, as individuals or in pairs with strong bonds (Wells et al., 1999; Wells, 2000). White-Sided Dolphin
The Atlantic white-sided dolphin is believed to cycle in August and September and calf in June and July, suggesting an 11-month gestation period. The Pacific white-sided dolphin is seasonally polyestrous with estrous activity and births occurring from July through September in the
0839_frame_C11 Page 206 Tuesday, May 22, 2001 11:09 AM
206
CRC Handbook of Marine Mammal Medicine
United States. No information is available describing physiological control of their seasonality. Captive Pacific white-sided dolphins have exhibited an approximately 12-month gestation period (Dalton and Robeck, unpubl. data). Killer Whale
Killer whales are polyestrous. Estrus and conception occur throughout the year, with a slight, nonsignificant, seasonal increase in activity during the spring from March through August (Matsue et al., 1971; Robeck et al., 1993). Nonlactational periods of anestrus have ranged from 3 to 24 months in mature healthy females (Duffield et al., 1995; Robeck, 1996). Duffield et al. (1995) used biweekly progesterone data to describe a calving interval in captive killer whales of 32 to 58 months. Robeck (1996) found that the mean calving interval in females that were nonsuccessful at calf rearing (due to stillbirth or unsuccessful nursing) was 33 months, whereas in females that nursed successfully it was 50 months. The minimum calving interval observed for resident wild killer whales off the northern Pacific Coast of the Unites States was 36 months (Balcomb et al., 1982). Recent estimates from resident whales of this region place calving intervals from 24 months to 12 years (Olesiuk et al., 1990), with the average calving interval for wild populations estimated at 8.6 years (Balcomb et al., 1982) and 10.3 years (Bigg, 1982). The reduced calving interval of captive whales compared with wild whales is probably explained, to some extent, by nutritional and environmental differences (Matkin and Leatherwood, 1986). A decrease in reproductive productivity in response to adverse or seasonal nutritional and environmental conditions is well documented in other species (Bronson, 1988). Another possible explanation for the calving interval differences is that early postpartum or peripartum neonatal calf mortality might go unnoticed in wild killer whales. False Killer Whale
The false killer whale is polyestrous with no strong evidence for seasonality (Robeck et al., 1994b; Atkinson et al., 1999). Information on wild animals suggests that they can become pregnant any time of the year and have an estimated gestation period of 12 to 15 months (Comrie and Adams, 1938; Purves and Pilleri, 1978). Robeck et al. (1994b) described a gestation period of 14 months in a captive animal that produced a normal calf. If gestation lasts 14 months and lactation 6 to 12 months (with lactational anestrus), one could estimate a calving interval of 2.5 to 3.5 years. Atkinson et al. (1999) noted possible pseudopregnancy and prolonged anestrus in captive false killer whales with no access to males. Beluga
The beluga is seasonally polyestrous, breeding in the wild in April and May (Brodie, 1971b). Captive animals have conceived from February to June (Calle et al., 1996). This difference may be the result of latitudinal differences and associated photoperiod effects on breeding activity, although there is no evidence to confirm this. Calving in wild belugas has been observed from July to September, and in captive animals from May through September. Gestation lengths have been estimated at 14.5 months for wild populations and 15 to 17 months for captive ones (Brodie, 1971a; Calle et al., 1996). Animals have been observed to nurse for 2 years, and at least one animal conceived during the spring season after a previous summer birth; thus, lactational anestrus may not occur in this species. The calving interval for captive animals is as little as 3 years (Brodie, 1971a; Seargent, 1973; 1980; Braham, 1984). Estrous Cycle and Ovarian Physiology Bottlenose Dolphin
Most of the published information on cetacean reproduction concerns the most common cetacean in captivity, the bottlenose dolphin (Robeck et al., 1994a). Existing endocrine data has come from weekly or biweekly blood sampling of trained captive animals. This type of
0839_frame_C11 Page 207 Tuesday, May 22, 2001 11:09 AM
Reproduction
207
FIGURE 2 Mean (±S.D.)(n = 35 estrous cycles) pattern of development for the dominant follicle prior to ovulation in four T. t. aduncus. FD = Follicle diameter. (From F. Brook, Hong Kong Polytechnic University, Kowloon, Hong Kong, 1997, 339.)
sampling frequency is sufficient to describe seasonality or estrous cycle patterns, but is not adequate to the pulsatile endocrine activity that occurs in proximity to ovulation, or other important ovarian events. At best, one could hope to catch an estrogen surge, but without serial sampling, few to no conclusions can be drawn. Urinary and fecal sampling or other noninvasive techniques that can be performed daily offer the best hope for describing and eventually predicting ovarian and endocrine relationships. Urinary endocrine monitoring offers great promise, but, until recently, the only species that had been reliably trained for this procedure was the killer whale, although many facilities have now reported success in training bottlenose dolphins. A wealth of information on bottlenose dolphin reproductive physiology and follicular dynamics has recently been collected through the use of sonographic ovarian analysis (Brook, 1997; 2000; Robeck et al., 1998; 2000). However, this technique, has yet to be combined in adequate endocrine monitoring to describe how hormonal events relate to ovulation. Harrison and Ridgway (1971) reported on the gonadal activity of 22 female bottlenose dolphins. In these animals, most of the follicles were 2 mm or less in diameter with no follicles greater than 5 mm, although there was an accessory CL formed from a luteinized follicle 10 mm in diameter. Brook (1997) used ultrasonography to follow follicular activity in bottlenose dolphins (T. t. aduncus) and provide the first real-time description of folliculogenesis in cetaceans. Multiple 2- to 3 mm-diameter follicles were often observed on the ovary, regardless of ovarian activity. Once a follicle was larger than 3 mm, it could be classified as developing. In 32% of observed cycles (n = 37), more than one follicle developed beyond 4 mm in diameter. The dominant or primary follicle appeared 1 to 2 days prior to ovulation, when it was distinguished from other follicles by its size. Only one follicle was seen to ovulate, subordinate follicles regressing either before or just after ovulation. Ovulation occurred at a mean of 8 days after the dominant follicle reached 10 mm in diameter (Figure 2). Preovulatory follicles ranged in size from 18 to 28 mm, with a mean of 20.9 mm (Figure 2).
0839_frame_C11 Page 208 Tuesday, May 22, 2001 11:09 AM
208
CRC Handbook of Marine Mammal Medicine
There appeared to be a loose correlation between the size of the dolphin and the size of the preovulatory follicle in this population, although the number of females studied in detail was small and this remains to be confirmed. There is evidently significant individual variation in preovulatory follicle size and it is essential to assess each animal over time in order to use follicular size to predict ovulation. The maximum diameter of “normal” CLs (i.e., not associated with pregnancy or pseudopregnancy) observed ranged from 21 to 36 mm. Again, the largest CLs were seen in the larger females. Estrous cycle length in T. t. aduncas is about 30 days. For T. t. truncatus, cycle lengths of 21 to 42 days have been estimated from serum hormone levels (Sawyer-Steffan and Kirby, 1980; Kirby and Ridgway, 1984; Schroeder, 1990b). Periods of anestrus not associated with gestation or lactation occur in Tursiops (Brook, 1997). At these times, ovulation does not occur and the ovaries appear to “shut down.” Periods of anestrus of up to 27 months have been documented in T. t. aduncas, but the significance of this phenomenon is not understood. Killer Whale
The only cetacean species in which detailed information on gonadotropic hormones has been collected is the killer whale. Walker et al. (1988) used urinary progesterone and estrogen metabolites, and bioactive FSH, to describe endocrine changes that occurred during two estrous cycles. Based on their results, they predicted a wave of follicular activity that begins before peak estrogen levels, but the temporal relationship between peak plasma estrogen and ovulation could not be determined. Urinary LH levels can be quantitatively detected in killer whales although twice-daily urine samples are necessary to describe the LH peak or surge consistently (Robeck et al., 1990; Robeck, 1996). Recent data suggest the LH surge occurs around 12 hours after the peak estrogen surge (Robeck et al., unpubl. data). The mean estrous cycle length based on the beginning of luteal phases was 41.2 days (range 19 to 49 days), the follicular phase lasts around 18 days, and the luteal phase lasts around 20 days (Robeck, 1996). Anestrus periods as long as 2 years, which are not associated with gestation or lactation, have been observed in killer whales (Robeck, unpubl. data). Copulatory activity of killer whales has been compared with qualitative estimates of vaginal mucus secretion and endocrine data (Robeck, 1996). A higher percentage of mucus secretions and copulations occurred around peak levels of EC rather than peak levels of LH, and heavy vaginal mucus secretion was often associated with estrus or receptivity. Although mild mucus secretion occurred during various phases of the estrous cycle, all of the heavy vaginal secretion occurred during periods of detectable EC. Thus, it appears that in the killer whale, as with other terrestrial species, estrogens, presumably produced from developing follicles, are responsible for stimulating sexual activity (probably by changing female receptivity) and producing secretory changes (i.e., vaginal and cervical mucus secretions) required for conception. Limited observations with killer whale ovaries suggest a different pattern for developing follicles from that in the bottlenose dolphin. Follicles destined to ovulate appear to develop over two cycles, with the size of the follicle ranging from 2.5 to 4.5 cm at the start of the follicular phase (Figure 3). As many as four preovulatory follicles have been observed on the ipsilateral ovary and at least two on the contralateral ovary (Robeck, unpubl. data). More observations are needed to understand better the range of patterns that naturally occur in this species. False Killer Whale
Prolonged periods of elevated serum progesterone or pseudopregnancy may occur with regularity in false killer whales. Serum progesterone and hydrolyzed conjugated progesterone in daily urine samples from two female false killer whales indicated prolonged luteal or pseudopregnant periods of elevated progestin for 378, 202, 36, and 24 days (Robeck et al.,
0839_frame_C11 Page 209 Tuesday, May 22, 2001 11:09 AM
Reproduction
209
FIGURE 3 Killer whale with a large dominant follicle approximately 3.5 cm in diameter. This follicle was 12 days from an ovulation that resulted in an artificial insemination (with cooled transported semen) pregnancy. The dotted lines represent the ovarian length (9.9 cm). (From T. Robeck, unpubl. data.)
1994b). Atkinson et al. (1999) measured weekly serum progesterone concentrations and observed a prolonged period of ovarian activity from March to December. Periods of anestrus not associated with gestation or lactation of 3 to 10 months have been observed in false killer whales (Atkinson et al., 1999). Suckling (Lactational) Suppression of Estrus
During a 10-year period of observations on one group of bottlenose dolphins (T. t. aduncus), ovulation during lactation was never observed (Brook, unpubl. data). On one occasion, a female was accompanied by a 1.5-year-old calf, but suckling was not observed for several weeks. This animal was seen to ovulate once, but then her calf slid over the enclosure wall and stranded on the poolside. Although physically unharmed, intensive suckling behavior resumed when the calf was returned to the mother, and continued for some time. The mother did not cycle again for several months until suckling stopped again. Robeck (1996) provides strong evidence of lactational, or suckling, suppression of estrous activity in killer whales. There were significant differences between postpartum return to estrus in lactating (mean 481.4 days; range 159 to 983 days) and nonlactating (mean 65.8 days; range 31 to 122 days) females. Lactation alone does not suppress estrus. This was demonstrated by West et al. (2000) when they collected milk samples from lactating dolphins with or without suckling calves for up to 402 days postpartum. Although these dolphins were lactating, cycling began after the calf had been weaned, or, if the calf was stillborn, within a relatively short period. When an animal is lactating, total suckling time can drop below the minimum threshold duration of stimuli required to suppress estrus, and the animal will return to estrus. This usually occurs in females with older calves that obtain most of their nutrition from fish, but will still occasionally nurse when presented with the opportunity. This threshold effect may be related either to decreased sucking stimuli or to a built-in time clock that reduces the hypothalamic inhibitory effects of suckling stimuli after a certain period postpartum, or a combination of the two. In general, lactational alteration of reproductive function is believed to be caused by suckling stimuli, which suppresses gonadotropin (particularly LH) secretion, preventing normal follicular maturation and ovulation (McNeilly, 1988). In dolphins and
0839_frame_C11 Page 210 Tuesday, May 22, 2001 11:09 AM
210
CRC Handbook of Marine Mammal Medicine
killer whales, therefore, it appears that suckling (which also helps to maintain lactation) plays an important role in regulating the calving interval. Corpora Albicantia and Asymmetry of Ovulation
Histological changes in ovarian structures in the bottlenose dolphin and other delphinids have been described in detail (Harrison, 1969; Benirschke et al., 1980; Perrin and Reilly, 1984). Corpora albicantia (CA) are believed to be retained indefinitely in pilot whales (Globicephala macrorhynchus), but are only retained when they have originated from corpora lutea of pregnancy in other species, such as bottlenose dolphins and Stenella spp. (Harrison, 1969; Marsh and Kasuya, 1984; Perrin and Reilly, 1984). This has recently been confirmed by analysis of ovaries from a bottlenose dolphin whose entire reproductive history, including ovulations and pregnancies, was documented by ultrasound (Brook, unpubl. data). Based on histological identification of CA, ovulation and pregnancy in the bottlenose dolphin occurred in the left ovary and left uterine horn more than 68% of the time (Ohsumi, 1964; Harrison and Ridgway, 1971). Brook (1997) found similar asymmetry with respect to ovulation in T. t. aduncus. Asymmetry exists in other cetaceans; yet the physiological mechanisms for this are unknown (Ohsumi, 1964; Perrin and Reilly, 1984; Bryden and Harrison, 1986). Pseudopregnancy
Pseudopregnancy occurs in bottlenose dolphins, killer whales, and false killer whales. The cause of pseudopregnancy in delphinids is unknown and may be multifactorial. In terrestrial species (without obligate embryonic diapause), the most common cause is early embryonic loss after the embryo has released pregnancy-specific proteins that are involved with MRP. Thus, the maternal uterus “believes” it is pregnant, release of prostaglandin is inhibited, and the CL maintains secretion. For pseudopregnancy to continue for any significant duration, however, there must be a source of gonadotropins to maintain the CL. As discussed below, in killer whales, it appears that at least early pituitary LH is responsible for CL growth and development. If fetal death occurs after placental formation, it may be a local source (Hobson and Wide, 1986). Using ultrasound, Jensen (2000) described early fetal abortion in a bottlenose dolphin. Although data are inconclusive, it appears that the fetus died approximately 3 to 4 weeks before CL progesterone secretion stopped, and that the abortion of the dead fetus coincided with basal progesterone concentrations. Ultrasound evaluation of early embryonic loss and how, or if, the timing of such events affects the endocrine system may help determine whether it plays a role in pseudopregnancy. Pseudopregnancy occurs with some frequency in females without access to males. If pseudopregnancy only occurred in females without access to males, then it would be easy to blame the unnatural social groups found in managed environments as the cause for these conditions. Cowan (2000), however, reported a number of wild dolphins having luteal cysts that could result in pseudopregnancy. In killer whales, pseudopregnancy tends to occur in animals that have cycled multiple times (more than four cycles) without becoming pregnant. It is not dependent on age, but once an animal has experienced pseudopregnancy, it appears more likely to experience it a second time. Although killer whales will cycle multiple times during a season, this polyestrous activity occurs only in the absence of a fertile male, and as such, would probably not occur in wild populations. The rate of pseudopregnancy among wild animals is not known.
0839_frame_C11 Page 211 Tuesday, May 22, 2001 11:09 AM
Reproduction
211
Management of pseudopregnancy via prostaglandin F2α administration is a viable option for returning females to the breeding pool and maximizing their reproductive potential (Robeck et al., 2000). Pregnancy Bottlenose Dolphin
The use of ultrasound to monitor pregnancy in captive cetaceans provides valuable data on fetal morphology, development, and well-being and on maternal gestation length in bottlenose dolphins, although there remains a need for normal data (see Chapter 26, Ultrasonography) (Williamson et al., 1990; Taverne, 1991; Brook, 1994; Stone et al., 1999; Sweeney et al., 2000). Gestation periods in Delphinidae vary. The gestation period for bottlenose dolphins has been estimated at 12+ months. Recent data in T. t. aduncus with known conception dates places these values at 370 ± 11 days (Brook, 1997). Plasma progesterone levels recorded during pregnancy in bottlenose dolphins range from 2.0 to 56.0 ng/ml (Sawyer-Steffan and Kirby, 1980; Schroeder and Keller, 1989). Killer Whale
Robeck (1996) used high-performance liquid chromatography (HPLC) to describe progesterone metabolite secretion during the luteal phase, and early, mid, and late pregnancy in killer whales. The presence of only one major immunoreactive metabolite during these periods provides evidence for the presence of a single source of progesterone. These data support the commonly proposed hypothesis that maintenance of pregnancy relies heavily on luteal production of progesterone. Recent data demonstrated an increase in the frequency of LH surges soon after conception but after initial luteal progesterone levels had begun to increase (Robeck, 1996). This increase in high-amplitude LH secretion was not observed during the luteal phases of nonconceptive cycles. A similar increase in LH secretion during the early luteal phase has been observed in Asian elephants (Elephas maximus) (Brown et al., 1991). However, unlike the killer whale, this increased LH secretion is not limited to conceptive cycles. The hypothesized significance of these early-luteal-phase LH surges in the elephant was to aid in the formation of a critical mass of luteal tissue necessary for the maintenance of pregnancy (Brown et al., 1991). Since these LH surges were observed after progesterone had begun to rise, they may be needed for stimulating maintenance of the existing CL or for formation of accessory luteal structures. Similarly, the killer whale may require additional LH release for correct formation of the CL of pregnancy (Robeck, 1996). This differential secretion of LH during the early progesterone secretion of a conceptive cycle rather than a nonconceptive cycle should result in the formation of two different types of CL structures. The CL that has been supported by additional LH secretion should theoretically be developed to a greater degree than the one without this additional stimulation. This theory is supported by the presence of two types of luteal scars or CA on the ovaries of odontocetes (Harrison, 1969; Fisher and Harrison, 1970; Harrison et al., 1972). Type 1 CAs are typically 5 to 10 mm in diameter and represent the remnants of a well-vascularized and organized CL. Type 2 CAs are usually smaller, 3 to 5 mm, and appear to represent a less well developed or organized CL. Although there is some debate over the significance of these histologically distinct CAs, type 1 CAs are believed to be associated with pregnancy and type 2 CAs are believed to represent anovulatory luteinized follicles or nonconceptive ovulations, (Harrison and Ridgway, 1971; Gaskin et al., 1984; Marsh and Kasuya, 1984).
0839_frame_C11 Page 212 Tuesday, May 22, 2001 11:09 AM
212
CRC Handbook of Marine Mammal Medicine
Beluga
Gestation length in the beluga has been estimated as 14.5 to 17 months. Little is known about the physiology of pregnancy in this species. Pregnancy appears to be maintained by the CL of pregnancy. Accessory corpora (luteinized follicles) have been found in 11 to 15% of pregnant belugas, but because of their low incidence, they are obviously not required to maintain pregnancy. They also indicate that follicular growth can occur in pregnant animals, possibly during the next breeding season, when they would normally come into estrus. Calle et al. (1996) pooled mean monthly gestational plasma hormone levels for captive animals; progesterone levels ranged from 0.97 ± 1.14 to 42.86 ± 12.00 ng/ml and estrogen ranged from 13.93 ± 11.62 to 30.62 ± 12.43 pg/ml. Pregnancy Diagnosis
Pseudopregnancy occurs with such regularity in dolphins, killer whales, and false killer whales that an animal cannot be confirmed pregnant without the use of ultrasonography (see Chapter 26, Ultrasonography). Despite the regular occurrence of pseudopregnancy, it is still a relatively newly described phenomenon that undoubtedly has always occurred, and may have led to an overestimation of abortion rates. Because of its recent recognition, and the slow integration of ultrasound into clinical practice, data are insufficient to allow accurate descriptions of its frequency and to determine which class of animal is most susceptible. Parturition
The mechanism of control of parturition in cetaceans is unknown; however, there appears to be an interaction between hormones produced by the fetal–placental uterine axis. Six major hormones, and probably others, appear to be intertwined during the induction of parturition. These hormones include estrogens, progesterone, adrenal steroids, oxytocin, relaxin, and prostaglandins. Stages of Parturition
Early stages of pregnancy generally have similar behavioral components. The most common behavioral signs are listed in Table 2. The table was designed as a quick reference to some important periparturient events. Many of these events, such as first nursing, can have extreme variability in length, so it is important to remember these guidelines cannot replace careful clinical observation of each situation. For example, if when using the table to determine interval to first nursing, it may be comforting to know that to the authors’ knowledge bottlenose dolphin calves have lived even after failing to nurse for up to 48 hours. However, the level of comfort of the clinician attending a parturient cetacean should be dependent upon the behavior, condition, and activity of the cow and calf. A predictor of parturition not on the table is a decrease in rectal temperature 24 hours prior to stage-two labor. These data have recently been collected for both the bottlenose dolphin and the killer whale. It requires minimal training to condition the animals to obtain a daily body temperature and may provide an objective indicator for predicting parturition (Katsumata et al., 1999a; Terasawa et al., 1999). Recognition of the onset of parturition is an important management tool. Most reproductiverelated problems (dystocia, stillbirth, weak calf, poor maternal care) occur, and can be observed, in the first few hours after delivery. Induction of Parturition
Although the hormonal control of parturition is not understood, administration of hormones in appropriate combinations can result in the induction of parturition, sometimes with less-than-satisfactory results (Catchpole, 1991). Induction of parturition should not be
115 min (n = 1)
>5 h F (93%, n = 15) 8–12 Usually after stage 3, >12 h <48 h
188.8 (20–600) min F (98.1%) <12 Often after stage 3, <12 h
Lactation (months) Birth to first fish (months)
Calf behavior critical, <36 h 26.6 (18 to 36) 5.8 (2.5 to 27)
94.3 (45–240) min
15–24 3–6
228 min (n = 1)
Length of stage-2 labor For animals with live calves For animals with dead calves Presentation Birth to stage 3 (h) Birth to nursing Normal Maximum
Flukes appear, VD
8–12 2–3
48 h
F (100%, n = 5) 6–20 <15 h
See bottlenose dolphin
Unknown 12 months See bottlenose dolphin
29–35 days 12 months MD, VD, DA, CT, DBT
Estrous cycle length Gestation length Stage-1 labor signs (within 24 h of stage 2) Stage-2 labor signs
Seasonal polyestrous July–Oct.
Polyestrous All year, peak spring–fall 39–45 days 17 months See bottlenose dolphin See bottlenose dolphin 60–240 min
White-Sided Dolphin (Lagenorhynchus c obliquidens)
Polyestrous All year, peak spring–fall
Killer Whale b (Orcinus orca)
Reproductive pattern Period of activity
Characteristic
Bottlenose Dolphin a (Tursiops truncatus)
TABLE 2 Reproductive Parameters of Cetaceans
18
Unknown
F (n = 1) 5.8 7h
?
165 min (n = 1)
Flukes appear
Variable 14 months Arching
Polyestrous All year
False Killer Whale d (Pseudorca crassidens)
f
g
(Continued)
24–36 10 (6–23)
33 h
F (14% HF) 6.2–8.3 <18 h
>2 days
See bottlenose dolphin 392 (136–870) min
Unknown 14.5 months Arching, VD
Seasonal polyestrous Feb.–June
Beluga (Delphinapterus e leucas)
0839_frame_C11 Page 213 Tuesday, May 22, 2001 11:09 AM
Reproduction 213
2.5 yr 4 yr 5.8 yr 5.8–12 yr 10 yr 10–12 yr
2.9 yr 3.6 yr 4 yr 7–10 yr 8 yr 8–10 yr
Killer Whale b (Orcinus orca)
6–8 yr
Unknown 3 yr 3 yr 3–6 yr Unknown
White-Sided Dolphin (Lagenorhynchus c obliquidens)
8–14 yr
Unknown Unknown 5 yr 8–14 yr Unknown
False Killer Whale d (Pseudorca crassidens)
8–9 yrs
3 yr 6 yr 6–7 yr Unknown
Beluga (Delphinapterus e leucas)
214
Key: CI = calving interval; CT = contractions; DA = decreased appetite; DBT = decrease in basal temperature; F = flukes first; HF = head first; MD = milk discharge; VD = vaginal discharge. a From Andrews et al., 1997; Duffield et al., 2000; Joseph et al., 2000; Sweeney et al., 2000; Wells, 2000. b From Duffield et al., 1995; Robeck, 1996; McBain, Reidarson and Walsh, pers. comm. c From Sergeant et al., 1980, Dalton et al. 1995; Rogan et al., 1997. d From Comrie and Adams, 1938; Purves and Pilleri, 1978; Robeck et al., 1994b; Atkinson et al., 1999; Walsh, M. 2000; pers. comm. e From Brodie, 1971a; Braham, 1984; Dalton et al., 1994; 1996; Calle et al., 1996. f Both calves had to be manually extracted. g The authors have had one calf go 5 days without nursing; however, intensive management and intravenous IgG were required to keep the calf alive.
CI, nonviable calf CI, viable calf Youngest mature female Sexual maturity: female Youngest mature male (sired a calf) Sexual maturity: male
Characteristic
Bottlenose Dolphin a (Tursiops truncatus)
TABLE 2 Reproductive Parameters of Cetaceans (Continued)
0839_frame_C11 Page 214 Tuesday, May 22, 2001 11:09 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C11 Page 215 Tuesday, May 22, 2001 11:09 AM
Reproduction
215
attempted, therefore, unless the clinician feels it is the only recourse available. With a wide range in gestational lengths within species, and a usually speculative conception date, induction of “overdue” calves is never indicated. In the authors’ clinical experience, and in most cases, attempting to induce delivery of an apparently dead, in utero fetus is not indicated. If uterine infection is the cause of the dead fetus, the cow can be placed on antibiotics until she aborts the fetus. At that time, uterine, placental, and fetal cultures can be obtained to ensure effective treatment. In addition, the postpartum uterus is easily catheterized for local treatment. However, if the clinician feels induction is necessary, prostaglandin F2α has been used successfully to induce parturition in a beluga (Robeck, unpublished data). In this case, 40 mg PgF2α IM, BID, for 4 days caused progesterone to decrease to less than 1 ng/ml and stage-two labor to commence 7 days after the final injection. Another attempt at induction of parturition was of a midterm fetus in a bottlenose dolphin. The animal had a serious systemic infection, and based on a history of difficult pregnancies, it was believed that the fetus posed a risk to the cow’s health. Thus, multiple doses of prostaglandin F2α were administered until a response was observed. No response in circulating progesterone was observed until a single dose of 60 mg was used. The animal finally went into labor, but was unable to pass the calf and died during manual extraction. The reader must understand that the efficacy of these protocols is not well established, so sound clinical judgment should be employed. Early (60-day or less) unwanted pregnancies may be a situation where the chance of success at inducing abortion is greater than the risks. Because prostaglandin has been effective for CL lysis in pseudopregnant animals (Robeck et al., 2000), one can only speculate that application of these protocols in early gestation might be successful.
Male Cetacean Reproduction Sexual Maturity Bottlenose Dolphin
Postmortem assessment of sexual maturity in males is based on testis weight, diameter of the seminiferous tubules, presence of spermatozoa in the seminiferous tubules, and presence of seminal fluid in the epididymis (Perrin and Reilly, 1984). Observations of the gonads of bottlenose dolphins from Florida waters suggested that the age of sexual maturity for males was 10 to 13 years (Seargent et al., 1973; Perrin and Reilly, 1984; Cockcroft and Ross, 1990), but may begin as early as 9 years (Cockcroft and Ross, 1990). Males recovered on the east coast of South Africa were estimated to attain sexual maturity at 14.5 years of age (Cockcroft and Ross, 1990). Normal ejaculate was obtained from a 7-year-old captive T. t. aduncus (Brook, 1997). Captive animals are maintained under artificial social conditions that often allow younger animals opportunities to breed successfully. In the wild, the presence of a physically dominating male appears to exclude reproductively mature, but physically immature, males from successfully mating until they reach at least 20 years of age (Duffield and Wells, 1991). White-Sided Dolphin
Sexual maturity occurs when males reach 2 to 4 m in length and 6 to 8 years of age (Sergeant et al., 1980; Rogan et al., 1997). Killer Whale
Wild killer whales have been estimated to reach sexual maturity at 15 to 16 years of age and 6 to 7 m in length (Bigg, 1982; Christensen, 1984). By evaluating serum testosterone in biweekly samples, Robeck et al. (1995) concluded that male killer whales are fertile as early as 10 years of age. Younger animals, however, were not included in the study, so the earliest age when mature testosterone levels were produced could not be determined. Katsumata et al. (1999b)
0839_frame_C11 Page 216 Tuesday, May 22, 2001 11:09 AM
216
CRC Handbook of Marine Mammal Medicine
used similar methods to estimate age of sexual maturity (based on testosterone) in one male at 12 years of age. Testosterone concentrations for this animal were below 1 ng/ml until reaching maturity at 12 years. Beluga
Sexual maturity was estimated at 8 to 9 years of age in belugas (Brodie, 1971a). Seasonality
Influences on male seasonality have yet to be investigated. Presence or absence of mature cycling females and other males, dominance hierarchies, size of the breeding population, and environmental or nutritional cues may all play some role in modification of seasonal levels of fertility. Bottlenose Dolphin
Harrison and Ridgway (1971) found evidence for seasonal variation in testosterone levels of bottlenose dolphins, which were elevated to 14 to 24 ng/ml in September and October, as well as in April and May. Peak testosterone levels correlated well with peak breeding activity. Schroeder and Keller (1989) measured serum testosterone levels and sperm production in a 19-year-old bottlenose dolphin. Blood samples were collected twice monthly, and ejaculate was obtained twice weekly, over a 28-month period. Testosterone levels ranged from 1.1 to 54.1 ng/ml, with increasing levels from April to a peak in July in two consecutive seasons (Schroeder, 1990b). Peak sperm production and density, however, occurred during the breeding season, late August through October, when testosterone levels were lowest. Other seasonally reproductive species exhibit peak sperm production after serum testosterone peaks (Byers et al., 1983; Asher et al., 1987; Matsubayashi et al., 1991). This delay may represent the observed inhibitory effects that high testosterone can have on spermatogenesis (Matsumoto, 1990; Tom et al., 1991). Submaximal concentrations of testosterone may be required for optimum sperm recruitment. This is supported by the observation that normal spermatogenesis can occur in the presence of low intratesticular testosterone concentrations (Cunningham and Huckins, 1979). The delay may also represent the normal lag time from spermatocyte recruitment (which is maximally stimulated during peak testosterone) to sperm maturation in dolphins (Byers et al., 1983; Asher et al., 1987). Kirby (1990) summarized data of serum testosterone levels in bottlenose dolphins and reported that twice weekly samples from five male dolphins over periods of 6 to 24 months allowed classification of individuals as immature, pubescent, or sexually mature. Testosterone levels in mature animals (13 to 15 years of age) fluctuated between 2 and 5 ng/ml, rising above 10 ng/ml in the breeding season. Puberty in males has been estimated as the time when testosterone levels first rise from less than 1 ng/ml to 10 ng/ml. In contrast, Brook et al. (2000) and Brook (1997) determined that mature male T. t. aduncus can exhibit testosterone levels below 1.0 ng/ml, and found sonographic testicular echo texture a more reliable indicator of maturation. More significantly, they did not find changes in testicular echo pattern with season, and only a slightly seasonal pattern of testosterone production. Data from Brook et al. (1996; 2000) support the basic presumption that temperate animals would have less nutritional or environmental pressures for the development of seasonal breeding patterns. Thus, although numerous studies show increases in fecundity during predictable periods, dolphins remain fertile throughout the year, and can only be classified as facultatively seasonally polyestrous. Social patterns as opposed to environmental patterns (photoperiod, temperature) may have been the overwhelming pressure behind the development of the slightly seasonal trends. White-Sided Dolphin
Research conducted in Japan indicated that at least one male Pacific white-sided dolphin had a well-defined breeding seasonal where sperm was only collected from May to September
0839_frame_C11 Page 217 Tuesday, May 22, 2001 11:09 AM
Reproduction
217
(Yoshioka et al., 2000). The data included testosterone levels and sperm production and illus8 trated peak sperm concentrations in June of 19.3 × 10 /ml (mean from May to September = 8 3.8 × 10 /ml ± 0.65) to total azospermia from November through April. Although data are limited and different from bottlenose dolphins or seasonal breeders, peak testosterone occurred simultaneously with peak sperm production in May and June. From this information and assuming it holds true for the species as a whole, Pacific white-sided dolphins have a short seasonal reproductive period, synchronized with female cyclicity and regulated by an unknown physiological mechanisms. Killer Whale
No significant seasonal changes in testosterone levels were observed in biweekly serum samples from five male killer whales 10 years old or older, although mean testosterone was significantly lower in October. Testosterone concentrations ranged from a low in October of 1.4 ng/ml to a high in April of 2.2 ng/ml, with peak levels occurring from March to July (Robeck et al., 1995). As would be expected, no significant seasonal patterns have been observed in sperm concentration voluntarily collected from a captive male killer whale (Robeck, unpubl. data). Thus, in agreement with observed calving periods and female cyclic activity, killer whale males appear to be fertile throughout the year, with possible peak fertility occurring in the spring and summer (Robeck et al., 1995; Katsumata et al., 1999b). False Killer Whale
The only reproductive data from male false killer whales are testosterone levels that have no obvious seasonal trend (Robeck et al., 1994b). Beluga
In 11 captive male belugas 3 to 21 years old, mean circulating testosterone concentrations were lowest in September (0.9 ng/ml) and highest 6 months later in March (4.95 ng/ml) (Dalton et al., 1994). Mean testosterone levels gradually rose throughout the fall and were elevated (>3.5 ng/ml) from January through April, then declined to the nadir in September (Calle et al., 2000). The relationship between circulating testosterone and spermatozoa production is unknown, although if belugas are physiologically similar to other seasonal mammals, sperm production should peak 1 to 2 months after peak testosterone. If this proves true, captive beluga males should have peak sperm production in May or June.
Contraception and Control of Aggression Females
To the best of the authors’ knowledge, the only method of contraception attempted in female Delphinidae involves the use of the oral progestin, altrenogest (Regu-Mate®, Hoechst Roussel Vet, Melbourne, Australia), which is a relatively safe contraceptive. Altrenogest has been used effectively in several different animals to regulate the estrous cycle without producing any detrimental side effects (e.g., reduced fertility or abnormal behavioral patterns). It was developed for use in the mare (Squires et al., 1979; 1983; Webel and Squires, 1982), but has since proved effective in the sow (Kraeling et al., 1981; Stevenson and Davis, 1982), the giraffe, and the okapi (Loskutoff, pers. comm.). Regu-Mate has been used long term without any clinical evidence of damaged fertility in a killer whale (Young and Huff, 1996), Pacific white-sided dolphin, and bottlenose dolphins (Asa, 2000; Dougherty et al., 2000). It must be administered daily (0.05 mg/kg) by mouth and should (although no data exist to confirm this) be effective after 2 days of administration. Progestins typically do not inhibit follicular growth; thus animals on Regu-Mate may still exhibit behavioral estrus.
0839_frame_C11 Page 218 Tuesday, May 22, 2001 11:09 AM
218
CRC Handbook of Marine Mammal Medicine
Males
Most efforts in marine mammal contraception have been primarily to control fertility and aggression in males. In male bottlenose dolphins, the GnRH agonist leuprolide acetate (Lupron®, Tap Pharmaceuticals, Inc., Deerfield, IL) has been successfully used to cause azospermia and is currently the only recommended form of contraception for male bottlenose dolphins (Briggs, 2000). Its mechanism of action has previously been discussed in the pinniped contraception section. If the primary objective for its administration is the reduction of circulating testosterone and related aggression, then the clinician should understand that initial serum testosterone concentration may double, and a measured increase in aggression may be observed. Serum testosterone should subside by day 10, and reach basal concentrations from day 14 to 20. The major disadvantages of its use include the need for monthly or bimonthly injections, and cost. A newer generation of GnRH agonist, Deslorelin (Peptech Ltd, North Ryde NSW, Australia) has shown good activity for as long as a year in carnivores (Jochle, pers. comm.). Its application for marine mammals is under investigation.
Reproductive Abnormalities in Cetaceans Cystic follicles with varying degrees of luteinization were reported in the short-finned pilot whale (Marsh and Kasuya, 1984). Cystic follicles have been known to produce estrogens and progesterone depending on the degree of luteinization that occurs (Youngquist, 1986; Carriere et al., 1995). By using ultrasound, cystic follicles have since been visualized in bottlenose (Brook, unpubl. data; Jensen, unpubl. data; Robeck, unpubl. data) and Pacific white-sided dolphins (Robeck, unpubl. data). Luteinized cystic follicles may be partially responsible for pseudopregnancy that can occur in at least three delphinid species (Figure 4). Prolonged luteal phases in domestic animals have been associated with uterine infection or inflammation, early embryonic loss, and diestrus ovulations (Hinrichs, 1977). No clinical evidence exists to suggest an inflammatory process as causing prolonged or erratic luteal phases in the authors’ cases, although frequent and timely ultrasound examinations during ovarian activity may help explain these phenomena.
FIGURE 4 Luteinized follicular cyst in a T. t. aduncus. (From F. Brook, unpubl. data.)
0839_frame_C11 Page 219 Tuesday, May 22, 2001 11:09 AM
Reproduction
219
Dystocia with fetal death has occurred in cetaceans. In these situations, intervention was usually delayed until fetal death had occurred, so the only remaining concern was for the cow. Chapter 30, Intensive Care, reviews treatments that have been used for dystocia in various cetacean species. By far the most frequent pathology associated with reproduction is stillbirth. A recent survey revealed 8% abortion and 8.8% stillbirth rates in bottlenose dolphins from 1995 through 2000 (Joseph et al., 2000). Only a few females were responsible for a high percentage of stillbirths and neonatal deaths. These females should be identified, and environmental or physiological conditions that may be contributing to poor reproductive success should be changed. Furthermore, as Miller and Bossart (2000) point out in their review of reproductive-related pathology in bottlenose dolphins, the fetus and placenta should be submitted for culture and histology, in an effort to determine potential infectious causes for reproductive failures.
Artificial Insemination Artificial insemination (AI) can be an important and powerful tool for genetic management of captive populations. However, it is usually most effective when applied to populations that are reproducing successfully. AI does not replace, but rather enhances, reproductive efficiency. Neither can it be viewed as the sole solution to infertility or other reproductive abnormalities (Lasley and Anderson, 1991; Wildt, 2000). AI has recently been successful in at least three individuals of two different species; killer whales and bottlenose dolphins (T. t. aduncus) (Robeck unpubl. data; Brook, unpubl. data). The development of AI in these two species is of no surprise as they are the two cetacean species in which most of the basic reproductive physiological research has occurred. Although these successes provide insight into what might be accomplished when these techniques become routine, many challenges remain before that vision can be realized. There are many techniques that must be improved or investigated (depending upon the species) before AI can be developed in other cetacean species. These techniques include semen collection, handling, and storage, ovulation detection, estrus synchronization, and insemination techniques. Perhaps the biggest obstacle to applying any successful AI techniques to cetaceans is the intense management that must occur. It is the job of investigators, not only to develop AI and related technologies, but also to use methodologies that can be applied to a wide range of husbandry situations with minimal additional equipment and training. Once this has been accomplished, assisted reproductive techniques will truly make an impact on captive cetacean management. Semen Collection and Storage
Much has been written about early successes in freezing semen (Hill and Gilmartin, 1977; Fleming et al., 1981; Seager et al., 1981; Schroeder and Keller, 1989). This section reviews some of this work, but focuses on recent and current, often unpublished, work that has been performed since these earlier trials. The sensitivity of semen to cryopreservation and to various cryopreservation methods varies among species and individuals (Watson, 1979; Senger, 1986; Howard et al., 1991). Schroeder (1990b) found the post-thaw motility of semen frozen with lactose-based egg yolk extender to be greater than that of semen frozen in a fructose-based extender. His extender was composed of 11% lactose or fructose, 6% glycerol, and 20% egg yolk (1000 IU/ml Penicillin G and 1.25 mg/ml streptomycin sulfate were included in the extender). Few other studies with dolphin semen have attempted to evaluate and/or compare other major variables that can have important influences on the success of cryopreservation attempts. These variables include the effects of cryoprotectants and diluents on in vitro longevity at varying
0839_frame_C11 Page 220 Tuesday, May 22, 2001 11:09 AM
220
CRC Handbook of Marine Mammal Medicine
TABLE 3 A Simple Method for Cryopreservation of Cetacean Semen 1. Warm Extender A (without glycerol) to 35°C in water bath in preparation for extension. 2. Once semen is in the laboratory, determine total motility (TM), percent progressive motility (PPM), and rate of forward motility (RFM). TM and PPM are determined by visual 6 estimation of extender-diluted semen (diluted to ∼25 × 10 ). RFM is judged on a scale of 0 to 5: 0 = no forward motility; 1 = little forward movement; 2 = movement and poor progression; 3 = slow forward progression; 4 = steady forward progression; 5 = rapid forward progression. 3. Slowly (over 5 min) dilute semen with equal volume of extender (1:1 dilution). Take care to mix semen gently while the extender is added. 4. Place diluted semen into a conical vial and store at 5°C for 2 hours. 5. Place a volume of Extender A that is equal in volume and initial temperature to the extended semen in the refrigerator at the same time. The temperature of these two vials (extender and extended semen) should remain the same. 6. Place a vial of Extender B (with 14% glycerol) into the refrigerator. The vial should contain enough glycerolated Extender B to extend the maximum amount of Extender A 1:1. 7. Determine the concentration of the raw semen. Based on this concentration, determine how much additional extender (Extender A) must be added to make the concentration 200 to 300 million sperm/ml. Slowly add the necessary amount of Extender fraction A to achieve the desired concentration. 8. Place extended semen and Extender B in an ice water bath for 30 min; then slowly add an equal volume of Extender B to Extender A (a ratio of 1:1). 9. Incubate the glycerolated semen at 3°C (ice-water bath) for 1 hour. 10. Fill straws with semen, minimizing exposure to the warm air, seal, and then place back into ice-water bath until all straws are filled. Float Styrofoam platform in liquid nitrogen. Dry and load straws on freezing rack and place on floating lid. Straws should be approximately 8 cm above liquid nitrogen. After 10 min, plunge into liquid nitrogen.
temperatures, cooling rates, alternative freezing methods (straws vs. pellets), freezing curves, varying thaw temperatures, and effects of cryopreservation on acrosomal and/or plasma membrane integrity (Pursel and Park, 1985; Pontbriand et al., 1989; Bwanga, 1991; Pickett et al., 1992; Curry, 2000; Holt, 2000). Recently, Yoshioka et al. (2000) evaluated the effects of extender composition on post-thaw motility when Pacific white-sided and bottlenose dolphin sperm were frozen using the pelleting method described by Schroeder and Keller (1990). They found that the non-sugar-based extenders (egg yolk citrate) resulted in significant increases in post-thaw motility in both species. Durrant et al. (2000) provided the first descriptions of the effects of different freezing rates, incubation times with glycerol either prior to or after cooling, and freezing with or without cooling below room temperature. Their most effective freezing protocol was a medium freezing rate (12.8°C/min) with glycerol (4%) added prior to freezing after a 30 min cool to 4°C. They also illustrate the importance of comparing postfreezing motility score values to prefreezing values as a percentage. This eliminates the effect that differences in ejaculate quality between and within animals can have on post-thaw motility. Ongoing work with killer whale and Pacific white-sided dolphin semen indicates that they can be frozen successfully using straws. A simple method being developed for killer whales that has also been successful in Pacific white-sided dolphins is outlined in Table 3. It must be remembered when applying this method to a novel species that freezing curves and the most effective extender will vary with each species. Table 3 provides only a beginning. Postthaw motility as high as 70% in both species has been recorded (Robeck, unpubl. data). Ongoing research in Japan has shown high post-thaw motility when pelleting semen from
0839_frame_C11 Page 221 Tuesday, May 22, 2001 11:09 AM
Reproduction
221
Pacific white-sided dolphins (Yoshioka et al., 2000). Pelleting dolphin semen has a high rate of success, and methods have been discussed above, with details provided in the previous references. No methods have been published, however, that detail freezing cetacean semen in straws. Epididymal spermatozoa remain structurally intact, retaining motility in the tail of the epididymis for hours after death (Hopkins et al., 1988; Marmar, 1998). Successful collection and storage of post-mortem epididymal spermatozoa has been accomplished in a few species (Howard et al., 1986; Hopkins et al., 1988; Goodrowe and Hay, 1993). The concentration and motility of the spermatozoa vary with species, health of the animal before its death (traumatic event vs. chronic debilitation), length of time after death it is collected, environmental conditions at death, and handling of gonads once collected. Cornell and Leibo (pers. comm.) were able to collect and cryopreserve epididymal spermatozoa with 10% motility 24 hours post-mortem from a male bottlenose dolphin. After 72 hours at 4°C in Test-Y (Graham et al., 1972) extender they cryopreserved (at −10°C/min) four straws of semen in Test-Y and 10% glycerol. After 10 min at −196°C, they thawed (250°C/min) and evaluated the semen. The thawed semen had a 3 to 5% post-thaw motility. The ability to collect spermatozoa from wild or captive animals that die incidentally could provide managers another method to store and judiciously to infuse genetic material into captive populations (Wildt, 1989; Wildt et al., 1997; Kraemer, 2000). Manipulation and Control of Ovulation
Populations of bottlenose dolphins tend to exhibit seasonally bimodal peaks of reproductive activity or calf production. However, individual animals within these populations can be polyestrus throughout the year, anestrous, or pseudopregnant. Attempts to maximize the reproductive potential of these populations are difficult when potential breeding females are experiencing anestrus or pseudopregnancy. In addition, unpredictable estrous patterns reduce reproductive managers’ control of potential breeding events. Two basic methods of controlling ovulation in any mammalian species include induction of ovulation and estrus synchronization. Induction of Ovulation
Multiple attempts to induce ovulation in dolphins with exogenous gonadotropins have been performed with wide variations in response (Sawyer-Steffan et al., 1983; Schroeder and Keller, 1990). Because success was defined as elevated serum progesterone concentrations posttreatment, the authors were unable to determine if elevated progesterone reflected normal postovulatory luteinization. Similar doses of exogenous gonadotropins in other species commonly result in multiple ovulation, follicular luteinization, or other ovarian abnormalities (Hansel, 1985; Sreenan, 1988). In an effort to determine whether induced ovulation was normal, Schroeder and Keller (1990) allowed a reproductively successful male dolphin access to five exogenously induced females. Although breeding activity was observed, none of the females became pregnant. Two animals in this group were diagnosed as having persistent CLs. The lack of postinduction pregnancy after natural insemination and ovarian abnormalities (persistent CLs) provides strong evidence that these protocols were not effective. Robeck et al. (1998) used transabdominal ultrasonography to evaluate the response of bottlenose dolphins to ovulation-induction protocols. The results indicated that (1) bottlenose dolphins can be sensitive to exogenous gonadotropins, as multiple follicular recruitment of follicles occurred; (2) no physical evidence of ovulation was detected, but if ovulation were to occur, there was a good potential for multiple ovulations; and (3) until further ultrasonographic studies can be conducted to evaluate the effects of titrated doses of exogenous gonadotropins, induction protocols should be considered unsuitable for AI procedures in bottlenose dolphins.
0839_frame_C11 Page 222 Tuesday, May 22, 2001 11:09 AM
222
CRC Handbook of Marine Mammal Medicine
Robeck et al. (2000) attempted ovulation induction with three additional animals. Prior to exogenous gonadotropin administration, however, ultrasound was used to classify the dolphins as either anestrus or cycling. Cycling animals had follicles >5 mm (Brook, 1997). All three animals were placed on altrenogest at 1.5 ml/50 kg (110 lb), PO (oral), SID, for 16 days. After a single dose of 1500 IU of PG600® (Intervet America, Inc., Millsboro, DE) IM and 17.6 mg FSH intramuscularly (IM) on day 14, animal 2 and 3 responded with increased cortical activity, or antral follicle activity. After a second dose of 1800 IU PG600 was administered on day 22, animal 2 exhibited no ovarian change, and animal 3 had grown three follicles >25 mm. Animal 3 was administered two doses of 100 µg of GnRH (Cystorelin®, Merial Ltd, Harlow, Essex, U.K.) 10 days apart. GnRH administration did not stimulate ovulation despite the presence of follicles similar in size to preovulatory follicles previously characterized for T. t. aduncus (Brook, 1997). This may indicate that either the follicle was not preovulatory and/or that the dose of GnRH was ineffective. Administration of GnRH to animals that have nonpreovulatory follicles usually results in luteinization (Hennington et al., 1982; Valle et al., 1986). However, with this animal neither luteinization nor ovulation occurred, which probably indicated an insufficient dose of GnRH. These attempts at inducing ovulation in dolphins indicate that further investigations are needed to evaluate the differential sensitivity of the dolphin hypothalamic–pituitary–ovarian (HPO) axis to exogenous gonadotropins during anestrus or estrus, and at different stages of follicular growth. Synchronization of Ovulation
Attempts at synchronizing ovulation are most effective when used with normal, cycling animals. The ovarian response to exogenous hormones is variable both among and within species. In domestic species, the most effective methodologies use progestagens (Davis et al., 1979; Squires et al., 1979; Wright and Malmo, 1992) with or without estrogens (estradiol valerate) (Heersche et al., 1979; Odde, 1990) and/or prostaglandin F2α (Bunch et al., 1977; King et al., 1982; Odde, 1990). In some domestic species synthetic or natural prostaglandin F2α are commonly used in ovulatory synchronization protocols because of their luteolytic effect on receptive CL (between day 5 and 15 of the estrous cycle in cattle) (King et al., 1982). The many methodologies employed for estrus synchronization in various species are beyond the scope of this chapter (for review, see Wright, 1981; Odde, 1990; Wright and Malmo, 1992). Recently, the oral progestin altrenogest (Regu-Mate) has been evaluated as an estrus (ovulatory) synchronization tool in killer whales and bottlenose dolphins (Robeck, 2000; Robeck et al., 2000). In these studies, three dolphins and two killer whales were placed on Regu-Mate for as long as 31 days. Both of the killer whales and one of the dolphins were cycling prior to administration of the hormone. The time from progesterone withdrawal to estrus in the dolphins and killer whales was a mean 17.6 and 21.3 days, respectively. In both dolphins and killer whales, Regu-Mate appeared to cause a delay or suppression of ovarian activity after the hormone was withdrawn. The mean length of this suppression appeared to be similar to the length of the animals’ normal luteal phase. After this interval was reached, folliculogenesis and ovulation often occurred. All three dolphins placed on Regu-Mate returned to estrus within 1 week of each other. Although this interval is prolonged and too variable compared with traditional estrus-synchronization methods, it was effective for coordinating ovulation in a group of females during intensive AI trials. Receptivity of the cetacean CL to luteolytic doses of prostaglandin F2α is currently under investigation. Thus far, limited data indicate that PGF2α can be effective at disrupting normal CL function (Robeck et al., 2000). Three sonographically diagnosed nonpregnant animals with persistently elevated progesterone were administered an initial dose of 25 mg Lutalyse® (Pharmacia
0839_frame_C11 Page 223 Tuesday, May 22, 2001 11:09 AM
Reproduction
223
& Upjohn Co., Peapack, NJ) SID or BID (twice a day). Serum progesterone was determined 1 week after the initial dose. Two animals responded after the initial dose; the other one had to be given two additional doses of 25 mg Lutalyse 6 hours apart. Two of the animals went on to cycle normally, and have since become pregnant. Side effects of PgF2α administration generally consisted of apparent abdominal discomfort, nausea, and, on two occasions, inappetence for the remainder of the day. All obvious abdominal discomfort was gone within 1 hour, and all animals returned to normal behavior by the following day. The data suggest that nonpregnant animals with a history of elevated progesterone (>3 ng/ml) should be considered candidates for prostaglandin treatment. The results also demonstrate that these hormones can be administered safely. Further research is required to determine when and if a CL of diestrus is sensitive to exogenous prostaglandin F2α and what effect it will have on subsequent cycles. Insemination Techniques
Schroeder and Keller (1990) attempted to artificially inseminate five bottlenose dolphins in conjunction with the ovulation-induction protocol described above. For the procedure, freshly collected semen was placed external to the cervix in the spermathecal recess of the female using a flexible fiber optic laryngoscope (Schroeder, 1985; 1990; Schroeder and Keller, 1990). Based on serum progesterone levels, two of the five artificially inseminated animals were diagnosed as pregnant. Both pregnancies were believed to have spontaneously terminated in the first trimester. As was mentioned above, recent evidence using ultrasound indicates that when dolphins respond to exogenous gonadotropins, they do so with multiple follicular development. This increases the likelihood of multiple ovulation. Obviously, multiple ovulation, and potentially multiple embryos, would not be advantageous. Thus, without further research, ovulationinduction trials should be considered inappropriate for artificial or natural breeding. Recently, AI using cooled, transported semen and fresh, extended semen has been successful in the killer whale and the bottlenose dolphin, respectively (Robeck, unpubl. data; Brook, unpubl. data) (Figure 5). Each method used different indicators for determining when insemination should occur in relationship to ovulation. With the killer whale, urinary endocrine data were used to determine when the preovulatory estrogen surge had occurred. How this hormonal event relates to ovulation has yet to be determined, but ongoing ultrasonographic examinations should help determine this association. With bottlenose dolphins, ultrasonographic follicular evaluation was used to estimate the time of ovulation (Figure 6). Both
FIGURE 5 A 72-day-old T. t. aduncus fetus that was conceived through artificial insemination using fresh extended semen from a male located on site. The white arrows represent the CL of pregnancy. ac = amniotic cavity. (From F. Brook et al., unpubl. data.)
0839_frame_C11 Page 224 Tuesday, May 22, 2001 11:09 AM
224
CRC Handbook of Marine Mammal Medicine
FIGURE 6 Pre- and postovulation in a female T. t. aduncus. The sonogram on the left shows a 2-cm preovulatory follicle. The sonogram on the right was taken 12 hours after the one on the left. Fluid can still be seen in the recently ovulated follicle (black arrow). White arrows indicate ovarian dimensions. (From F. Brook, unpubl. data.)
techniques require intensive monitoring and have their limitations. Endocrine data require that (1) the animal be trained for urine collection; (2) an assay system be validated for the species in question; (3) the assay be rapid and provide results twice daily; and (4) the animal should have extensive hormonal profiling prior to inseminations. This profiling will help the manager predict when the animal will return to estrus and how long, generally, the animal will be in estrus before the estrogen surge occurs. Similar intensive animal monitoring is required when relying on ultrasonography. For this procedure to be effective (1) animals must be trained for regular voluntary sonogram exams or be restrained for the procedure; (2) there must be an ultrasound unit of minimal quality on site; and (3) the normal range of preovulatory follicular size for each animal should be determined. Future Applications
Kraemer (2000) and Wildt (2000) give good descriptions of current reproductive biotechnology and its realistic applications in exotic species. The short-term applications of biotechnology revolve around AI. The use of these technologies takes on more significance for long-term genetic management as the procedures become more refined, and can be applied in many different situations. Refinements of AI sophistication include successful insemination with fresh extended, cooled, transported, frozen, post-mortem epididymal, and sexed semen. The only successful method of AI in bottlenose dolphins relied on the most basic form. This involves collecting semen from the male on site (the male is usually in a different holding pool than the female), extending the semen to help protect and provide nutrients to maintain viability, and inseminating within a few hours of collection. Although this method has limited application for marine mammals as a whole, it enables park managers to house mature males and females separately. This type of social arrangement is most often observed in wild populations, and may have other benefits for population management (Wells, 2000). The second level of improvement for AI technologies is the use of cooled transported semen. This method, which was recently validated in killer whales, opens the door for
0839_frame_C11 Page 225 Tuesday, May 22, 2001 11:09 AM
Reproduction
225
meaningful application of AI to the marine mammal community (Robeck, unpubl. data). This method is so effective that the equine AI industry has been built around its use (Samper, 1997). This method involves collection, extension, and shipping of semen to an off-site location. Most shipping systems are designed to cool the semen in transit to provide longer periods of viability. Once on location, the cooled semen is deposited into the female that is approaching ovulation. For this system to work, managers must be able to predict when the female will ovulate within a couple of days, be able to collect semen routinely from the donor male, and develop extenders that will allow semen to remain viable during, and for at least 3 days after, the cooling process. The next area of progress in AI technology is in the successful use of cryopreserved semen. Developing this methodology requires tremendous effort to develop a system that will allow managers to cryopreserve semen with minimal loss in its fertility. Although most research with cryopreserved semen uses post-thaw motility and membrane integrity to evaluate success of the procedure, the ultimate and often only meaningful test is to determine its fertility. Once it can be demonstrated that frozen semen can be used to inseminate a marine mammal successfully, the door will be opened for long-term genetic management. This technique will allow shipment of semen across international borders, thus effectively opening up the captive gene pool to worldwide contributions. It will also allow long-term storage of valuable genetic material that can be selectively reintroduced into the gene pool generations after the donor is deceased. It may also allow the use of cryopreserved semen collected postmortem from the epididymis of stranded delphinids. Harvesting of genetic material from animals that would otherwise be lost to wild populations would greatly increase the genetic diversity of captive populations without the need for additional wild live captures. This technique has already been used successfully in domestic and exotic terrestrial animals and has been applied for in vitro fertilization techniques in minke whales (Balaenoptera acutorostrata) (Fukui et al., 1997a,b). Finally, use of sex-determined semen (sorted by chromosomal content) for successful AI in marine mammals would revolutionize animal management procedures. The ability to control sex ratios would allow optimum utilization of the limited resources available to managed species. Although in its infancy, this technique uses flow cytometry to sort semen based on nuclear content or X- vs. Y-bearing spermatozoa. The current sorting rates are around 900 live sperm/second; thus, the technique produces far fewer sperm than are required for transcervical inseminations. However, the technique has been used together with laparoscopic insemination in cattle to produce conception rates of 30% with liquid semen and 51% with frozen semen, and producing 97% females (Seidel et al., 1999). This technique appears to be a long way from being implemented in marine mammals; however, the recent abilities to perform laproscopic procedures (see Chapter 27, Endoscopy) and to monitor ovarian activity place this procedure within reach, although many challenges still remain before the sorting system is validated for delphinid semen.
Acknowledgments The authors thank Denise Greig for editorial assistance in preparing this chapter for publication. In addition, the authors acknowledge all those, far too numerous to mention individually, whose work has contributed to their increasing knowledge of reproduction in marine mammals and, in particular, those colleagues who have directly supported the authors’ efforts over the years. This article is SeaWorld San Antonio technical contribution no. 2000-05-T.
0839_frame_C11 Page 226 Tuesday, May 22, 2001 11:09 AM
226
CRC Handbook of Marine Mammal Medicine
References Amoroso, E.C., Bourne, G.H., Harrison, R.J., Harrison Matthews, L., Rowlands, I.W., and Sloper, J.C., 1965, Reproductive and endocrine organs of foetal, newborn and adult seals, J. Zool., 147: 430–486. Asa, C., 2000, Current methods of contraception in zoos, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 191–200. Asher, G.W., Day, A.M., and Barrell, G.K., 1987, Annual cycle of liveweight and reproductive changes of farmed male fallow deer (Dama dama) and the effect of daily administration of melatonin in summer on the attainment of seasonal fertility, J. Reprod. Fertil., 79: 353–362. Atkinson, S., 1997, Reproductive biology of seals, Rev. Reprod., 2: 175–194. Atkinson, S., and Gilmartin, W.G., 1992, Seasonal testosterone pattern in Hawaiian monk seals, J. Reprod. Fertil., 96: 35–39. Atkinson, S., Gilmartin, W.G., and Lasley, B.L., 1993, Testosterone response to a gonadotropin-releasing hormone agonist in Hawaiian monk seals (Monachus schauinslandi), J. Reprod. Fertil., 97: 35–38. Atkinson, S., Becker, B.L., Johanos, T.C., Pietraszek, J.R., and Kuhn, B.C.S., 1994, Reproductive morphology and status of female Hawaiian monk seals fatally injured by adult male seals, J. Reprod. Fertil., 100: 225–230. Atkinson, S., Ragen, T.J., Gilmartin, W.G., Becker, L., and Johanos, T.C., 1998, Use of GnRH agonist to suppress testosterone in wild male monk seals (Monachus schauinslandi), Mar. Mammal Sci., 112: 178–182. Atkinson, S., Combelles, C., Vincent, D., Nachtigall, P., Pawloski, J., and Breese, M., 1999, Monitoring of progesterone in captive female false killer whales, Pseudorca crassidens, Gen. Comp. Endocrinol., 115: 323–332. Baird, D.T., and Smith, K.B., 1993, Inhibin and related peptides in the regulation of reproduction, in Oxford Review of Reproductive Biology, Milligan, S.R. (Ed.), Oxford University Press, Oxford, U.K., 191–232. Balcomb, K.C.I., Boran, J.R., and Heimlich, S.L., 1982, Killer whales in greater Puget Sound––A population ideally suited for statistical modeling, Rep. Int. Whaling Comm., 32: 681–686. Baldwin, R.L., and Miller, P.S., 1991, Mammary gland development and lactation, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, San Diego, CA, 385–412. Belanger Auclair, C., Ferland, L., Caron, S., and Labrie, F., 1980, Time-course of the effect of treatment with a potent LHRH agonist on testicular steroidogenesis and gonadotropin receptor levels in the adult rat, J. Steroid Biochem., 13: 191–196. Benirschke, K., Johnson, M.L., and Benirschke, R.J., 1980, Is ovulation in dolphins, Stenella longirostris and Stenella attenuata always copulation induced? Fish. Bull., 78: 507–528. Bigg, M.A., 1969, Clines in the pupping season of the harbor seal, Phoca vitulina, J. Fish. Res. Board Can., 26: 449–455. Bigg, M.A., 1973, Adaptations in the breeding of the harbour seal, Phoca vitulina, J. Reprod. Fertil. Suppl., 19: 131–142. Bigg, M.A., 1982, An assessment of killer whale (Orcinus orca) stocks off Vancouver Island, British Columbia, Rep. Int. Whaling Comm., 32: 655–666. Blue, B.J., Pickett, B.W., Squires, E.L., McKinnon, A.O., Nett, T.M., Amann, R.P., and Shiner, K.A., 1991, Effect of pulsatile or continuous administration of GnRH on reproductive function of stallions, J. Reprod. Fertil. Suppl., 44: 145–154. Boyd, I.L., 1983, Luteal regression, follicle growth and the concentration of some plasma steroids during lactation in grey seals (Halichoerus grypus), J. Reprod. Fertil., 69: 157–164. Boyd, I.L., 1991, Changes in plasma progesterone and prolactin concentrations during the annual cycle and the role of prolactin in the maintenance of lactation and luteal development in the Antarctic fur seal (Arctocephalus gazella), J. Reprod. Fertil., 91: 637–647.
0839_frame_C11 Page 227 Tuesday, May 22, 2001 11:09 AM
Reproduction
227
Boyd, I.L., Lockyer, C., and Marsh, H., 1999, Reproduction in marine mammals, in Biology of Marine Mammals, Reynolds, J.E., III, and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 218–286. Braham, H.W., 1984, Review of reproduction in the white whale, Delphinapterus leucas, narwhale, Monodon monoceros, and irrawaddy dolphin, Orcaella brevirostris, with comments on stock assessment, in Reproduction in Whales, Dolphins and Porpoises, Special Issue 6, Perrin, W.F., Brownell, R.J., Jr., and DeMaster, D.P. (Eds.), International Whaling Commission, Cambridge, U.K., 6: 81–90. Briggs, M., 2000, Contraception in bottlenose dolphins (Tursiops truncatus), in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 201–204. Brodie, P.F., 1971a, A reconsideration of aspects of growth, reproduction and behavior of the white whale (Delphinapterus leucus) with reference to the Cumberland Sound, Baffin Island, population, J. Fish. Res. Board Can., 28: 1309–1318. Brodie, P.F., 1971b, A reconsideration of aspects of growth, reproduction, and behavior of the white whale (Delphinapterus leucas), with reference to the Cumberland Sound, Baffin Island, population, J. Fish. Res. Board Can., 28: 1309–1319. Bronson, F.H., 1988, Seasonal regulation of reproduction in mammals, in The Physiology of Reproduction, Knobil, E., Neill, J., Ewing, L.L., Greenwald, G.S., Markert, C.L., and Pfaff, D.W. (Eds.), Raven Press, New York, 2323–2351. Brook, F., 1994, Ultrasound diagnosis of anencephaly in the fetus of a bottlenose dolphin (Tursiops aduncas), J. Zoo Wildl. Med., 25: 569–574. Brook, F., 1997, The Use of Diagnostic Ultrasound in Assessment of the Reproductive Status of the Bottlenose Dolphin, Tursiops aduncas, in Captivity and Applications in Management of a Controlled Breeding Programme, Ph.D. thesis, Hong Kong Polytechnic University, Kowloon, Hong Kong, 339 pp. Brook, F., 2000, Sonographic testicular and ovarian assessment in the bottlenose dolphin, Tursiops truncatus aduncus, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 207–222. Brook, F.M., Kinoshita, R., Brown, B., and Metreweli, C., 2000, Ultrasonographic imaging of the testis and epididymis of the bottlenose dolphin, Tursiops truncatus aduncus, J. Reprod. Fertil., 119: 233–240. Brown, J.L., Schoenemann, H.M., and Reeves, J.J., 1986, Effect of FSH treatment on LH and FSH receptors in chronic cystic-ovarian-diseased dairy cows, J. Anim. Sci., 62: 1063–1071. Brown, J.L., Citino, S.B., Bush, M., Lehnhardt, J., and Phillips, L., 1991, Cyclic patterns of luteinizing hormone, follicle-stimulating hormone, inhibin, and progesterone secretion in the Asian elephant (Elephas maximus), J. Zoo Wildl. Med., 22: 49–57. Brown, R.G., Bowen, W.D., Eddington, J.D., Kimmins, W.C., Mezei, M., Parsons, J.L., and Pohajdak, B., 1997a, Temporal trends in antibody production in captive grey, harp and hooded seals to a single administration immunocontraceptive vaccine, J. Reprod. Immunol., 35: 53–64. Brown, R.G., Bowen, W.D., Eddington, J.D., Kimmins, W.C., Mezei, M., Parsons, J.L., and Pohajdak, B., 1997b, Evidence for a long-lasting single administration contraception vaccine in wild grey seals, J. Reprod. Immunol., 35: 43–51. Bryden, M.M., and Harrison, R.J., 1986, Gonads and reproduction, in Research on Dolphins, Bryden, M.M., and Harrison, R. (Eds.), Clarendon Press, Oxford, U.K., 149–159. Bunch, T.D., Foote, W.C., and Whitaker, B., 1977, Interspecies ovum transfer to propagate wild sheep, J. Wildl. Manage., 41: 726–730. Bwanga, C.O., 1991, Cryopreservation of boar semen II: Effects of cooling rate and duration of freezing point plateau on boar semen frozen in mini- and maxi-straws and plastic bags, Acta Vet. Scand., 32: 455–461.
0839_frame_C11 Page 228 Tuesday, May 22, 2001 11:09 AM
228
CRC Handbook of Marine Mammal Medicine
Byers, S.W., Dowsett, K.F., and Glover, T.D., 1983, Seasonal and circadian changes of testosterone levels in the peripheral blood plasma of stallions and their relation to semen quality, J. Endocrinol., 99: 141–150. Calle, P.P., Cook, R.A., Robeck, T.R., Young, S.J.F., and Jones, M.H., 1996, Circulating gestational progesterone and estradiol concentrations in beluga whales (Delphinapterus leucas), Proc. Am. Assoc. Zoo Vet., 340–341. Calle, P.P., Monfort, S.L., Dunn, J.L., Jensen, E., Boehm, R., Young, S.J.F., and Robeck, T.R., 2000, Seasonal testosterone secretion in male white whales (Delphinapterus leucas), in Proceedings of American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine Joint Meeting, 49–50. Carriere, P.D., Amaya, D., and Lee, B., 1995, Ultrasonography and endocrinology of ovarian dysfunctions induced in heifers with estradiol valerate, Theriogenology, 43: 1061–1076. Catchpole, H.R., 1991, Hormonal mechanisms in pregnancy and parturition, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, New York, 361–380. Chrisp, P., and Goa, K.L., 1990, Naferlin: A review of its pharmacodynamic and pharmacokinetic properties, and clinical potential in sex hormone-related conditions, Drugs, 39: 523–551. Christensen, I., 1984, Growth and reproduction of killer whales, Orcinus orca, in Norwegian coastal waters, in Reproduction in Whales, Dolphins and Porpoises, Special Issue 6, Perrin, W.F., Brownell, R.L., and DeMaster, D.P. (Eds.), International Whaling Commission, Cambridge, 6: 253–258. Cockcroft, V.G., and Ross, G.J.B., 1990, Age growth, and reproduction of bottlenose dolphins Tursiops truncatus from the east coast of southern Africa, Fish. Bull., 88: 289–302. Comrie, L.C., and Adams, A.B., 1938, The female reproductive system and corpora lutea of the false killer whale, Pseudorca crassidens, Trans. R. Soc. Edinburgh, 59: 521–531. Conn, P.M., McArdle, C.A., Andrews, W.V., and Huckle, W.R., 1987, The molecular basis of gonadotropinreleasing hormone (GnRH) action in the pituitary gonadotrope, Biol. Reprod., 36: 17–35. Conn, P.M., Huckle, W.R., Andrews, W.V., and McCardle, C.A., 1988, The molecular mechanism of action of gonadotropin releasing hormone (GnRH) in the pituitary, Recent Prog. Horm. Res., 43: 29–61. Cowan, D., 2000, Pathologic evidence of reproductive disease and/or dysfunction in wild cetaceans, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 163–174. Craig, A.M., 1964, Histology of reproduction and the estrous cycle in the female fur seal, Callorhinus ursinus, J. Fish. Res. Board Can., 21: 773–811. Cunningham, G.R., and Huckins, C., 1979, Persistence of complete spermatogenesis in the presence of low intra-testicular concentrations of testosterone, Endocrinology, 105: 177–184. Cupps, P.T., 1991, Reproduction in Domestic Animals, 4th ed., Academic Press, New York. Curry, M.R., 2000, Cryopreservation of semen from domestic livestock, Rev. Reprod., 5: 46–52. Dalton, L.M., Robeck, T.R., Calle, P.P., and Cook, R.A., 1994, Observations at parturition in eight beluga whales (Delphinapterus leucas), in Proceedings of the Annual International Association for Aquatic Animal Medicine Conference, 1–2. Dalton, L.M., Robeck, T.R., and Young, W.G., 1995, Growth and development of a Pacific white-sided dolphin (Legemorhynchus obliquidens), in Proceedings of the Annual International Association for Aquatic Animal Medicine Conference, 89–91. Daniel, J.C., 1971, Growth of the preimplantation embryo of the northern fur seal and its correlation with changes in uterine protein, Dev. Biol., 26: 316–322. Daniel, J.C., 1974, Circulating levels of oestradiol-17B during early pregnancy in the Alaskan fur seal showing an oestrogen surge preceding implantation, J. Reprod. Fertil., 37: 425–428. Davis, D.L., Knight, J.W., Killian, D.B., and Day, B.N., 1979, Control of estrus in gilts with a progestogen, J. Anim. Sci., 49: 1507–1509.
0839_frame_C11 Page 229 Tuesday, May 22, 2001 11:09 AM
Reproduction
229
Dougherty, M., Bossart, G., and Renner, M., 2000, Case study: Calf born thirteen months post-ReguMate contraception with social complications, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 41–42. Duffield, D.A., and Wells, R.S., 1991, The combined application of chromosome, protein and molecular data for the investigation of social unit structure and dynamics in Tursiops truncatus, in Genetic Ecology of Whales and Dolphins, Hoelzel, A.R. (Ed.), Report of the International Whaling Commission, Special Issue 13, Cambridge, U.K., 155–169. Duffield, D.A., Odell, D.K., McBain, J.F., and Andrews, B., 1995, Killer whale (Orcinus orca) reproduction at SeaWorld, Zoo Biol., 14: 417–430. Duffield, D.A., Shell, D., and Dudley, M., 2000, Demographic analysis of breeding bottlenose dolphins in North American zoological facilities: 1976–1998, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 139–156. Durrant, B.S., Russ, K.D., Proulx, J.R., Reddy, M.L., and Ridgway, S.H., 2000, Current techniques for semen cryopreservation in bottlenose dolphins (Tursiops truncatus), in Report from the Bottlenose Dolphin Reproductive Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), June 3–6, San Diego, CA, American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 239–248. Fay, F.H., 1981, Walrus, Odobenus rosmarus, in Handbook of Marine Mammals, Vol. 1: The Walrus, Sea Lions, Fur Seals and Sea Otter, Ridgway, S.H., and Harrison, R.J. (Eds.), Academic Press, New York, 1–24. Fay, F.H., Ray, G.C., and Kibal’chich, A.A., 1981, Time and location of mating and associated behavior of the Pacific walrus, Odobenus rosmarus divergens Illiger, in Soviet-American Cooperative Research on Marine Mammals, Vol. 1, Pinnipeds, Fay, F.H., and Fedoseev, G.A. (Eds.), NOAA Technical Report NMFS 12: 89–99. Fisher, H.D., and Harrison, R.J., 1970, Reproduction in the common porpoise (Phocoena phocoena) of the North Atlantic, J. Zool. Soc. London, 161: 471–486. Fleming, A.D., Yanagimachi, R., and Yanagimachi, H., 1981, Spermatozoa of the Atlantic bottlenosed dolphin, Tursiops truncatus, J. Reprod. Fertil., 63: 509–514. Frick, J., Bartsch, G., and Weiske, W.M., 1977, The effects of monthly depot medroxy progesterone acetate and testosterone on human spermatogenesis uniform dosage levels, Contraception, 15: 649–655. Fukui, Y., Mogoe, T., Ishikawa, H., and Ohsumi, S., 1997a, Factors affecting in vitro maturation of minke whale (Balaenoptera acutorostrata) follicular oocytes, Biol. Reprod., 56: 523–528. Fukui, Y., Mogoe, T., Ishikawa, H., and Ohsumi, S., 1997b, In vitro fertilization of in vitro matured minke whale (Balaenoptera acutorostrata) follicular oocytes, Mar. Mammal Sci., 13: 395–404. Ganong, W.F., 1991, Role of the nervous system in reproduction, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, San Diego, CA, 1–23. Gardiner, K.J., Boyd, L., Follett, B.K., Racey, A., and Reijnders, P.J., 1999, Changes in pituitary, ovarian, and testicular activity in harbour seals (Phoca vitulina) in relation to season and sexual maturity, Can. J. Zool., 77: 211–221. Gaskin, D.E., Smith, G.J.D., Watson, A.P., Yasui, W.Y., and Yurick, D.B., 1984, Reproduction in the porpoises (Phocoenidae): Implications for management, in Reproduction in Whales, Dolphins and Porpoises, Perrin, W.F., Brownell, R.L., and DeMaster, D.P. (Eds.), International Whaling Commission, Cambridge, U.K., 135–148. Gilmartin, W.G., DeLong, R.L., Smith, A.W., Sweeney, J.C., DeLappe, B.W., Riseborough, R.W., Griner, L.A., Dailey, M.D., and Peakall, D.B., 1976, Premature parturition in the California sea lion, J. Wildl. Dis., 12: 104–115. Goodman, R.L., 1988, Neuroendocrine control of the ovine estrous cycle, in The Physiology of Reproduction, Vol. 2, Knobil, E., and Neill, J.D. (Eds.), Raven Press, New York, 1929–1969.
0839_frame_C11 Page 230 Tuesday, May 22, 2001 11:09 AM
230
CRC Handbook of Marine Mammal Medicine
Goodrowe, K.L., and Hay, M., 1993, Characteristics and zona binding ability of fresh and cooled domestic cat epididymal spermatozoa, Theriogenology, 40: 967–975. Graham, E.F., Crabo, B.G., and Brown, K.I., 1972, Effect of some zwitter ion buffers on the freezing and storage of spermatozoa I. Bull, J. Dairy Sci., 55: 372–378. Griffiths, D.J., 1984a, The annual cycle of the testis of the elephant seal (Mirounga leonina) at Macquarie Island, J. Zool. Soc. London, 203: 193–204. Griffiths, D.J., 1984b, The annual cycle of the epididymis of the elephant seal (Mirounga leonina) at Macquarie Island, J. Zool., 203: 181–191. 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-8, National Marine Fisheries Service, U.S. Department of Commerce, Silver Spring, MD, 44 pp. Hansel, W., 1985, Advances in physiology of growth, reproduction and lactation, Cornell Vet., 75: 56–76. Harrison, R.J., 1968, General physiology of the pinnipedia, in The Behavior and Physiology of Pinnipeds, Harrison, R.J., Hubbard, R.S., Peterson, C.E., and Schusterman, R.J. (Eds.), Appleton-CenturyCrofts, New York, 211–296. Harrison, R.J., 1969, Reproduction and reproductive organs, in The Biology of Marine Mammals, Anderson, H.T. (Ed.), Academic Press, New York, 253–348. Harrison, R.J., and Ridgway, S.H., 1971, Gonadal activity in some bottlenose dolphins (Tursiops truncatus), J. Zool., 165: 355–366. Harrison, R.J., Brownell, R.J., Jr., and Boice, R.C., 1972, Reproduction and gonadal appearances in some odontocetes, in Functional Anatomy of Marine Mammals, Vol. 1, Harrison, R.J. (Ed.), Academic Press, New York, 361–469. Heath, R.B., 1985, The effects of environment on the breeding system of the California sea lion (Zalophus californianus), in Proceedings of the Sixth Biennial Conference on the Biology of Marine Mammals, Vancouver, British Columbia, Canada. Heersche, G., Kirakofe, G.H., DeBenedetti, R.C., Wen, S., and McKee, R.M., 1979, Synchronization of estrus in beef heifers with a norgestomet implant and prostaglandin F2a, Theriogenology, 11: 197–103. Hendricks, D.M., 1991, Biochemistry and physiology of the gonadal hormones, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, San Diego, CA, 81–118. Hennington, D.L., Kreider, J.L., and Potter, G.D., 1982, The effects of GnRH on induction of follicular development and ovulation in anovulatory and ovulatory mares, Theriogenology, 17: 635–643. Hill, H.J., and Gilmartin, W.G., 1977, Collection and storage of semen from dolphins, in Breeding Dolphins: Present Status, Suggestions for the Future, Ridgway, S.H., and Benirschke, K. (Eds.), U.S. Department of Commerce, NTIS PB-273-673, Washington, D.C., 205–210. Hinrichs, K., 1977, Irregularities of the estrous cycle in mares, in Current Therapy in Large Animal Theriogenology, Youngquist, R.S. (Ed.), W.B. Saunders, Philadelphia, 166–171. Hobson, B.M., and Boyd, I.L., 1984, Gonadotropin and progesterone concentrations in placentae of grey seals (Halichoerus grypus), J. Reprod. Fertil., 72: 521–528. Hobson, M., and Wide, L., 1986, Gonadotrophin in the term placenta of the dolphin (Tursiops truncatus), the California sea lion (Zalophus californianus), the grey seal (Halichoerus grypus) and man, J. Reprod. Fertil., 76: 637–644. Holt, W.V., 2000, Fundamental aspects of sperm cryobiology: The importance of species and individual differences, Theriogenology, 53: 47–58. Hopkins, S.M., Armstrong, D.L., Hummel, S.K.C., and Junior, S., 1988, Successful cryopreservation of gaur (Bos gaurus) epididymal spermatozoa, J. Zoo Wildl. Med., 19: 195–201. Howard, J.G., Bush, M., and Wildt, D.E., 1986, Semen collection, analysis and cryopreservation in nondomestic mammals, in Current Therapy in Theriogenology 2, Morrow, D.A. (Ed.), W.B. Saunders, Philadelphia, 1047–1053.
0839_frame_C11 Page 231 Tuesday, May 22, 2001 11:09 AM
Reproduction
231
Howard, J.G., Bush, M., Morton, C., Morton, F., Wentzel, K., and Wildt, D.E., 1991, Comparative semen cryopreservation in ferrets (Mustela putorius furo) and pregnancies after laparoscopic intrauterine insemination with frozen-thawed spermatozoa, J. Reprod. Fertil., 92: 109–118. Iwasa, M., and Atkinson, S., 1997, Analysis of corpora lutea to estimate reproductive cycles of wild Hawaiian monk seals (Monachus schauinslandi), Mar. Mammal Sci., 12: 182–198. Iwasa, M., Atkinson, S., and Kamiya, S., 1997, Lipofuscin granular cells in regressing corpora lutea and corpora albicantia of ovaries from wild Hawaiian monk seals, Mar. Mammal Sci., 13: 326–332. Jensen, E., 2000, Embryonic/early fetal loss in the Atlantic bottlenose dolphin (Tursiops truncatus), in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 273–278. Joseph, B.E., Duffield, D.A., and Robeck, T.R., 2000, Summary data on reproduction of bottlenose dolphins in controlled environments, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 43–56. Katsumata, E., Katsumata, H., Tobayama, T., and Usuki, 1999a, Body temperature in reared killer whales, Orcinus orca: Useful predictive method of confinement in females? in Recent Progress in Molecular and Comparative Endocrinology Asia and Oceania Society of Comparative Endocrinology, Korea Kwangiu, Republic of Korea, 448–452. Katsumata, E., Katsumata, H., Tobayama, T., and Usuki, S., 1999b, Changes in serum steroid hormone levels in reared killer whales, Orcinus orca, in Recent Progress in Molecular and Comparative Endocrinology Asia and Oceania Society of Comparative Endocrinology, Korea Kwangiu, Republic of Korea, 116–122. Kenagy, G.J., and Trombulak, S.C., 1986, Size and function of mammalian testes, J. Mammal., 75: 1–22. King, M.E., Kiracofe, G.F., Stevenson, J.S., and Schalles, R.R., 1982, Effect of stage of the estrous cycle on interval to estrus after PGF2α in beef cattle, Theriogenology, 18: 191–197. Kirby, V.L., 1982, Testosterone levels in captive bottlenose dolphins, Naval Oceans Systems Center Contract Report, N66001-82-0098 J. Kirby, V.L., 1990, Endocrinology of marine mammals, in CRC Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 303–351. Kirby, V.L., and Ridgway, S.H., 1984, Hormonal evidence of spontaneous ovulation in captive dolphins (Tursiops truncatus and Delphinus delphis), Rep. Int. Whaling Comm., Spec. Issue 6: 459–464. Kirkpatrick, J.F., Turner, J.W., Jr., and Perkins, A., 1982, Reversible fertility control in feral horses, J. Equine Vet. Sci., 2: 114–118. Kirkpatrick, J.F., Liu, I.K.M., and Turner, J.W., Jr., 1990, Remotely-delivered immunocontraception in feral mares, Wildl. Soc. Bull., 18: 326–330. Kirkpatrick, J.F., Turner, J.W., Jr., Liu, I.K.M., and Fayrer-Hosken, R., 1996, Application of pig zona pellucida immunocontraception to wildlife fertility control, J. Reprod. Fertil., Suppl. 50: 183–189. Kiyota, M., Yamaguchi, Y., Nishikawa, F., and Kohyama, K., 1999, Cytological changes in vaginal smear and epithelium associated with the reproductive cycle in northern fur seal, Callorhinus ursinus, Bull. Nat. Resour. Inst. Far Seas Fish., 36: 17–25. Knobil, E., and Neill, J.D., 1988, The Physiology of Reproduction, Vol. 1 and 2, Raven Press, New York. Kraeling, R.R., Dziuk, P.J., Pursel, V.G., Rampacek, G.B., and Webel, S.K., 1981, Synchronization of estrus in swine with ally trenbolone (RU-2267), J. Anim. Sci., 52: 831–835. Kraemer, D.C., 2000, Current advances in reproductive biotechnology, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 95–100. Lasley, B.L., and Anderson, G.B., 1991, Where does biotechnology fit in captive breeding programs? Zoo Biol., 10: 195–196. Laws, R.M., 1955, The elephant seal (Mirounga leonina Linn.), III. The physiology of reproduction, Falkland Island Dependencies Surveys Scientific Reports, No. 15.
0839_frame_C11 Page 232 Tuesday, May 22, 2001 11:09 AM
232
CRC Handbook of Marine Mammal Medicine
Lindsay, D.R., 1991, Reproduction in sheep and goat, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, San Diego, CA, 491–512. Mahi-Brown, C.A., Yanagimachi, R., and Nelson, M.L., 1988, Ovarian histopathology of bitches immunized with porcine zonae pellucidae, Am. J. Reprod., Immunol. Microbiol., 18: 94. Mann, D.R., Gould, K.G., and Collins, D.C., 1984, Influences of continuous gonadotropin-releasing hormone (GnRH) agonist treatment on luteinizing hormone and testosterone secretion, the responce to GnRH, and the testicular response to human chorionic gonadotropin in male rhesus monkeys, J. Clin. Endocrinol., 58: 262–267. Mariana, C., Monniaux, M.A., Driancourt, M.A., and Mauleon, P., 1991, Folliculogenesis, in Reproduction in Domestic Animals, Cupps, P.T. (Ed.), Academic Press, San Diego, CA, 119–173. Marmar, J.L., 1998, The emergence of specialized procedures for the acquisition, processing and cryopreservation of epididymal and testicular sperm in connection with intracytoplasmic sperm injection, J. Androl., 19: 517–526. Marsh, H., and Kasuya, T., 1984, Changes in the ovaries of short-finned pilot whales, Globicephala macrorhynchus, off the Pacific coast of Japan, in Reproduction in Whales, Dolphins and Porpoises, Perrin, W.F., Brownell, R.L., and DeMaster, D.P. (Eds.), International Whaling Commission, Cambridge, U.K., 311–336. Matkin, C.O., and Leatherwood, S., 1986, General biology of killer whales, in Behavioral Biology of Killer Whales: Zoo Biology Monographs, Vol. 1, Kirkevold, B., and Lockard, J.S. (Eds.), Alan R. Liss, New York, 35–68. Matsubayashi, K., Watanabe, G., Taya, K., Katakai, Y., Sasamoto, S., Suzuki, J., and Nozaki, M., 1991, Seasonal changes in plasma concentrations of immunoreactive inhibin and testicular activity in male Japanese monkeys, Biol. Reprod., 44: 822–826. Matsue, H., Arimure, A., Nair, M.G., and Schally, A.V., 1971, Synthesis of the porcine LH and FSHreleasing hormone by solid-phase method, Biochem. Biophys. Res. Commun., 45: 822–827. Matsumoto, A.M., 1990, Effects of chronic testosterone administration in normal men: Safety and efficacy of high dosage testosterone and parallel dose-dependent suppression of LH, FSH and sperm production, J. Clin. Endocrinol. Metab., 70: 282–287. McNeilly, A.S., 1988, Suckling and the control of gonadotropin secretion, in The Physiology of Reproduction, Knobil, E., Neill, J., Ewing, L.L., Greenwald, G.S., Markert, C.L., and Pfaff, D.W. (Eds.), Raven Press, New York, 2323–2351. Miller, D., and Bossart, G., 2000, Reproductive and fatal neonatal diseases in cetaceans from Florida oceanarium: 1979–1999, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck,T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 183–190. Miller, E.H., Pitcher, K.W., and Loughlin, T.R., 1999, Bacular size, growth and allometry in the largest extant otariid, the Steller sea lion (Eumetopias jubatus), J. Mammal., 81: 134–144. Morejohn, G.V., 1975, A phylogeny of otariid seals based on morphology of the baculum, Rapp. P. V. Reun. Cons. Int. Explor. Mer, 169: 49–56. Nett, T.M., Crowder, M.E., Moss, G.E., and Duello, T.M., 1981, GnRH-receptor interaction. V. Down regulation of pituitary receptors for GnRH in ovariectomized ewes by infusion of homologous hormone, Biol. Reprod., 24: 1145–1155. Noonan, L.M., Ronald, K., and Raeside, J.I., 1991, Plasma testosterone concentrations of captive male hooded seals (Cytophora cristata), Can. J. Zool., 69: 2279–2282. Norman, A.W., and Litwack, G., 1987, Hormones, Academic Press, San Diego, CA, 565–605. Odde, K.G., 1990, A review of synchronization of estrus in postpartum cattle, J. Anim. Sci., 68: 817–830. Ohsumi, S., 1964, Comparison of maturity and accumulation rate of corpora albacantia between the left and right ovaries in cetacea, Tokyo Whale Res. Inst., 18: 123–149. Olesiuk, P.F., Bigg, M.A., and Ellis, G.M., 1990, Life history and population dynamics of resident killer whales (Orcinus orca) in the coastal waters of British Columbia and Washington State, Rep. Int. Whaling Comm., Spec. Issue 12: 209–243.
0839_frame_C11 Page 233 Tuesday, May 22, 2001 11:09 AM
Reproduction
233
Perrin, W.F., and Reilly, S.B., 1984, Reproductive parameters of dolphins and small whales of the family Delphinidae, Rep. Int. Whaling Comm., Spec. Issue 6: 97–134. Perrin, W.F., Brownell, R.L., and DeMaster, D.P. (Eds.), 1984, Reproduction in Whales, Dolphins and Porpoises, Rep. Int. Whaling Comm., Spec. Issue 6, 495 pp. Pickett, B.W., Berndtson, W.E., and Sullivan, J.J., 1992, Influence of seminal additives and packaging systems on fertility of frozen bovine spermatozoa, J. Anim. Sci., 70: 12–47. Pietraszek, J.R., and Atkinson, S., 1994, Concentrations of oestrone sulfate and progesterone in plasma and saliva, vaginal cytology, and bioelectric impedance during the oestrous cycle of the Hawaiian monk seal, Mar. Mammal Sci., 10: 430–441. Plant, T.M., 1988, Puberty in primates, in The Physiology of Reproduction, Vol. 2, Knobil, E., and Neill, J.D. (Eds.), Raven Press, New York, 1763–1788. Pontbriand, S.M., Howard, J.G., Schiewe, M.C., Stuart, L.D., and Wildt, D.E., 1989, Effect of cryoprotective diluent and method of freeze-thawing on survival and acrosomal integrity of ram spermatozoa, Cryobiology, 10: 255–263. Pursel, V.G., and Park, C.S., 1985, Freezing and thawing procedures for boar spermatozoa, in Deep Freezing of Boar Semen, Johnson, L.A., and Larsson, K. (Eds.), Swedish University of Agricultural Science, Uppsala, 147–166. Purves, P.E., and Pilleri, G., 1978, The functional anatomy and general biology of Pseudorca crassidens with a review of the hydrodynamics and acoustics in cetacea, Invest. Cetacea, 67: 227. Reijnders, P.J., 1986, Reproductive failure in harbour seals feeding on fish from polluted coastal waters, Nature, 324: 456–457. Reijnders, P.J.H., 1991, Progesterone and oestradiol-17β concentration profiles throughout the reproductive cycle in harbor seals (Phoca vitulina), J. Reprod. Fertil., 90: 403–409. Renfree, M.B., and Calaby, J.H., 1981, Background to delayed implantation and embryonic diapause, J. Reprod. Fertil., Suppl. 29: 1–9. Richkind, M., and Ridgway, S.H., 1975, Estrogens, corticosteroid and progestagen patterns in the pregnant and non-pregnant bottle-nosed dolphin Tursiops truncatus, following the intramuscular administration of nih-fsh-ovine-s9, J. Steroid Biochem., 15–20. Riedman, M., 1990, The Pinnipeds, Seals, Sea Lions, and Walrus, University of California Press, Berkeley, 222–263. Robeck, T.R., 1996, Defining Basic Reproductive Physiology, Including Reproductive Anatomy, Reproductive Endocrinology and Estrus Induction of Captive Delphinids, Dissertation, Texas A&M University, UMI Dissertation Service, 89–166. Robeck, T.R., 2000, Advances in the understanding and manipulation of bottlenose dolphin reproduction, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 109–132. Robeck, T.R., McBain, J.M., Dalton, L.M., Walsh, M.T., Czekala, M., and Kraemer, D.C., 1990, Monitoring reproductive endocrinology of captive killer whales, Orcinus orca, in Proceedings of the Annual International Association for Aquatic Animal Medicine Conference, 21: 48–54. Robeck, T.R., Schneyer, A.L., McBain, J.M., Dalton, L.M., Walsh, M.T., Czekala, N.M., and Kraemer, D.C., 1993, Analysis of urinary immunoreactive steroid metabolites and gonadotropins for characterization of the estrous cycle, breeding period, and seasonal estrus activity of captive killer whales (Orcinus orca), Zoo Biol., 12: 173–187. Robeck, T.R., Curry, B.E., McBain, J.F., and Kraemer, D.C., 1994a, Reproductive biology of the bottlenose dolphin (Tursiops truncatus) and the potential application of advanced reproductive technologies, J. Zoo Wildl. Med., 25: 321–336. Robeck, T.R., Gross, T., Walsh, M.T., Campbell, T., and McBain, J., 1994b, Preliminary results on radioimmunoassay determinations of post enzyme hydrolysis urinary progestin concentrations in the false killer whale (Pseudorca crassidens), in Proceedings of the 25th Annual International Association for Aquatic Animal Medicine, Vallejo, CA, 161.
0839_frame_C11 Page 234 Tuesday, May 22, 2001 11:09 AM
234
CRC Handbook of Marine Mammal Medicine
Robeck, T.R., Gross, T., and McBain, J., 1995, Preliminary results on radioimmunoassay determination of serum testosterone concentrations in the killer whale (Orcinus orca), in Proceedings 26th International Association for Aquatic Animal Medicine, Mystic Aquarium, 72–72. Robeck, T.R., McBain, J.F., Mathey, S., and Kraemer, D.C., 1998, Ultrasonographic evaluation of the effects of exogenous gonadotropins on follicular recruitment and ovulation induction in the Atlantic bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 6–13. Robeck, T.R., Jensen, E., Brook, F., Rouke, N., Rayner, C., and Kinoshita, R., 2000, Preliminary investigations into ovulation manipulation techniques in delphinids, in Proceedings of the American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, 222–225. Rogan, E., Baker, R., Jepson, P.D., Berrow, S., and Kiely, O., 1997, A mass stranding of white-sided dolphins (Lagenorhynchus acutus) in Ireland: Biological and pathological studies, J. Zool. Soc. London, 242: 217–227. Rommel, S.A., Early, G.A., Matassa, K.A., Pabst, D.A., and McLellan, W.A., 1995, Venous structures associated with thermoregulation of phocid seal reproductive organs, Anat. Rec., 243: 390–402. Ross, G., 1977, The taxonomy of bottlenosed dolphins, Tursiops species, in South African waters, Ann. Cape Prov. Mus. Nat. Hist., 11: 135–194. Samper, J.C., 1997, Techniques for artificial insemination, in Current Therapy in Large Animal Theriogenology, Youngquist, R.S. (Ed.), W.B. Saunders, Philadelphia, 36–42. Sawyer-Steffan, J.E., and Kirby, V.L., 1980, A study of serum steroid hormone levels in captive female bottlenose dolphins, their correlation with reproductive status, and their application to ovulation induction in captivity, National Technical Information Service, PB80-177199, Springfield, VA. Sawyer-Steffan, J.E., Kirby, V.L., and Gilmartin, W.C., 1983, Progesterone and estrogens in the pregnant and non-pregnant dolphin, Tursiops truncatus, and the effects of induced ovulation, Biol. Reprod., 28: 897–901. Schroeder, J.P., 1985, Artificial insemination of the bottlenose dolphin Tursiops truncatus, Proc. Am. Assoc. Zoo Vet., 122–124. Schroeder, J.P., 1990a, Breeding bottlenose dolphins in captivity, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 435–446. Schroeder, J.P., 1990b, Reproductive aspects of marine mammals, in CRC Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 353–369. Schroeder, J.P., and Keller, K.V., 1989, Seasonality of serum testosterone levels and sperm density in Tursiops truncatus, J. Exp. Zool., 249: 316–321. Schroeder, J.P., and Keller, K.V., 1990, Artificial insemination of bottlenose dolphins, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 447–460. Schurmayer, T.H., Knuth, U.A., Freischem, C.W., Sandow, J., Akhtar, F.B., and Nieschlag, E., 1984, Suppression of pituitary and testicular function in normal men by constant gonadotropin-releasing hormone agonist infusion, J. Clin. Endocrinol., 59: 19–24. Seager, S., Gilmartin, W., Moore, L., Platz, C., and Kirby, V., 1981, Semen collection (electroejaculation), evaluation and freezing in the Atlantic bottlenose dolphin (Tursiops truncatus), Proc. Am. Assoc. Zoo Vet., 136. Seargent, D.E., 1973, Biology of the white whales (Delphinapterus leucas) in western Hudson Bay, J. Fish. Res. Board Can., 30: 1065–1090. Seargent, D.E., Caldwell, D.K., and Caldwell, M.C., 1973, Age, growth, and maturity of bottlenose dolphin (Tursiops truncatus) from northeast Florida, J. Fish. Res. Board Can., 30: 1009–1011. Seargent, D.E., St. Aubin, D.J., and Geraci, J.R., 1980, Life history and northwest Atlantic status of the Atlantic white-sided dolphin, Lagenorhynchus acutus, Cetology, 37: 2–12. Seidel, G.E., Cran, D.G., Herickhoff, L.A., Schenk, J.L., Doyle, S.P., and Green, R.D., 1999, Inseminations of heifers with sexed frozen or sexed liquid semen, Theriogenology, 51: 400. Senger, P.L., 1986, Principles and procedures for storing and using frozen bovine semen, in Current Therapy in Theriogenology 2, Morrow, D.A. (Ed.), W.B. Saunders, Philadelphia, 162–174.
0839_frame_C11 Page 235 Tuesday, May 22, 2001 11:09 AM
Reproduction
235
Sharp, D., Robinson, G., Cleaver, B., and Porter, M., 1997, Role of photoperiod in regulating reproduction in mares: Basic and practical aspects, in Current Therapy in Large Animal Theriogenology, Youngquist, R.S. (Ed.), W.B. Saunders, Philadelphia, 71–78. Silvia, W.J., Lewis, G.S., McCracken, J.A., Thacher, W.W., and Wilson, L., Jr., 1991, Hormonal regulation of uterine secretion of prostaglandin F2α during luteolysis in ruminants, Biol. Reprod., 45: 655–663. Sisk, C.L., and Desjardins, C., 1986, Pulsatile release of luteinizing hormone and testosterone in male ferrets, Endocrinology, 119: 1195–1203. Squires, E.L., Stevens, W.B., McGlothlin, D.E., and Pickett, N.W., 1979, Effects of an oral progestin on the estrous cycle and fertility of mares, J. Anim. Sci., 49: 729–735. Squires, E.L., Heesemann, S.K., Webel, S.K., Shideler, K., and Voss, J.L., 1983, Relationship of altrenogist to ovarian activity, hormone concentrations and fertility of mares, J. Anim. Sci., 56: 901–909. Sreenan, J.M., 1988, Embryo transfer: Its uses and recent developments, Vet. Rec., 122: 624–629. Stevenson, J.S., and Davis, D.L., 1982, Estrus synchronization and fertility in gilts after 14- or 18-day feeding of altrenogest beginning at estrus or diestrus, J. Anim. Sci., 55: 119–123. Stone, L.R., Johnson, R.L., Sweeney, J.C., and Lewis, M.L., 1999, Fetal ultrasonography in dolphins with emphasis on gestational aging, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, E.R. (Eds.), W.B. Saunders, Philadelphia, 501–506. Sundaram, K., Connell, K.G., Bardin, C.W., Samojlik, E., and Schally, A.V., 1982, Inhibition of pituitarytesticular function with [D-Trp] luteinizing hormone-releasing hormone in rhesus monkeys, Endocrinology, 110: 1308–1314. Sweeney, J.C., Krames, B., Krames, J., and Stone, R., 2000, Stages of parturition, normal early calf development, and food energy requirements of the cow, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 289–296. Taverne, M.A.M., 1991, Applications of two-dimensional ultrasound in animal reproduction, Wien. Tierärztl. Monatsschr., 78: 341–345. Temte, J.L., 1985, Photoperiod and delayed implantation in the northern fur seal (Callorhinus ursinus), J. Reprod. Fertil., 73: 127–131. Temte, J.L., 1991, Precise birth timing in captive harbor seals (Phoca vitulina) and California sea lions (Zalophus californianus), Mar. Mammal Sci., 7: 145–156. Terasawa, F., Yokoyama, Y., and Kitamure, M., 1999, Rectal temperature before and after parturition in bottlenose dolphins, Zoo Biol., 18: 153–156. Theodorou, J., and Atkinson, S., 1998, Monitoring total androgen concentrations in saliva from captive Hawaiian monk seals (Monachus schauinslandi), Mar. Mammal Sci., 14: 304–310. Tom, L., Bhasin, S., Salameh, W., Peterson, M., Steiner, B., and Swerdloff, R.S., 1991, Male contraception: Combined GnRH antagonist and testosterone enanthate, Clin. Res., 39: 91A. Valle, E.R., Cruz, L.C., Cmarik, G.F., Ott, R.S., Peterson, L.A., and Kesler, D.J., 1986, The effect of GnRH and its method of administration on ovulation response, corpus luteum function and fertility of beef heifers synchronized with norgestomet and PGF 2α, J. Anim. Sci., 63: 132. Walker, L.A., Cornell, L., Dahl, K.D., Czekala, N.M., Dargen, C.M., Joseph, B.E., Hsueh, A.J.W., and Lasley, B.L., 1988, Urinary concentrations of ovarian steroid hormone metabolites and bioactive follicle-stimulating hormone in killer whales (Orcinus orca) during ovarian cycles and pregnancy, Biol. Reprod., 39: 1013–1020. Watson, P.F., 1979, The preservation of semen in mammals, in Oxford Reviews of Reproductive Biology, Vol. 1, Finn, C.A. (Ed.), Clarendon Press, Oxford, U.K., 283–350. Webel, S.K., and Squires, E.L., 1982, Control of the oestrous cycle in mares with altrenogest, J. Reprod. Fertil., Suppl. 32: 193–198. Wells, R., 2000, Reproduction in wild bottlenose dolphins: Overview of patterns observed during a longterm study, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 57–74.
0839_frame_C11 Page 236 Tuesday, May 22, 2001 11:09 AM
236
CRC Handbook of Marine Mammal Medicine
Wells, R.S., Boness, D.J., and Rathbun, G.B., 1999, Behavior, in Biology of Marine Mammals, Reynolds, J.E., III, and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 324–422. West, K.L., Atkinson, S., Carmichael, M.J., Sweeney, J.C., Kraemes, B., and Krames, J., 2000, Concentrations of progesterone in milk from bottlenose dolphins during different reproductive states, Gen. Comp. Endocrinol., 117: 218–224. Wildt, D.E., 1989, Reproductive research in conservation biology: Priorities and avenues for support, J. Zoo Wildl. Med., 20: 391–395. Wildt, D.E., 2000, Reproductive technology for conserving endangered species: Reality check, what is feasible and what is not, in Report from the Bottlenose Dolphin Breeding Workshop, Duffield, D.A., and Robeck, T.R. (Eds.), American Zoological Association Marine Mammal Taxon Advisory Group, Silver Spring, MD, 101–108. Wildt, D.E., Rall, W.F., Critser, J.K., Monfort, S.L., and Seal, U.S., 1997, Genome resource banks: “Living collections” for biodiversity conservation, Bioscience, 47: 698–704. Williamson, P., Gales, N.J., and Lister, S., 1990, Use of real-time B-mode ultrasound for pregnancy diagnosis and measurement of fetal growth in captive bottlenose dolphins (Tursiops truncatus), J. Reprod. Fertil., 88: 543–548. Wright, J.M., 1981, Non-surgical embryo transfer in cattle, embryo–recipient interaction, Theriogenology, 15: 433–56. Wright, P.J., and Malmo, J., 1992, Pharamacologic manipulation of fertility, Vet. Clin. North Am. Food Anim. Pract., 8: 57–89. Yoshioka, M., Tobayama, T., Inoue, S., and Katsumata, E., 2000, Semen collection, evaluation and cryopreservation of the bottlenose dolphin (Tursiops truncatus) and the Pacific white sided dolphin (Lagenorhychus obliquidens), unpublished manuscript. Young, S.J.F., and Huff, D.G., 1996, Fertility management in a female killer whale (Orcinus orca) with altrenogets (Regu-Mate), Proceedings International Association for Aquatic Animal Medicine, Baltimore, MD, 66–66. Youngquist, R.S., 1986, Cystic follicular degeneration in the cow, in Current Therapy in Theriogenology, Morrow, D.A. (Ed.), W.B. Saunders, Philadelphia, 243–246. Youngquist, R.S., 1997, Current Therapy in Large Animal Theriogenology, W.B. Saunders, Philadelphia.
0839_frame_C12.fm Page 237 Tuesday, April 9, 2002 1:36 PM
12
Immunology Donald P. King, Brian M. Aldridge, Suzanne Kennedy-Stoskopf, and Jeffrey L. Stott
Introduction The immune system is primarily a series of defense mechanisms that function to protect the body against the potential harmful effects of foreign microorganisms. In recent years, there have been rapid advances in the field of immunology. With these advances have come new methods for preventing and treating infectious disease. Although marine mammal immunology is a relatively recent field of scientific endeavor, it is already possible to perform reliable and pertinent studies to address specific aspects of health and disease in these species. Immune system monitoring and serological diagnostic assays have clear roles in the management of disease in individual marine mammals. In addition to clinical assessment, there are a number of other reasons to consider immunological parameters in marine mammals. The concept that the status and well-being of the aquatic environment are reflected in the immune systems of marine mammals has gained considerable acceptance within the last decade. Furthermore, there has also been a strong interest in genetic markers of immunological diversity, since many believe that the successful management of endangered populations may require assessment of genetic diversity. This chapter reviews the most recent advances in marine mammal immunology and immunodiagnostics. It concludes with what might be considered a typical approach to defining immunological dysfunction in a marine mammal. To date, clinical and experimental evidence support the notion that the immune systems of marine mammals share all the major identifiable components that have been described in detail for key terrestrial species, such as humans and rodents. However, it is likely that marine mammals possess some unique immunological features that reflect the adaptations required for survival and function in the aquatic environment. These adaptions may, in turn, reflect the spectrum of microbial pathogens that inhabit marine ecosystems, or may comprise homeostatic mechanisms that maintain immune function despite physiological extremes, such as hypoxia, hyperbaric pressures, or cold temperatures, that have been shown to be immunosuppressive in other species (Shinomiya, 1994; Knowles et al., 1996; Shepard and Shek, 1998; Brenner et al., 1999). Until more-detailed studies are performed, and immunological adaptations to the marine environment are documented, it is useful to refer to a generalized model of a mammalian immune system to understand how marine mammals mount a protective response to invading pathogens.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
237
0839_frame_C12.fm Page 238 Tuesday, May 22, 2001 11:09 AM
238
CRC Handbook of Marine Mammal Medicine
Overview of the Immune System The immune system has been classically divided into the innate and adaptive (or acquired) immune responses. Whereas the innate immune system is static relative to the quantity and quality of a response, the adaptive response gains quantity and quality (immunological memory) upon repeated exposure to the pathogen. Although these divisions are descriptively useful, it is important to realize that successful host defense responses rely on close orchestration between these two arms. To help the reader fully appreciate the progression of the immunological processes involved in pathogen clearance and host protection, a general inflammatory response will be described first, followed by the sequence of events that would occur following exposure to a pathogen.
Innate Immunity and the Inflammatory Response The normal, healthy, mammalian host is exposed to a vast number of potentially pathogenic microorganisms each day. Since clinical infectious disease is relatively uncommon in normal individuals, defense against these organisms must be a constant process. The majority of microorganisms are repelled by innate host defenses that include nonimmunological anatomical and physiological barriers (e.g., mucociliary blanket), antimicrobial factors (e.g., complement, lysozyme, lactoferrin, defensins, and reactive oxygen and nitrogen intermediates), and immunological effector cells (e.g., neutrophils, eosinophils, macrophages, and natural killer, or NK, cells). Many of these immune mechanisms act immediately following microbial invasion, particularly against those pathogens possessing identifiable structures such as lipopolysaccharide (LPS) present on Gram-negative bacteria, or double-stranded viral RNA. To be effective, the mammalian immune system possesses molecules capable of recognizing and neutralizing an enormous repertoire of infectious agents. Recognition is one of the key steps in the stimulation of early-induced immune responses that function to keep the infection under control, while the antigen-specific cells of the adaptive immune response are recruited and activated. At many portals for potential infection (e.g., mucosal surfaces), there are a number of locally produced antimicrobial peptides and cells that are sufficient to repel or eliminate a small pathogen load. However, in the event of an infection that can overwhelm these in situ defense mechanisms, an inflammatory process is initiated, aimed at destroying and eliminating the offending pathogen and at healing damaged tissues. Occasionally, the nature or extent of the localized inflammation may be severe enough to evoke a number of systemic inflammatory processes termed the acute-phase response, which serves to produce inflammatory mediators and recruit more inflammatory cells to the site of infection. The most important effector cells in these early phases of the immune response are phagocytes (tissue macrophages and migrating neutrophils). These not only trap, engulf, and destroy microbes, but also secrete cytokines that initiate the systemic acute-phase response and recruit additional leukocytes to magnify the local inflammatory response. The recruitment of cells involves chemotaxis and an increase in vascular endothelial cell and immune cell-adhesion molecule expression. These factors, in conjunction with an increased local blood flow and increased vascular permeability, lead to an accumulation of leukocytes, immunoglobulins, and other blood proteins in the infected tissue.
Adaptive Immune Response If a pathogen evades or overwhelms the innate defense mechanisms of the host, causing the foreign antigen to persist beyond the first several days of infection, an adaptive immune response is initiated. In contrast to the innate immune responses, the adaptive response produces effector cells (B- and T-lymphocytes) and molecules (immunoglobulins), which are highly specific to the
0839_frame_C12.fm Page 239 Tuesday, May 22, 2001 11:09 AM
Immunology
239
antigens of the invading microbe. In addition, the antigen-specific lymphocytes of the adaptive immune response are capable of swift clonal expansion and of a more rapid and effective immune response on subsequent exposures to the pathogen (immunological memory). The trigger for the adaptive immune response, the activation and proliferation of lymphocytes, takes place in organized lymphoid tissues. There are three major portals by which an invading pathogen can enter the host, namely, via a mucosal surface (respiratory tract, gastrointestinal tract), through the skin, or by direct inoculation into the bloodstream. At each of these portals are organized lymphoid tissues (mucosal-associated lymphoid tissue, regional lymph nodes, and spleen, respectively), which provide the organized microenvironment in which the intricate events of the adaptive immune response are closely coordinated. Microscopic investigations of the marine mammal immune system reveal that the morphology of the lymphoid organs is similar to terrestrial mammals, but with a few unique attributes (Romano et al., 1993; 1994; Cowan and Smith, 1995; 1999; Cowan, 1999; Smith et al., 1999). At these lymphoid sites, pathogens are trapped and engulfed by phagocytic cells. Some of the lymphoid cells are specialized for processing microbial antigens into small peptides, and presenting these peptides in association with highly polymorphic glycoproteins, called major histocompatibility (MHC) proteins, on their cell surface. The ability of the immune system to recognize and respond to such a vast array of foreign proteins is determined to a large degree by the number and structural diversity of the MHC molecules present in an individual. The polymorphic nature of these MHC proteins ensures maintenance of the host’s immunological vigor by minimizing the ability of a pathogen to avoid presentation by selective mutation. It is speculated that genetically restricted species, such as those that have been subjected to a “population bottleneck,” will lack MHC diversity. This is an area of increasing interest among marine mammal researchers (Gyllensten et al., 1990; Slade, 1992; Murray and White, 1998; Hoelzel et al., 1999; Zhong et al., 1999). The immunogenic peptides of the invading pathogens bound to the cell-surface MHC molecules are recognized by the highly specific receptors of T-helper lymphocytes, which by specific patterns of cytokine secretion stimulate either B lymphocyte expansion and antibody production (humoral immunity) or activation of macrophages (delayed-type hypersensitivity), and expansion and activation of cytotoxic T lymphocytes. The subsets of lymphocytes with these polarized patterns of cytokine production are T-helper1 and T-helper2 cells, respectively.
Cytokines The initiation, maintenance, and amplification of the immune response are regulated by soluble mediators called cytokines. Cytokines are the soluble messengers of the immune system and have the capacity to regulate many different cells in an autocrine, paracrine, and endocrine fashion. The predominant proinflammatory cytokines are interleukin-6 (IL-6), IL-1, and tumor necrosis factor alpha (TNF-α). These cytokines have a number of systemic effects, including body temperature elevation (fever), neutrophil mobilization, and stimulation of acute-phase protein production in the liver. Cytokines can also be immune effectors. Interferon-α (IFN-α) and INF-β are produced by a number of different cell-types following viral infection. They interfere with viral replication and can therefore limit the spread of viruses to uninfected cells. Additional cytokines such as IL-2, IL-4, IL-5, IL-10, IL-12, IL-15, and IFN-γ are pivotal in directing the development of both humoral and cellular immune responses. By using existing biological assays, it is possible to assay cytokine-like activity in mitogen-stimulated cultures (King et al., 1993b; 1995). Furthermore, cytokine transcripts from a number of marine mammals have been recently cloned and the DNA sequence determined (Table 1). The identification of these sequences will facilitate the development of molecular techniques for examining
0839_frame_C12.fm Page 240 Tuesday, May 22, 2001 11:09 AM
240
CRC Handbook of Marine Mammal Medicine
TABLE 1 Published Marine Mammal Cytokines Cytokine
Species
a
cDNA Clone (base pairs)
GSDB Accession Number
Reference
IL-1α
Bottlenose dolphin (Tursiops truncatus)
906
AB028215
Inoue et al., 1999c
IL-1β
Bottlenose dolphin (T. truncatus)
818
AB028216
Inoue et al., 1999c
IL-2
Killer whale (Orcinus orca) Beluga (Delphinapterus leucas) Northern elephant seal (Mirounga angustirostris) Gray seal (Halichoerus grypus) Manatee (Trichechus manatus latirostris)
455
AF009570
Ness et al., 1998
465
AF072870
658
U79187
St-Laurent et al., 1999 Shoda et al., 1998
468
AF072871
450
U09420
St-Laurent et al., 1999 Cashman et al., 1996
IL-4
Bottlenose dolphin (T. truncatus)
528
AB020732
IL-6
Killer whale (O. orca) Beluga (D. leucas) Harbor seal (Phoca vitulina) Southern sea otter (Enhydra lutris nereis)
670
L46803
King et al., 1996
627
AF076643
682
L46802
St-Laurent et al., 1999 King et al., 1996
676
L46804
King et al., 1996 Herman, unpubl. data
IL-10
Killer whale (O. orca)
548
U93260
IFN-γ
Killer whale (O. orca) Bottlenose dolphin (T. truncatus)
144
—
548
AB022044
a
Inoue et al., 1999b
King, unpubl. data Inoue et al., 1999a
Genome Sequence Database.
cytokine gene expression during infectious disease. These techniques have great potential for improving the ability to measure immune cell activity in marine mammals.
Immunodiagnostics Inflammation Monitoring the changes associated with inflammation is a key component of diagnostic tests that establish the overall health of an animal. Unfortunately, in part because of the presence of blubber, the cardinal signs classically used to define inflammation in humans and domestic species can be difficult to recognize in some marine mammals. However, experimental and clinical data from human and veterinary medicine demonstrate that changes in the concentrations of specific proteins (collectively referred to as acute-phase proteins) can aid the detection and quantification of inflammation. In human medicine, the acute-phase response can be assessed by measuring the erythrocyte sedimentation rate (ESR) of blood collected into anticoagulant. This method is, in part, an
0839_frame_C12.fm Page 241 Tuesday, May 22, 2001 11:09 AM
Immunology
241
indirect measurement of fibrinogen and has been successfully adapted for use in a variety of cetacean species. Unfortunately, as a result of variable serum lipid content, this method is unreliable for detecting inflammation in pinnipeds. Determination of serum iron concentration can also be used as an indirect measure of inflammation in cetaceans, but has not been evaluated in pinnipeds (see Chapter 19, Clinical Pathology). Although these methods are widely used in marine mammal medicine, current efforts are directed at identifying inflammation at earlier stages. The characteristics of ideal markers are that they exhibit dramatic changes in serum concentrations early in a systemic inflammatory response and that they are not influenced by other physiological changes, such as malnutrition or handling stresses. Recent approaches in this field employ the measurement of the specific protein mediators of the acute-phase response such as IL-6. This cytokine is produced by macrophages at the site of tissue damage and injury. Unlike most other cytokines that possess only local activity, IL-6 enters the systemic circulation and is a key player in the induction of acute-phase protein synthesis in the liver. Recent studies using a murine bioassay system have suggested that IL-6 may prove to be a valuable indicator of inflammation in a number of marine mammal species (King et al., 1993b). Of the many acute-phase proteins, C-reactive protein (CRP) and serum amyloid A have been targeted for use in clinical medicine. The potential utility of CRP has been highlighted in a recent study in harbor seals (Phoca vitulina) that measured increases of CRP in excess of 100-fold associated with clinical signs of inflammatory disease, compared with apparently healthy animals (Funke et al., 1997).
Cellular Immunity Classical differential white blood cell counts can morphologically distinguish and enumerate major leukocyte subpopulations into lymphocytes, monocytes, eosinophils, and neutrophils (see Chapter 19, Clinical Pathology). These cells, although ultimately derived from the same progenitor bone marrow stem cell population, make different functional contributions to the immune system. There is a wide range of immunological techniques that can be used to evaluate the cellular immune system. Broadly speaking, these assays can be divided into those that measure the phenotypic qualities of leukocytes (lymphocyte subpopulations and the cellsurface density of adhesion proteins) and those that assess functional aspects of the cells. Recently, a major use of these assays in marine mammals has been to examine immunological dysfunction arising from the presence of environmental pollutants (de Swart et al., 1995; 1996; Lahvis et al., 1995; Ross et al., 1995a,b). Furthermore, since the immune system is acutely sensitive, these methods have the potential to measure the influence of many internal and external stresses that affect marine mammals (see Chapter 13, Stress). The peripheral blood represents the most convenient sampling window for the assessment of the cellular immune system. In many circumstances, cells can also be isolated from tissues, such as spleen and lymph nodes collected during post-mortem examination, and can be subsequently used in phenotypic assays and/or in vitro functional testing. Tissues and blood collected into anticoagulant should be transported to the laboratory and used as soon as possible after collection. A major requirement of such assays is the availability of purified and viable mononuclear leukocytes. Classically, cells should be purified and either cryopreserved and stored in liquid nitrogen or placed in culture within 24 hours of sample acquisition. The recent introduction of CPT tubes (Becton Dickinson, Franklin Lakes, NJ) has provided researchers with a novel method of obtaining mononuclear cells from peripheral blood without the need to use density-gradient techniques. Centrifugation of the CPT vacutainer tubes at 1800 g results in the mononuclear cells being permanently separated from granulocytes and red blood cells. The length of centrifugation must be determined for each species to optimize
0839_frame_C12.fm Page 242 Tuesday, May 22, 2001 11:09 AM
242
CRC Handbook of Marine Mammal Medicine
cell yield and purity (i.e., horse blood is typically centrifuged for 7 min, cow blood for 30 min, whereas most cetacean and pinniped blood is centrifuged for 18 to 20 min). Immunophenotyping refers to methods that delineate multiple leukocyte subpopulations in the blood. Analysis of the density of cell-surface adhesion and activation antigens on these leukocyte populations has also become possible. Alone or in combination, these techniques are finding increasing application in identifying subtle immunological perturbations caused by infectious agents and other tissue insults. This analysis is performed using flow cytometry and requires characterized markers, usually monoclonal antibodies that are specific for unique determinants expressed on the various leukocyte populations (also commonly referred to as CD, cluster of differentiation or leukocyte differentiation antigens). Unfortunately, only a limited number of the anti-CD markers available from academic and commercial sources crossreact with marine mammal white blood cells (Romano et al., 1992; De Guise et al., 1997b). As might be expected, the ability of these reagents to cross-react usually parallels phylogenetic relationships. Consequently, it is more likely that antibovine CD reagents will work for cetacean blood and that anticanine/feline reagents will work for pinniped samples. To perform a full complement of analyses, a number of species-specific monoclonal antibodies have been developed and are currently being characterized for pinnipeds and cetaceans (De Guise et al., 1998). Future development in this area will likely see the extension of this panel of reagents. Since these reagents are usually produced in a serendipitous manner using immunizations with mixed cell populations, cloning and expression studies such as those performed for beluga (Delphinapterus leucas) CD4 (Romano et al., 1999) may be required to allow the development of some antibodies against individual cell determinants.
Functional Immune Testing In Vitro
The capacity of lymphocytes to proliferate in response to antigen is central to the success of the adaptive immune system. This mechanism allows small numbers of antigen-specific lymphocytes to be rapidly increased to counteract an invading pathogen. In vitro blastogenesis assays mimic this response and measure the ability of isolated blood cells to proliferate in response to broadspectrum mitogenic stimulation. These assays have been successfully adapted and optimized for use with marine mammal samples (Mumford et al., 1975; Colgrove, 1978; de Swart et al., 1993; Lahvis et al., 1993; Ross et al., 1993; De Guise et al., 1996). Differential use of mitogens such as the plant lectins (concanavalin A, phytohemagglutinin, pokeweed mitogen) and bacterial lipopolysaccharide can serve as a relative measure of T- and B-lymphocyte responsiveness. A useful alternative to these traditional blastogenesis assays is to measure the expression of the IL2 receptor on lymphocytes by flow cytometry. This technique, adapted for harbor seals and bottlenose dolphins (Tursiops truncatus), uses labeled recombinant human IL-2, which binds to upregulated IL-2 receptors expressed on activated lymphocytes (DiMolfetto-Landon et al., 1995; Erickson et al., 1995). These proliferation assays have the capacity to be modified for measuring pathogen-specific T-cell responses. Assays to assess function of additional leukocytes such as phagocytosis (De Guise et al., 1995) and NK cell function (Ross et al., 1995b; De Guise et al., 1997a) have also been described for harbor seals and belugas. In Vivo
In marine mammals, challenge experiments with pathogens are rarely feasible or ethical as a means of studying immune function. The delayed-type hypersensitivity (DTH) skin test represents the only practical in vivo method of assessing cellular immune function. This procedure
0839_frame_C12.fm Page 243 Tuesday, May 22, 2001 11:09 AM
Immunology
243
involves inoculating an antigen intradermally into the individual under investigation and monitoring the local immune response over the following 48 to 72 hours. The monitoring procedure can be as simple as measuring changes in skin thickness at the site of antigen inoculation. More complete information can be obtained by histopathological examination of a skin biopsy taken from the same region. A DTH response is characterized by a γ-INFassociated influx of macrophages. The γ-INF is secreted by TDTH helper cells. For this reason, this assay can be used for measuring antigen-specific immune cell responsiveness. Since animals must be housed for up to several weeks, this approach is not practicable for many field situations involving marine mammals. Decreased DTH responses to ovalbumin were measured in a Dutch study examining the effects of environmental contaminants on harbor seal immune responsiveness (Ross et al., 1995a). The results implicated an immunosuppressive effect of pollutants upon cellular components of the immune system.
Humoral Immunity Immunoglobulins (antibodies) are soluble, antigen-specific effector molecules of the adaptive immune response. These proteins are produced by B-lymphocytes of the humoral immune system. Distinct classes of immunoglobulin molecules (IgG, IgM, IgA, and, of lesser importance in the peripheral circulation, IgE and IgD) have been identified in most mammalian species studied. Because of their relatively high concentration in serum, purification and characterization of these proteins are often the first tasks undertaken by comparative immunologists. To date, several studies have characterized immunoglobulin molecules with characteristic component heavy and light chains, using sera collected from a selected number of cetacean and pinniped species (Nash and Mach, 1971; Cavagnolo and Vedros, 1978; Carter et al., 1990). Binding of immunoglobulin proteins to unique determinants (epitopes) on foreign proteins is an important mechanism by which pathogens are targeted for subsequent elimination from the body. By measuring changes in the circulating levels of antigen-specific immunoglobulin, exposure to infectious agents can be documented. This can be used in epidemiological studies of infectious disease and to enhance the management and prevention of disease outbreaks by identifying naive unexposed animals. It must be emphasized, however, that pathogen-specific antibody levels do not necessarily confirm the presence of an active pathogen. Serum is the most readily obtainable and conveniently sampled source for measuring systemic humoral immune responses. Carefully collected sera can often be stored over prolonged periods at −70°C without seriously compromising their performance in diagnostic assays. Sera can also be stored for long periods at −20°C, providing the freezer is not frost-free. Serum stored in frostfree freezers will become desiccated with time and should no longer be used for the detection and measurement of antibodies. As a general rule, it is good practice to dispense sera into multiple aliquots (of appropriate volume), because this will minimize the freezing/ thawing of samples. For larger organizations, a dedicated serum bank will prove valuable for long-term monitoring of individuals and for epidemiological studies of disease in populations. Access to good quality sera is particularly important when performing retrospective serological studies.
Measurement of Pathogen-Specific Antibodies (Serodiagnostics) Vast arrays of laboratory-based assays have been developed to measure pathogen-specific antibodies. Although they may vary in specificity and sensitivity, most of the approaches described below provide useful serological information when performed with appropriate controls. These controls increase the confidence of the assay data and assist with the interpretation of the results.
0839_frame_C12.fm Page 244 Tuesday, May 22, 2001 11:09 AM
244
CRC Handbook of Marine Mammal Medicine
When possible, assays should be performed with established positive and negative reference sera. Unfortunately, prior exposure to pathogens, particularly for free-ranging marine mammals, is rarely documented. In these cases, designated hyperimmune sera from a closely related species can be substituted. For example, commercially available canine distemper virus (CDV) immune sera have been successfully used to validate a morbillivirus seroassay for use in harbor seals (Ham Lammé et al., 1999). To discriminate actively infected animals from those with prior exposure to the pathogen in question, it is important to use paired sera that have been collected at least 14 to 21 days apart. For many assays, a fourfold increase in antibody titer between these time points is indicative of active infection. In the absence of defined clinical signs of disease in a population, care and consideration must be taken before serological evidence alone can confirm the presence or absence of a pathogen. Since microbiology of marine mammal diseases is in its infancy, there are probably many microorganisms yet to be discovered (see Chapter 15, Viral Diseases; Chapter 16, Bacterial Diseases). Therefore, the possibility that the test used is detecting a similar agent that shares common structural domains with the agent for which it was designed should not be excluded. When possible, serological studies should be performed in concert with other independent methods such as viral/bacterial isolation or molecular identification of genomic sequences.
Serum/Virus Neutralization Test The serum/virus neutralization test (SNT/VNT) is an in vitro assay that estimates the amount of pathogen-specific antibody that neutralizes the replication and subsequent cytopathic effect of a defined dose of virus. In recent years, SNTs have been successfully developed and used to monitor exposure to a number of marine mammal-specific viruses including morbilliviruses (Visser et al., 1990; Van Bressem et al., 1993), herpesviruses (Borst et al., 1986), and caliciviruses (Smith, 1987). An advantage of this test is that it is sensitive and highly specific (e.g., defining viral serotypes) for the viral pathogen being employed in the assay. However, false positives can arise as a result of the presence of serum constituents that are somewhat toxic to cells and directly inhibit virus replication. These substances are common in samples collected postmortem. Further limitations of SNTs are that they can be lengthy assays to perform, requiring up to 7 to 14 days before they can be evaluated, and require cell culture expertise and equipment that is not easily adapted for field situations. SNTs require that the laboratory must have access to appropriate isolates of the virus in question and cells in which this virus can replicate, limiting its use to a small number of specialized laboratories. Although not necessarily a negative attribute, the neutralization assay is serotype-specific and will not necessarily detect antibody to closely related viruses.
Precipitation/Agglutination Techniques These are traditional serodiagnostic techniques that exploit the ability of antibodies to form visible aggregates with antigen. The precipitation reaction employs soluble antigen (e.g., agar gel immunodiffusion, AGID), whereas the agglutination reaction utilizes particulate antigen (e.g., bacterial agglutination reaction) or soluble antigen bound to inert particles (e.g., latex beads). The advantage of these tests is that they are cheaper than SNT/VNT, since specialized, and therefore often expensive, equipment is not required. Furthermore, since specific reagents are not necessary, existing assays for human and domestic species are usually easily adapted for use with marine mammal sera. Examples of the use of these methods with marine mammal sera include recent studies to determine the prevalence of antibodies to Brucella (Tryland et al., 1999), as well other studies investigating the serology of Leptospira (Vedros et al., 1971). A disadvantage of these tests is that, since the formation of large immune complexes is inhibited
0839_frame_C12.fm Page 245 Tuesday, May 22, 2001 11:09 AM
Immunology
245
by excess amounts of antibody or antigen (prozone effect), careful titrations must be performed to optimize the assays.
Enzyme-Linked Immunosorbent Assay In recent years, enzyme-linked immunosorbent assays (ELISA) have been increasingly used in serological diagnostics. The basis for these assays is that the antigen in question is immobilized onto a solid phase, usually to a specially treated 96-well plastic plate. Antigen-specific immunoglobulin is detected by stepwise incubations with the test sera followed by an antispecies secondary reagent covalently linked to an enzyme reporter such as horseradish peroxidase or alkaline phosphatase. Although the approach described above (indirect ELISA) normally requires purified antigen, ELISA methods can be modified by the use of pathogenspecific reagents, so that antigen is captured from solution (trapping ELISA) prior to the addition of the test sera. A limited number of monoclonal and polyclonal species-specific secondary antibodies for pinniped immunoglobulin are available (Carter et al., 1990; King et al., 1993a). In the absence of species-specific reagents, staphylococcal protein A (SPA) and/or streptococcal protein G (SPG) (Ross et al., 1994; Reidarson et al., 1998) can be used. These are commercially available bacterial cell wall components that have been shown to bind the Fc portion of most mammalian immunoglobulin molecules. For most marine mammal species tested to date, SPA appears to be the preferred reagent for ELISA. In addition to these valuable reagents, development of further monoclonal antibody markers is anticipated in the near future. This should allow the subsequent establishment of new and more sensitive serological tests for these species. These exquisitely sensitive techniques are rapid to perform, adaptable to field situations, inexpensive, and can be easily applied to a large number of samples. However, assay specificity is dependent on the degree of antigen purity and is therefore easily compromised.
Total Immunoglobulin Even in a hyperimmunized individual, the component of immunoglobulin that is specific for one particular antigen is usually less than 5%. Therefore, changes in the concentration of total immunoglobulin (classes and subclasses) are not usually indicative of the progression of an immune response. However, total immunoglobulin concentrations do have a diagnostic utility. Measurement of IgG concentrations in serum is performed in clinical situations to determine if passive transfer of immunoglobulin has occurred in neonates of species that are transiently hypogammaglobulinemic at birth. The clinical utility of total immunoglobulin concentrations has been demonstrated in animals with recurrent bacterial infections, suspected autoimmune disease, and lymphoproliferative disorders. In these instances, quantifying IgG is used to support a specific diagnosis, and is not used as an evaluation of specific immune function. Increases and decreases in serum levels of IgG are the consequences rather than the cause of complex events. A decrease in IgG does not necessarily mean an animal is functionally immunocompromised. A number of investigators have used a variety of techniques to quantify serum immunoglobulin concentrations of pinnipeds (Calvagnolo and Vedros, 1979; Carter et al., 1990; King et al., 1994; 1998; Marquez et al., 1998). Interestingly, serum IgG concentrations in pinnipeds appear to be significantly elevated compared with those of terrestrial carnivores. These studies provide the basis for the future establishment of clinically useful baseline values for these species. Further work in this area is still required to determine if low or abnormally elevated immunoglobulin levels are consistent with any recognized disease entities in marine mammals.
0839_frame_C12.fm Page 246 Tuesday, May 22, 2001 11:09 AM
246
CRC Handbook of Marine Mammal Medicine
Clinical Approach to Suspected Marine Mammal Immunological Disorders Immunological disorders can be broadly divided into those with immunological overactivity (autoimmunity, allergic, hypersensitivity) and those with immune system deficiency. Anecdotal reports on marine mammals suggest that immunological insufficiency is a more common concern for clinicians, so this subsection will focus on a clinical approach to cases in which a functional impairment of the immune system is suspected. A number of immunodeficiency classification schemes have been developed in other species. These may be categorized by etiology (primary or secondary) or by the predominant compartment of the immune system affected (humoral, cellular, combined). Since there is little published information regarding clinical immunology in marine mammals, it is useful to borrow these classification criteria, at least until a better understanding of factors and conditions that impair immune function in marine mammals is gained. The classification scheme proposed by the World Health Organization (WHO) is broadly based on which compartment of the immune system is involved in the deficiency. Primary immunodeficiencies are those caused by intrinsic defects (congenital or acquired). This category consists of a large number of inherited defects, but also includes intrinsic defects induced by environmental insults. In secondary immunodeficiencies (Table 2), there are no intrinsic abnormalities in the development or function of B or T cells, but, instead, an external factor or condition interferes with immune function. These include viral-induced immunodeficiencies and those arising from stress, malnutrition, neoplasia, parasitic infections, or iatrogenic factors. Clinical investigations of suspected immunodeficiency should be directed at identifying which compartments of the immune system are affected. This is the first step in determining an underlying cause for the abnormality. Immunodeficiency syndromes, by definition, are characterized by an unusual susceptibility to infection. This susceptibility may include frequent infections with common or opportunistic microbes, unusually severe infections, or the failure of an infection to respond to antibiotics to which the suspect organism is susceptible. The type and extent of the infection provides the first clue to the nature of the immune dysfunction. For example, recurrent infections with pyogenic bacteria are likely caused by defects in B-lymphocytes or humoral (antibody-mediated) immunity. Severe fungal infections are more compatible with T-lymphocyte deficiencies. The repeated formation of abscesses with low-grade pathogens may suggest a neutrophil deficiency. If one or more of these scenarios is present, then it is reasonable to suspect a compromised immune system. The next tasks are to confirm this diagnosis using a stepwise clinical approach (Figure 1), to examine the possible cause of the immune compromise. In many marine mammal species, specific information regarding the immune system is difficult to obtain by physical examination, because of the difficulty in palpating external lymph nodes. However, an assessment of the size and state of the lymphoid organs (thymus, spleen, lymph nodes) can be useful, particularly in detecting primary immunodeficiency states, and may circumvent the need for extensive laboratory evaluations. If possible, this information should be acquired using radiographic or ultrasonographic imaging of these organs (see Chapters 24 through 28, Diagnostic Imaging). Information on the immune system can be obtained from routine hematological examinations (white blood cell counts with leukocyte differentials) and clinical serum chemistry analyses (see Chapter 19, Clinical Pathology). An immunological abnormality must be suspected when a persistent lymphopenia or neutropenia is observed. A marked or progressive hypo- or hyperglobulinemia, in the absence of serum albumin changes, can also indicate an immune dysfunction. It is important to emphasize that the presence of any abnormality should be confirmed by repeated sampling and comparison with healthy, age-matched individuals.
0839_frame_C12.fm Page 247 Tuesday, May 22, 2001 11:09 AM
247
Immunology
TABLE 2 Possible Causes of Secondary Marine Mammal Immunodeficiencies Inciting Cause
Possible Mechanism
a
Follow-Up Tests
Failure of passive transfer
Immunoglobulin deficiency
Serum immunoglobulin levels
Malnutrition (protein, caloric, and/or micronutrient) (Chandra, 1997)
Multifactorial and complex: impaired antibody production, cell-mediated immunity, phagocyte function, and complement activity
Response to nutritional supplementation Lymphocyte proliferation
Trauma/surgery (Page and Ben-Elihau, 2000)
Acute-phase response Cytokine imbalance Pain (neuroendocrine)
Response to analgesics Acute-phase proteins
Viral infection (see Chapter 15)
Varies with etiological agent (e.g. lymphoid depletion, suppression of lymphocyte proliferation, downregulation of MHC expression)
Identification of viral agent Lymph node biopsy Lymphocyte proliferation Flow cytometry
Hormonal (e.g., endocrine imbalance, pregnancy) (Mellor and Munn, 2000) (see Chapter 10)
Varies with hormone (e.g., pregnancy can invoke a cytokine imbalance)
Detection of pregnancy Flow cytometry Lymphocyte proliferation
Bacterial infection (Song et al., 2000) (see Chapter 16)
Cytokine imbalance
Identification of pathogen Acute-phase proteins Lymphocyte imbalance
Stress (Elenkov et al., 1999) (see Chapter 13)
Hormonally induced changes in cytokines, lymphocyte function, and expression of cell-surface proteins
Flow cytometry Lymphocyte proliferation
Neoplasia/malignancy (see Chapter 23)
Quantitative and qualitative alterations in humoral and cell-mediated immunity
Flow cytometry Serum immunoglobulin levels Lymph node/bone marrow biopsy
Drug-induced (e.g., corticosteroids)
Varies with drug; corticosteroids affect cytokine production
Flow cytometry Lymphocyte proliferation
a
Tests with abnormal results should be repeated and results compared with those of age-matched control animals.
Although these simple hematological and serum protein changes are clearly not pathognomonic for immune deficient states, they do provide strong justification for pursuing more specific and reliable immunological testing. Furthermore, there are a large number of immunological disorders that will not be detected by changes in absolute leukocyte numbers or serum globulin levels. Many of the species-specific assays for reliably examining the immune systems of marine mammals are not available through routine diagnostic laboratories. However, there are a number of research laboratories that are able to provide advice and to perform these services. A broad evaluation of the cellular immune system can be obtained by immunophenotypic analysis and lymphocyte function tests. A detailed serum immunoglobulin profile will provide useful information concerning the antibody-producing capabilities of the immune system. In most cases, these tests will identify the nature and extent of an immune dysfunction. On occasion, the abnormality is more subtle, and its identification and characterization may require
0839_frame_C12.fm Page 248 Tuesday, May 22, 2001 11:09 AM
248
CRC Handbook of Marine Mammal Medicine
FIGURE 1 Clinical evaluation of the immune system.
the inoculation of an exogenous antigen and the measurement of the responses induced by the inoculation.
Conclusion In summary, there is a need to expand the knowledge and understanding of immunological disorders in marine mammals. By adopting a systematic approach to examining the immune system, it is possible to determine the nature and extent and, possibly, the etiology of immune dysfunction in an individual. This information will be vital in designing management and preventative strategies in susceptible populations.
Acknowledgments The authors thank Tracy Romano for her peer-review of this chapter.
References Borst, G.H., Walvoort, H.C., Reijnders, P.J., van der Kamp, J.S., and Osterhaus, A.D., 1986, An outbreak of a herpes virus infection in harbor seals (Phoca vitulina), J. Wildl. Dis., 22: 1–6. Brenner, I., Shepard, R.J., and Shek, P.N., 1999, Immune function in hyperbaric environments, diving and decompression, Undersea Hyperbaric Med., 26: 27–39. Carter, S.D., Hughes, D.E., and Baker, J.R., 1990, Characterization and measurement of immunoglobulins in the grey seal (Halichoerus grypus), J. Comp. Pathol., 102: 13–23.
0839_frame_C12.fm Page 249 Tuesday, May 22, 2001 11:09 AM
Immunology
249
Cashman, M.E., Ness, T.L., Roess, W.B., Bradley, W.G., and Reynolds, J.E., 1996, Isolation and characterization of a cDNA encoding interleukin 2 from the Florida manatee, Tricbechus manatus latirostris, Mar. Mammal Sci., 12: 89–98. Cavagnolo, R.Z., and Vedros, N.A., 1978, Identification and characterization of three immunoglobulin classes in the northern fur seal Callorhinus ursinus, Dev. Comp. Immunol., 2: 689–697. Cavagnolo, R.Z., and Vedros, N.A., 1979, Serum and colostrum immunoglobulin levels in the northern fur seal Callorhinus ursinus, Dev. Comp. Immunol., 3: 139–146. Chandra, R.K., 1997, The immune suppressive nature of pain, Semin. Oncol. Nurs., 1: 10–15. Colgrove, G.S., 1978, Stimulation of lymphocytes from a dolphin (Tursiops truncatus) by phytomitogens, Am. J. Vet. Res., 39: 141–144. Cowan, D.F., 1999, Involution and cystic transformation of the thymus in the bottlenose dolphin, Tursiops truncatus, Vet. Pathol., 31: 648–653. Cowan, D.F., and Smith, T., 1995, Morphology of the complex lymphoepithelial organs of the anal canal (“anal tonsil”) in the bottlenose dolphin, Tursiops truncatus, J. Morphol., 223: 263–268. Cowan, D.F., and Smith, T., 1999, Morphology of the lymphoid organs of the bottlenose dolphin, Tursiops truncatus, J. Anat., 194: 505–517. De Guise, S., Flipo, D., Boehm, J.R., Martineau, D., Beland, P., and Fournier, M., 1995, Immune functions in beluga whales (Delphinapterus leucas): Evaluation of phagocytosis and respiratory burst with peripheral blood leukocytes using flow cytometry, Vet. Immunol. Immunopathol., 47: 351–362. De Guise, S., Bernier, J., Dufresne, M.M., Martineau, D., Beland, P., and Fournier, M., 1996, Immune functions in beluga whales (Delphinapterus leucas): Evaluation of mitogen-induced blastic formation of lymphocytes from peripheral blood, spleen and thymus, Vet. Immunol. Immunopathol., 50: 117–126. De Guise, S.D., Ross, P.S., Osterhaus, A.D., Martineau, D., Beland, P., and Fournier, M., 1997a, Immune functions in beluga whales (Delphinapterus leucas): Evaluation of natural killer cell activity, Vet. Immunol. Immunopathol., 58: 345–354. De Guise, S., Bernier, J., Martineau, D., Beland, P., and Fournier, M., 1997b, Phenotyping of beluga whale blood lymphocytes using monoclonal antibodies, Dev. Comp. Immunol., 21: 425–433. De Guise, S., Erickson, K., Blanchard, M., Dimolfetto, L., Lepper, H., Wang, J., Stott, J.L., and Ferrick, D.A., 1998, Characterization of a monoclonal antibody that recognizes a lymphocyte surface antigen for the cetacean homologue to CD45R, Immunology, 94: 207–212. de Swart, R.L., Kluten, R.M., Huizing, C.J., Vedder, L.J., Reijnders, P.J., Visser, I.K., UytdeHaag, F.G., and Osterhaus, A.D., 1993, Mitogen and antigen induced B and T cell responses of peripheral blood mononuclear cells from the harbour seal (Phoca vitulina), Vet. Immunol. Immunopathol., 37: 217–230. de Swart, R.L., Ross, P.S., Timmerman, H.H., Vos, H.W., Reijnders, P.J., Vos, J.G., and Osterhaus, A.D., 1995, Impaired cellular immune response in harbour seals (Phoca vitulina) feeding on environmentally contaminated herring, Clin. Exp. Immunol., 101: 480–486. de Swart, R.L., Ross, P.S., Vos, J.G., and Osterhaus, A.D., 1996, Impaired immunity in harbour seals (Phoca vitulina) exposed to bioaccumulated environmental contaminants: Review of a long-term feeding study, Environ. Health Perspect., 104 Suppl. 4: 823–828. DiMolfetto-Landon, L., Erickson, K.L., Blanchard-Channell, M., Jeffries, S.J., Harvey, J.T., Jessup, D.A., Ferrick, D.A., and Stott, J.L., 1995, Blastogenesis and interleukin-2 receptor expression assays in the harbor seal (Phoca vitulina), J. Wildl. Dis., 31: 150–158. Elenkov, I.J., and Chrousos, G.P., 1999, Stress, cytokine patterns and susceptibility to disease, Baillieres Best Pract. Res. Clin. Endocrinol. Metab., 13: 583–595. Erickson, K.L., DiMolfetto-Landon, L., Wells, R.S., Reidarson, T., Stott, J.L., and Ferrick, D.A., 1995, Development of an interleukin-2 receptor expression assay and its use in evaluation of cellular immune responses in bottlenose dolphin (Tursiops truncatus), J. Wildl. Dis., 31: 142–149.
0839_frame_C12.fm Page 250 Tuesday, May 22, 2001 11:09 AM
250
CRC Handbook of Marine Mammal Medicine
Funke, C., King, D.P., Brotheridge, R.M., Adelung, D., and Stott, J.L., 1997, Harbor seal (Phoca vitulina) C-reactive protein (C-RP): Purification, characterization of specific monoclonal antibodies and development of an immuno-assay to measure serum C-RP concentrations, Vet. Immunol. Immunopathol., 59: 151–162. Gyllensten, U.B., Lashkari, D., and Erlich, H., 1990, Allelic diversification at the class II DQB locus of the mammalian major histocompatibility complex, Proc. Natl. Acad. Sci. U.S.A., 87: 1835–1839. Ham Lammé, K.D., King, D.P., Taylor, B., House, C., Jessup, D.A., Jeffries, S., Yochem, P., Gulland, F., Ferrick, D.A., and Stott, J.L., 1999, The application of immuno-assays for serological detection of morbillivirus exposure in free-ranging harbor seals (Phoca vitulina) and sea otters (Enhydra lutris) from the Western coast of the United States, Mar. Mammal Sci., 15: 601–608. Hoelzel, A.R., Stephens, J.C., and O’Brien, S.J., 1999, Molecular genetic diversity and evolution at the MHC DQB locus in four species of pinnipeds, Mol. Biol. Evol., 16: 611–618. Inoue, Y., Itou, T., Oike, T., and Sakai, T., 1999a, Cloning and sequencing of the bottlenose dolphin (Tursiops truncatus) interferon-gamma gene, J. Vet. Med. Sci., 61: 939–942. Inoue, Y., Itou, T., Sakai, T., and Oike, T., 1999b, Cloning and sequencing of a bottlenose dolphin (Tursiops truncatus) interleukin-4-encoding cDNA, J. Vet. Med. Sci., 61: 693–696. Inoue, Y., Itou, T., Ueda, K., Oike, T., and Sakai, T., 1999c, Cloning and sequencing of a bottlenose dolphin (Tursiops truncatus) interleukin-1alpha and -1beta complementary DNAs, J. Vet. Med. Sci., 61: 1317–1321. King, D.P., Hay, A.W., Robinson, I., and Evans, S.W., 1993a, The use of monoclonal antibodies specific for seal immunoglobulins in an enzyme-linked immunosorbent assay to detect canine distemper virus-specific immunoglobulin in seal plasma samples, J. Immunol. Methods, 160: 163–171. King, D.P., Robinson, I., Hay, A.W., and Evans, S.W., 1993b, Identification and partial characterization of common seal (Phoca vitulina) and grey seal (Haliochoerus grypus) interleukin-6-like activities, Dev. Comp. Immunol., 17: 449–458. King, D.P., Lowe, K.A., Hay, A.W., and Evans, S.W., 1994, Identification, characterization, and measurement of immunoglobulin concentrations in grey (Haliocherus grypus) and common (Phoca vitulina) seals, Dev. Comp. Immunol., 18: 433–442. King, D.P., Hay, A.W., Robinson, I., and Evans, S.W., 1995, Leucocyte interleukin-1-like activity in the common seal (Phoca vitulina) and grey seal (Halichoerus grypus), J. Comp. Pathol., 113: 253–261. King, D.P., Schrenzel, M.D., McKnight, M.L., Reidarson, T.H., Hanni, K.D., Stott, J.L., and Ferrick, D.A., 1996, Molecular cloning and sequencing of interleukin 6 cDNA fragments from the harbor seal (Phoca vitulina), killer whale (Orcinus orca), and southern sea otter (Enhydra lutris nereis), Immunogenetics, 43: 190–195. King, D.P., Sanders, J.L., Nomura, C.T., Stoddard, R.A., Ortiz, C.L., and Evans, S.W., 1998, Ontogeny of humoral immunity in northern elephant seal (Mirounga angustirostris) neonates, Comp. Biochem. Physiol. B, 121: 363–368. Knowles, R., Keeping, H., Nguyen, K., Graeber, T., D’Amico, R., and Simms, H., 1996, Hypoxemia/ reoxygenation down-regulates interleukin-8-stimulated regulation of CD16 and CD35 mRNA expression, Surgery, 120: 382–387. Lahvis, G.P., Wells, R.S., Casper, D., and Via, C.S., 1993, In vitro lymphocyte response of bottlenose dolphins (Tursiops truncatus): Mitogen-induced proliferation, Mar. Environ. Res., 35: 115–119. Lahvis, G.P., Wells, R.S., Kuehl, D.W., Stewart, J.L., Rhinehart, H.L., and Via, C.S., 1995, Decreased lymphocyte responses in free-ranging bottlenose dolphins (Tursiops truncatus) are associated with increased concentrations of PCBs and DDT in peripheral blood, Environ. Health Perspect., 103: 67–72. Marquez, M.E., Carlini, A.R., Slobodianik, N.H., Ronayne de Ferrer, P.A., and Godoy, M.F., 1998, Immunoglobulin M serum levels in females and pups of southern elephant seal (Mirounga leonina) during the suckling period, Comp. Biochem. Physiol. A, 119: 795–799. Mellor, A.L., and Munn, D.H., 2000, Immunology at the maternal–fetal interface: Lessons for T cell tolerance and suppression, Annu. Rev. Immunol., 18: 367–391.
0839_frame_C12.fm Page 251 Tuesday, May 22, 2001 11:09 AM
Immunology
251
Mumford, D.M., Stockman, G.D., Barsales, P.B., Whitman, T., and Wilbur, J.R., 1975, Lymphocyte transformation studies of sea mammal blood, Experientia, 31: 498–500. Murray, B.W., and White, B.N., 1998, Sequence variation at the major histocompatibility complex DRB loci in beluga (Delphinpterus leucas) and narwhal (Monodon monoceros), Immunogenetics, 48: 242–252. Murray, B.W., Malik, S., and White, B.N., 1995, Sequence variation at the major histocompatibility complex locus DQB in beluga whales (Delphinapterus leucas), Mol. Biol. Evol., 12: 582–593. Nash, D.R., and Mach, J.P., 1971, Immunoglobulin classes in aquatic mammals: Characterization by serologic cross-reactivity, molecular size and binding of human free secretory component, J. Immunol., 107: 1424–1430. Ness, T.L., Bradley, W.G., Reynolds, J.E., and Roess, W.B., 1998, Isolation and expression of the interleukin-2 gene from the killer whale, Orcinus orca, Mar. Mammal Sci., 14: 531–543. Page, G.G., and Ben-Elihau, S., 2000, Immune suppression in polymicrobial sepsis: Differential regulation of Th1 and Th2 responses by p38 MAPK, J. Surg. Res., 91:141–146. Reidarson, T.H., McBain, J., House, C., King, D.P., Stott, J.L., Krafft, A., Taubenberger, J.K., Heyning, J., and Lipscomb, T.P., 1998, Morbillivirus infection in stranded common dolphins from the Pacific Ocean, J. Wildl. Dis., 34: 771–776. Romano, T.A., Ridgway, S.H., and Quaranta, V., 1992, MHC class II molecules and immunoglobulins on peripheral blood lymphocytes of the bottlenose dolphin, Tursiops truncatus, J. Exp. Zool., 263: 96–104. Romano, T.A., Felten, S.Y., Olschowka, J.A., and Felten, D.L., 1993, A microscopic investigation of the lymphoid organs of the beluga, Delphinapterus leucas, J. Morphol., 215: 261–287. Romano, T.A., Felten, S.Y., Olschowka, J.A., and Felten, D.L., 1994, Noradrenergic and peptidergic innervation of lymphoid organs in the beluga, Delphinapterus leucas: An anatomical link between the nervous and immune systems, J. Morphol., 221: 243–259. Romano, T.A., Ridgway, S.H., Felten, D.L., and Quaranta, V., 1999, Molecular cloning and characterization of CD4 in an aquatic mammal, the white whale Delphinapterus leucas, Immunogenetics, 49: 376–383. Ross, P.S., Pohajdak, B., Bowen, W.D., and Addison, R.F., 1993, Immune function in free-ranging harbor seal (Phoca vitulina) mothers and their pups during lactation, J. Wildl. Dis., 29: 21–29. Ross, P.S., de Swart, R.L., Visser, I.K., Vedder, L.J., Murk, W., Bowen, W.D., and Osterhaus, A.D., 1994, Relative immunocompetence of the newborn harbour seal, Phoca vitulina, Vet. Immunol. Immunopathol., 42: 331–348. Ross, P.S., de Swart, R.L., Reijnders, P.J., Van Loveren, H., Vos, J.G., and Osterhaus, A.D., 1995a, Contaminant-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., Vedder, L.J., Van Loveren, H., Vos, J.G., Reijnders, P.J.H., and Osterhaus, A.D.M.E., 1995b, Suppression of natural killer activity in harbor seals (Phoca vitulina) fed Baltic Sea herring, Aquat. Toxicol., 34: 71–84. Shepard, R.J., and Shek, P.N., 1998, Cold exposure and immune function, Can. J. Physiol. Pharmacol., 76: 828–836. Shinomiya, N., Suzuki, S., Hashimoto, A., and Oiwa, H., 1994, Effects of deep saturation diving on the lymphocyte subsets of healthy divers, Undersea Hyperbaric Med., 21: 277–286. Shoda, L.K., Brown, W.C., and Rice-Ficht, A.C., 1998, Sequence and characterization of phocine interleukin 2, J. Wildl. Dis., 34: 81–90. Slade, R.W., 1992, Limited MHC polymorphism in the southern elephant seal: Implications for MHC evolution and marine mammal population biology, Proc. R. Soc. London, 249: 163–171. Smith, A.W., 1987, San Miguel Sealion virus, in Virus Infections of Carnivores, Appel, M., Ed., Elsevier Science Publication, Amsterdam, 481–489. Smith, A.W., Skilling, D.E., Benirschke, K., Albert, T.F., and Barlough, J.E., 1987, Serology and virology of the bowhead whale (Balaena mysticetus L.), J. Wildl. Dis., 23: 92–98.
0839_frame_C12.fm Page 252 Tuesday, May 22, 2001 11:09 AM
252
CRC Handbook of Marine Mammal Medicine
Smith, T.L., Turnball, B.S., and Cowan, D.F., 1999, Morphology of the complex laryngeal gland in the Atlantic bottlenose dolphin, Tursiops truncatus, Anat. Rec., 254: 98–106. Song, G.Y., Chung, C.S., Chaudry, I.H., and Ayala, A., 2000, IL-4-induced activation of the stat6 pathway contributes to the suppression of cell-mediated immunity and death in sepsis, Surgery, 128: 133–138. St-Laurent, G., Beliveau, C., and Archambault, D., 1999, Molecular cloning and phylogenetic analysis of beluga whale (Delphinapterus leucas) and grey seal (Halichoerus grypus) interleukin 2, Vet. Immunol. Immunopathol., 67: 385–394. Tryland, M., Kleivane, L., Alfredsson, A., Kjeld, M., Arnason, A., Stuen, S., and Godfroid, J., 1999, Evidence of Brucella infection in marine mammals in the North Atlantic Ocean, Vet. Rec., 144: 588–592. Van Bressem, M.F., Visser, I.K., de Swart, R.L., Orvell, C., Stanzani, L., Androukaki, E., Siakavara, K., and Osterhaus, A.D., 1993, Dolphin morbillivirus infection in different parts of the Mediterranean, Arch. Virol., 129: 235–242. Vedros, N.A., Smith, A.W., Schonewald, J., Migaki, G., and Hubbard, R.C., 1971, Leptospirosis epizootic among California sea lions, Science, 172: 1250–1251. Visser, I.K.G., Grachev, M.A., Orvell, C., DeVries, P., Broeders, J., van de Bildt, M.W.G., Groen, J., Teppema, J.S., Burger, M.C., Uyt de Haag, F.G.C.M., and Osterhaus, A.D.M.E., 1990, Comparison of two morbilliviruses from seals during outbreaks of distemper in northwest Europe and Siberia, Arch. Virol., 111: 149–164. Zhong, J.F., Harvey, J.T., and Boothby, J.T., 1999, Characterization of a harbor seal class I major histocompatability complex cDNA clone, Immunogenetics, 48: 422–424.
0839_frame_13fm Page 253 Tuesday, May 22, 2001 11:10 AM
13 Stress and Marine Mammals David J. St. Aubin and Leslie A. Dierauf
Introduction As early as 450 B.C., Hippocrates considered health to be a state of harmonious balance and disease a state of disharmony (Chrousos, 1988). The Oxford English Dictionary notes that the word stress first appeared in the literature in 1303, but did not occur in the context of biological science until 1936 (OED, 1999). In that year, the journal Nature published a short article entitled “A Syndrome Produced by Diverse Nocuous Agents” by Hans Selye (Selye, 1936). This article laid the groundwork for current stress research by describing a three-stage syndrome of (1) alarm and adaptation, (2) hormonal events, and (3) resistance, exhaustion, and death, where “the symptoms ... are independent of the nature of the damaging agent or the pharmacological type of drug employed” (Neylan, 1998). Moberg (1985; 1987a) further defined Selye’s three stages of stress as (1) recognition of the stressful stimulus, (2) the body’s actual response to the stimulus, and (3) the resulting consequences to the body. Is stress harmful? The answer is no and yes. When an individual can predict and control the threatening stressor, a coping mechanism can be established. It might even be argued that periodic activation of the stress response is beneficial to maintaining health in the same way that physically demanding exercise promotes fitness. However, when the responses to stress are uncontrolled, excessive, and prolonged, a state of distress results. Distress is not always deleterious, although it is unpleasant and uncomfortable (Goldstein, 1995). This chapter considers the diverse mammalian responses to stressors and examines manifestations of the stress response in marine mammals. The chapter addresses clinical approaches and indicators for assessing stress in these species and concludes by identifying needs for future research to sharpen diagnostic abilities and to allow better prediction of the long-term consequences of stress.
Stressors Stressors are not equally stressful to all individuals or species. The response to a given stressor depends on how an animal’s sensory systems receive and interpret information about the surrounding environment, the reaction to this information, and the degree of positive and negative feedback that occurs during the response (Lovallo, 1997a). Experience and acclimation will blunt the response to potentially stressful procedures; for example, bottlenose dolphins
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
253
0839_frame_13fm Page 254 Tuesday, May 22, 2001 11:10 AM
254
CRC Handbook of Marine Mammal Medicine
(Tursiops truncatus) can become quite tolerant to transportation in a stretcher. The introduction of novel stimuli into an animal’s environment constitutes a stress for some, but necessary enrichment for others. A new arrival in a pinniped colony can either enhance the social framework or precipitate stressful aggression. In a captive setting, where it is desirable to eliminate, or at least manage, potential stressors in an animal’s environment to optimize health, it is important to evaluate each case in the context of the species and individuals involved. In the wild, marine mammals encounter natural stressors daily. Predators, demanding meteorological and oceanographic conditions, intraspecific aggression, and even aspects of their normal activities, such as prolonged fasts or extended dives, are significant challenges to homeostasis and may elicit stress responses. Of greater concern is the impact of unnatural or anthropogenic stressors on the health of marine mammals, particularly species that are threatened or endangered. Increasingly, biologists and medical professionals are called upon to evaluate and provide opinions that might lead to important management decisions. For example, in 1997, an amendment to the U.S. Marine Mammal Protection Act, directed the National Marine Fisheries Service to conduct a review of the scientific literature on stress to provide a context for future research concerning the effects of stress on dolphins (Curry, 1999). Human activities such as vessel traffic, fishing, petroleum and mineral exploration and development, low-frequency sounds for ocean thermometry, and sonar systems are highly controversial, in terms of the degree to which they elicit damaging stress responses in marine mammals. Oil spills (Geraci and St. Aubin, 1990) and other environmental contaminants can be directly harmful, but more often the impact must be measured through subtle physiological changes considered indicators of stress. There are few experimental data to address these points, largely because it is difficult to identify “control” populations in the wild or to isolate the effects of one particular stressor in the midst of a substantially degraded habitat.
Stress Response and Regulation The literature on the stress response in mammals identifies four broad categories of interest: physiology, endocrinology, immunology, and neurology. There is considerable overlap among these, particularly since hormones alter physiological processes and immune responses, neurological stimulation elicits certain endocrine and physiological responses and is also linked to the immune system, and mediators of inflammation activate some endocrine pathways (Figure 1). Within each category, there is the important consideration of whether the response is acute or chronic, and whether the associated perturbations are beneficial or more damaging than the original stressor. Survival for the organism depends on feedback regulation of many of these systems, and when unchecked or stimulated to exhaustion, the result is distress and possibly death (Breazile, 1987). A significant challenge to studying stress in marine mammals, or any wild species, is to obtain baseline data representing an unstressed state. Chase, capture, restraint, and sampling procedures are recognized stressors that can influence analytes, sometimes within minutes. In captivity, cetaceans and pinnipeds can be trained to allow specimen collection with minimal disturbance, yielding data that are as close to resting as can be expected. At the very least, the slight deviations that might be encountered under such circumstances serve as controls for the same procedures that must be employed to assess stress in free-ranging individuals or in captive animals suspected of stress-related abnormalities. For those working in the field, rapid and efficient capture strategies can sometimes be designed to allow specimen collection of baseline quality.
0839_frame_13fm Page 255 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
255
FIGURE 1 Major changes to body systems under stress.
Neurological Factors The acute stress response begins with recognition of a stressor, and is initially orchestrated by the limbic and hypothalamic centers of the brain. Perception of a stressful stimulus produces fear and anxiety, which feed back to the limbic system of the brain. Corticotropinreleasing factor (CRF) is secreted from the hypothalamus (paraventricular nucleus) and the limbic system, and is the main neuropeptide regulator activating the hypothalamic–pituitary–adrenal (HPA) axis (Rivier, 1991). It also acts as a neurotransmitter, helping integrate the animal’s sensory, behavioral, and endocrinological responses to stimuli (Lovallo, 1997a). Direct innervation of the adrenal medulla results in the release of catecholamines to adjust physiological processes; neurological connections to lymph nodes serve to link the central nervous and immune systems. These elements of the stress response in marine mammals are examined below.
0839_frame_13fm Page 256 Tuesday, May 22, 2001 11:10 AM
256
CRC Handbook of Marine Mammal Medicine
Endocrine Factors The primary endocrine components of the stress response are derived from the autonomic nervous system (norepinephrine, or NEpi), the adrenal medulla (epinephrine, or Epi, and NEpi), the hypothalamus (CRF), the pituitary (adrenocorticotropic hormone, or ACTH), the adrenal gland (cortisol, corticosterone, and aldosterone), and the brain (NEpi and β-endorphins) (Dunn, 1995; 1996; Lovallo, 1997b). Secondarily, enkephalins, substance P, neuropeptide Y, prolactin, growth hormone, thyroid hormones, vasopressin, angiotensin II, vasoactive intestinal peptides, and other pituitary hormones become involved in a cascading fashion (Dunn, 1995; 1996; Breazile, 1988). For many of these, there are no specific data from marine mammals. Nevertheless, there has been significant progress in the last three decades in the understanding of how some of these hormones participate in the stress response in these animals (Table 1). Information on the normal function of marine mammal endocrine systems is presented in Chapter 10, Endocrinology. Catecholamines
Catecholamines (Epi and NEpi) are the first line of defense in an animal’s response to stress—the “fight or flight” reaction. Their effects are induced rapidly, and circulating levels can be altered by the mere anticipation of a stressful event. Unlike many other hormones, the changes that they elicit subside quickly. The physiological systems affected by catecholamines are many, but principally involve the cardiovascular system and energy metabolism, to prepare the organism for immediate action. Thomas et al. (1990) examined changes in catecholamine levels in captive belugas (Delphinapterus leucas) exposed to playbacks of high-amplitude noise from oil-drilling rigs. Although the animals’ initial response was to flee, there was little or no consistent effect on circulating levels of catecholamines (Epi: 0 to 101 pg/ml; NEpi: 160 to 604 pg/ml). St. Aubin and Geraci (unpubl. data) compared Epi and NEpi concentrations in 29 belugas sampled immediately after a 5- to 15-min pursuit, with those in 10 whales captured and held for repeated sampling over 5 days. Epi levels averaged 634 pg/ml at the time of capture, but only 76 pg/ml in 95 samples collected during the 5-day holding period. Average NEpi concentrations of 1423 pg/ml after capture declined only slightly to a mean of 1042 pg/ml. The latter hormone is generally more reflective of muscular activity and discharge from the sympathetic nervous system than anxiety or alarm. Recent investigations on stranded cetaceans have revealed a pattern of lesions suggestive of massive release of endogenous catecholamines (Turnbull and Cowan, 1998; Cowan, 2000). Contraction band necrosis in cardiac and skeletal muscle, along with injuries of ischemia and reperfusion in gut and kidney, are manifestations of an excessive and prolonged alarm response, with fatal consequences. These observations were thought to account for the abrupt deaths during handling of highly stressed, stranded marine mammals. The acute deaths of three ringed seals (Phoca hispida) exposed experimentally to an oil spill (Geraci and Smith, 1976) were less a function of the toxicity of the petroleum than of cardiac tissue hypersensitized to certain volatile hydrocarbons by the stress of the situation (St. Aubin, 1990). Glucocorticoids
Glucocorticoids (cortisol, corticosterone) have three functions in stress. They (1) alter carbohydrate metabolism to increase circulating substrates for energy; (2) permit catecholamines to act on metabolic pathways and blood vasculature; and (3) provide protective adaptations to distress by limiting immunological reactions, including inflammation, thus minimizing cell and tissue damage (Munck et al., 1984; Breazile, 1988). Cortisol is the dominant circulating
Increase
Neutrophils
Capture and handling
Capture and handling
Immune
Capture and handling Capture and handling Capture and handling Noise playback Capture and handling Noise playback Capture and handling Glucocorticoid administration
No change Increase No change No change Increase No change Increase Increase
Increase
Capture and handling Capture and handling Capture and handling
Decrease No change Decrease
Leukocytes
Insulin
Norepinephrine
Epinephrine
Reverse triidothyronine
Triiodothyronine
Capture and handling Capture and handling
No change Increase
Aldosterone
No change No change
Herpesvirus infection Stranding Capture and handling Capture and handling
Corticosterone Arginine vasopressin Thyroxine
Capture and handling
Endocrine
Stressor
Increase
Effect
Cortisol
Factor
Beluga Ringed seal (P. hispida) Beluga Bottlenose dolphin
Beluga Bottlenose dolphin Beluga Bottlenose dolphin Bottlenose dolphin Beluga Bottlenose dolphin Beluga Beluga Beluga Beluga Bottlenose dolphin
Beluga Bottlenose dolphin Bottlenose dolphin
Beluga (Delphinapterus leucas) Bottlenose dolphin (Tursiops truncatus) Harbor seal (Phoca vitulina) Pilot whale (Globicephala melas) Bottlenose dolphin Bottlenose dolphin
Species
TABLE 1 Stress Indicators in Marine Mammals (effects noted are in blood, unless otherwise indicated)
Stress and Marine Mammals
continued
St. Aubin and Geraci, 1989 Geraci and Smith, 1975 St. Aubin and Geraci, 1989 Medway and Geraci, 1964
St. Aubin and Geraci, 1988; 1992 St. Aubin et al., 1996 St. Aubin and Geraci, 1988; 1992 Orlov et al., 1988 St. Aubin et al., 1996 St. Aubin and Geraci, 1988; 1992 St. Aubin et al., 1996 Thomas et al., 1990 St. Aubin and Geraci, unpubl. Thomas et al., 1990 St. Aubin and Geraci, unpubl. Reiderson and McBain, 1999
St. Aubin and Geraci, 1989; 1992 Thomson and Geraci, 1986; St. Aubin et al., 1996 Gulland et al., 1999 Geraci and St. Aubin, 1987 Ortiz and Worthy, 2000 Thomson and Geraci, 1986; St. Aubin et al., 1996 St. Aubin and Geraci, 1989 Ortiz and Worthy, 2000 Ortiz and Worthy, 2000
Reference
0839_frame_13fm Page 257 Tuesday, May 22, 2001 11:10 AM
257
Bottlenose dolphin
Handling
Seven species of cetaceans
Disease, entrapment, habitat degradation
Increase Decrease Present (in tissue) Increase (in tissue)
Creatine kinase
Haptoglobins
Alkaline phosphatase Contraction band necrosis Stress responsive proteins
Disease Stranding
Disease
Disease Capture and handling Handling
No change Increase
Sodium
Dugong (Dugong dugon) Bottlenose dolphin Harp seal (Pagophilus groenlandicus) Ringed seal Bottlenose dolphin Harp seal Beluga Steller sea lion (Eumetopias jubatus) and harbor seal Bottlenose dolphin Various cetacean species
Capture Capture and handling Nutritional stress
Increase No change Decrease
Miscellaneous Diagnostics
Gray seal (Halichoerus grypus)
Ringed seal Bottlenose dolphin Beluga Bottlenose dolphin Ringed seal Bottlenose dolphin
Ringed seal Bottlenose dolphin Beluga Bottlenose dolphin
Species
Intradermal PHA injection
Glucocorticoid administration
Glucocorticoid administration Capture and handling
Glucocorticoid administration Capture and handling
Stressor
Fothergill et al., 1991 Turnbull and Cowan, 1998; Cowan, 2000 Southern, 2000
Marsh and Anderson, 1983 Ortiz and Worthy, 2000 Geraci 1972, Engelhardt and Geraci, 1978 Geraci et al., 1979 Ortiz and Worthy, 2000 St. Aubin et al., 1979 St. Aubin and Geraci, 1989 Zenteno-Savin et al., 1997
Medway and Geraci, 1964
Geraci and Smith, 1975 Medway et al., 1970 St. Aubin and Geraci, 1989 Medway and Geraci, 1964; Thomson and Geraci, 1986 Geraci and Smith, 1975 Medway et al., 1970 St. Aubin and Geraci, 1989 Thomson and Geraci, 1986 Geraci and Smith, 1975 Medway et al., 1970; Reidarson and McBain, 1999 Hall et al., 1999
Reference
258
Potassium
Decrease
Lymphocytes
Effect
Decreased proliferation (in tissue) No change
Decrease
Eosinophils
Factor
TABLE 1 (CONTINUED) Stress Indicators in Marine Mammals (effects noted are in blood, unless otherwise indicated)
0839_frame_13fm Page 258 Tuesday, May 22, 2001 11:10 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_13fm Page 259 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
259
glucocorticoid in all marine mammals studied to date, although corticosterone levels vary in parallel with those of cortisol in bottlenose dolphins (Ortiz and Worthy, 2000). Exogenous ACTH has been used to alter circulating levels of cortisol in some odontocetes and pinnipeds, providing a standard against which stress-induced changes can be measured (see Chapter 10, Endocrinology). No dose–response studies have been attempted, reflecting the cautious experimental approach that must often be used with marine mammals, especially cetaceans. Despite this caution, two bottlenose dolphins tested with ACTH died 2 and 5 days later, possibly as a cumulative effect of preexisting stress (Thomson and Geraci, 1986). Thus, it is difficult to define in absolute terms what the maximum potential is for glucocorticoid secretion in marine mammals. Even with such information, it is misleading to use the degree of corticosteroid elevation as a direct measure of the intensity of the stressor (Rushen, 1986). Capture and handling is a stressor that is of particular interest to those who must manipulate animals in captivity or in the wild. Thomson and Geraci (1986) compared cortisol levels in bottlenose dolphins calmly captured and sampled within 10 min, with those in dolphins subjected to 3 hours of pursuit prior to sampling. The cortisol levels of the former group averaged approximately 1.25 µ g/dl, whereas the latter showed concentrations of 2.5 µ g/dl. During the next 7 hours, when the animals were held in stretchers to simulate transport and allow the collection of serial samples, cortisol levels for the most part did not rise above 4.7 µ g/dl, with no clear differences seen based on the earlier treatment of the dolphins. In the calmly captured animals, cortisol levels rose steadily during the first 90 min to reach concentrations similar to those in the first samples obtained from the chased dolphins. The changes as a result of handling and restraint were comparable to those following ACTH administration. Overall, the elevations in cortisol were modest, compared with those in other species. Wild bottlenose dolphins unaccustomed to capture or, in the case of the population in Sarasota Bay, Florida, infrequently captured might be expected to exhibit a stronger glucocorticoid response to this stress, but no such difference was noted (St. Aubin et al., 1996; Ortiz and Worthy, 2000). The narrow range of cortisol concentrations in most cetaceans limits its utility as a stress indicator. Although Thomson and Geraci (1986) concluded that it was a good measure of adrenal activity in bottlenose dolphins, Ortiz and Worthy (2000) found that cortisol levels were no higher in free-ranging dolphins sampled more than 41 min after capture than in those sampled within 27 min. Either the animals were undisturbed by the procedure in the latter study or the specimens were drawn before changes in cortisol occurred. In ACTH stimulation studies in this species, cortisol levels rose only slightly during the first hour postinjection (Thomson and Geraci, 1986). Belugas sampled at capture and after a 3- to 5-hour transport to field holding facilities showed rising levels of cortisol, from 3.2 to 5.8 µ g/dl (St. Aubin and Geraci, 1989). When they were next sampled, 2 to 4 days later, concentrations were comparable to the lower values found immediately following capture. The dynamics of the cortisol response to handling stress were examined in more detail in belugas serially sampled every 6.5 hours over a 5-day period after capture (St. Aubin and Geraci, 1992). Following the hour-long process of lowering the water twice daily to access the whales, blood cortisol levels averaged 3.9 µ g/dl, whereas samples collected 6 hours after acclimation to shallow water showed a mean cortisol level of 2.7 µ g/dl. As noted following ACTH administration, a cortisol response to stress is expected after 1 to 2 hours, with a return to baseline levels 4 to 5 hours later, in the absence of continued stimulation (St. Aubin and Geraci, 1990). Extreme elevations in cortisol have been noted in marine mammals in distress. Stranded pilot whales (Globicephala melas) on the shore for more than 6 hours showed levels up to 16 µ g/dl,
0839_frame_13fm Page 260 Tuesday, May 22, 2001 11:10 AM
260
CRC Handbook of Marine Mammal Medicine
far in excess of any values recorded after ACTH stimulation or other handling (Geraci and St. Aubin, 1987). It is likely that these supraphysiological concentrations were the result of reduced hepatic clearance in animals in shock. Gulland et al. (1999) found that harbor seals (P. vitulina) infected with an adrenotropic herpesvirus showed elevated baseline cortisol levels that peaked at an average of 38.7 ± 16 µ g/dl within 2 hours of death. Mineralocorticoids
The mineralocorticoid aldosterone is not customarily considered as part of the stress response in most mammals. However, a series of studies and other fortuitous observations have revealed its particular role in stress in marine mammals. It has been postulated that the role of aldosterone in water conservation is beneficial to stressed marine mammals, especially those that may not soon have an opportunity to acquire water through feeding (see Chapter 10, Endocrinology). Stimulation by ACTH elicits a proportionally larger elevation in aldosterone in bottlenose dolphins (Thomson and Geraci, 1986), belugas (St. Aubin and Geraci, 1990), ringed and harp seals (Pagophilus groenlandicus) (St. Aubin and Geraci, 1986), and northern fur seals (Callorhinus ursinus) (St. Aubin et al., unpubl. data) than it does in terrestrial mammals. Consistent with these findings, capture and handling stress produce the same changes in belugas (St. Aubin and Geraci, 1989) and bottlenose dolphins (Thomson and Geraci, 1986; St. Aubin et al., 1996), although Ortiz and Worthy (2000) found no aldosterone release in the latter species during the time frame of their sampling. Aldosterone elevations, when they do occur, are highly variable, peaking in less than 1 hour in some cases and at 3 hours in others; still other animals show no residual elevation after 3 hours of continuous handling (Thomson and Geraci, 1986). The sensitivity of aldosterone to central stimulation from the pituitary and higher neurological centers in phocid seals provides a mechanism that is subject to exhaustion and failure during chronic stress. The result is hyponatremia (Geraci, 1972a), which can occur not only in salt-restricted environments, as might be expected, but also as a consequence of a variety of nonspecific stresses such as vitamin deficiency (Geraci, 1972b; Engelhardt and Geraci, 1978). In the wild, ringed seals in poor body condition from undetermined causes also exhibit hyponatremia, suggesting that they were chronically stressed (Geraci et al., 1979). Thyroid Hormones
The activity of the thyroid gland is modulated during stress to conserve resources for more urgent survival needs. Thyroid hormone (TH)-mediated mobilization of energy stores could be adaptive at such times, but not the thermogenic catabolism that accompanies this process. Ridgway and Patton (1971) recognized that capture stress profoundly affects TH balance in some cetaceans. In belugas, St. Aubin and Geraci (1988; 1992) noted decreased levels of triiodothyronine (T3) approximately 6 to 8 hours after capture, whereas changes in thyroxine (T4) did not occur until more than 20 hours later. There was no recovery in whales monitored for as long as 10 weeks. During the acute phase of the response, levels of reverse T3 (rT3) rose, suggesting a diversion of the metabolism of T4 to the inactive rT3 rather than to the physiologically potent T3. Administration of ACTH depressed T3 levels even farther. These changes are consistent with glucocorticoid-mediated effects in other mammals (Larsen et al., 1998). Cortisol is capable of inhibiting thyrotropin secretion from the anterior pituitary, and also the tissue monodeiodinating enzyme that is responsible for converting much of the T4 in the circulation to T3. Because of its short half-life in circulation, T3 declines relatively rapidly, whereas T4 shows a more gradual decrease from a larger pool of circulating hormone that is not being replenished from the thyroid gland. A similar pattern of change in T3, but not T4,
0839_frame_13fm Page 261 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
261
was observed in an abbreviated study on bottlenose dolphins (Orlov et al., 1988). The dramatic changes in TH in belugas may have been exaggerated by the coincident annual stimulation of thyroid activity at the time when the studies were performed (see Chapter 10, Endocrinology). Other Hormones
There is little information on the role of other hormones in the stress response of marine mammals. The dynamics of growth hormone, prolactin, insulin, and glucagon, among others, bear investigation, considering their importance in producing metabolic adjustments that are advantageous during stress. Reidarson and McBain (1999) noted an increase in insulin levels in two dolphins given glucocorticoids to stimulate appetite. Arginine vasopressin (AVP) was examined for its possible influence on ACTH, and concomitantly adrenocortical hormones, in captured bottlenose dolphins, but no relationship was found (Ortiz and Worthy, 2000).
Immunological Factors For many years, the potent anti-inflammatory and immunosuppressive properties of glucocorticoids were not readily reconciled with a general impression that the stress response better equips the organism to meet potentially threatening conditions (Munck et al., 1984). A fully charged immune system would seem to be the best defense against opportunistic pathogens. Yet, it is widely recognized that stress can render individuals more, rather than less, susceptible to disease (Levine, 1993; Leonard and Miller, 1995). The suppressive action of glucocorticoids on the immune system is necessary to keep in check a powerful complement of cells and cell mediators that eventually would be detrimental (Keller et al., 1991; McEwen et al., 1997). Some of the mediators released during inflammation stimulate CRF secretion from the hypothalamus and, consequently, increase ACTH and cortisol levels to abate the immune response (Lovallo, 1997a). The general organization of the immune system in marine mammals is considered in Chapter 12, Immunology. Assessment of the various cellular and biochemical components of this system is a rapidly expanding discipline, and has grown quickly from the long-standing reliance on leukocyte differential counts to lymphocyte phenotyping, cytokine analysis, and blastogenesis studies (see Chapter 12, Immunology) (DiMolfetto-Landon et al., 1995; Erickson et al., 1995; Nielsen, 1995; Blanchard et al., 1999). Leukocyte counts are a convenient, albeit “low-tech,” approach to recognizing stress in these animals. The classic stress leukogram (leukocytosis, neutrophilia, eosinopenia, lymphopenia) attributable to the action of glucocorticoids on the various cell lines was described to varying degrees in bottlenose dolphins subjected to transportation stress (Medway and Geraci, 1964), treated with glucocorticoids (Medway et al., 1970; Reidarson and McBain, 1999) or following ACTH administration (Thomson and Geraci, 1986), and in belugas after capture (St. Aubin and Geraci, 1989) or ACTH (St. Aubin and Geraci, 1990). Ringed seals stressed by capture in nets showed similar changes (Geraci and Smith, 1975). Dexamethasone suppressed lymphocyte proliferation in gray seals (Halichoerus grypus) injected intradermally with the mitogen phytohemagluttinin (PHA) (Hall et al., 1999). Taken together, these observations demonstrate that the immune systems of marine mammals display the same sensitivity as other species to stress-related hormonal changes, and that stress may compromise their ability to resist infection. The immune system is also subject to direct regulation by the central nervous system. In belugas, as in other mammals, lymphoid organs are innervated by noradrenergic and peptidergic fibers (Romano et al., 1994). Activation of central structures during the stress response therefore has the potential to affect immunological activity.
0839_frame_13fm Page 262 Tuesday, May 22, 2001 11:10 AM
262
CRC Handbook of Marine Mammal Medicine
Indicators of Acute and Chronic Stress To help diagnose and treat stress in marine mammals, interdisciplinary teams are working to develop clinically useful laboratory tests to quantify better acute, prepathological, and chronic stress reactions (Figure 2). Because the stress response is a series of complex interrelated events, differing from species to species and from individual to individual within each species, this is a daunting task. Where in the stress response should one look for valid indicators—at the start (i.e., early warning systems), midway, or at the end (end-point measurements)? Are negative results as valuable as positive results in testing for stress indicators? Should one be looking for direct or indirect measurements of stressful events? What is the best way to induce stress to study it? These are questions that must be answered by those engaged in stress research, prior to designing any study.
Acute Response Behavioral assessments are commonly used to recognize acute stress. Anxiety is often the first outward sign of an animal under stress. Chrousos and Gold (1992) and Dunn (1995) suggest this anxiety results from the release of norepinephrine from the noradrenergic neurons in the brain stem locus ceruleus. Dolphins disturbed by the presence of, and noise from, a large ship, positioned themselves as far away from it as possible, and showed “agitation, stress and fear” by tail-slapping, head-slapping, hyperactive swimming, bunching about, and thrashing (Norris et al., 1978; Norris and Dohl, 1980). In some situations, passivity rather than hyperactivity might signal stress, as noted in spinner (Stenella longirostris) and spotted dolphins (S. attenuata) that were encircled as part of the tuna fishery (Norris et al., 1978). The acute stages of the stress response are most often examined through analysis of blood constituents. In addition to, and as a consequence of, the hormonal changes described earlier, one typically sees ketosis, hyperlipemia, hyperglycemia, hyperaminoacidemia, and metabolic acidosis signaling increased hepatic gluconeogenesis, and lipid and protein catabolism; hematological changes follow the expected pattern. Exertional stress during capture and handling can lead to muscle damage and the release of diagnostically useful indicators such as creatine kinase, aminotransferases, and potassium (St. Aubin et al., 1979; Marsh and Anderson, 1983) (see Chapter 19, Clinical Pathology). Capture myopathy, and its pathognomonic signs (Spraker, 1993), should be considered following any procedure involving wildlife, including marine mammals.
FIGURE 2 Results of acute vs. chronic stress.
0839_frame_13fm Page 263 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
263
Chronic Response Chronic stress may occur if stressors are frequent, intermittent, and/or repetitive. Chronic stress can produce one of three responses: (1) habituation, in which the stress response decreases with each episode; (2) sensitization, where the stress response increases with each episode; or (3) desensitization, when there is no change (Dantzer and Mormede, 1995). In chronic stress, there is sustained activation of the HPA axis, producing repetitive, pulsatile secretions of glucocorticoids. The chronic effects of stress are difficult to diagnose, and even more difficult to relate back to specific stressful events. Nevertheless, it is a task commonly presented to medical professionals and biologists. In reality, chronic stress is probably of greater significance in terms of an animal’s well-being than short-term responses to transient stimuli. Impaired growth and reproduction, frequent infection, and pathological changes in organs are among the many consequences that can be linked to chronic stress. Stress can disrupt reproductive functions in many mammalian species. CRF, ACTH, glucocorticoids, and β-endorphins secreted in response to stressful stimuli can inhibit reproductive processes (Moberg, 1987b). Stress-induced elevations of glucocorticoids may affect the reproductive system by inhibiting hypothalamic secretion of gonadotropin-releasing hormone, blocking the release of luteinizing hormone (LH) and follicle-stimulating hormone (FSH), and altering the gonadal response to LH and FSH secretion from the anterior pituitary (Rivier and Rivest, 1991). At present, there is no specific information on these pathways in marine mammals. Furthermore, one can only speculate about the long-term consequences of lowered TH levels on growth and development in species such as belugas, in which TH can be substantially altered by stress (St. Aubin and Geraci, 1988; 1992). The immune system has many cellular components useful in measuring chronic stress. In vitro, mitogens such as PHA, concanavalin A, and pokeweed mitogen act as nonspecific stimulators of immune function, causing lymphocyte proliferation and activation. Such tests have been used in killer whales (Orcinus orca), bottlenose dolphins, harbor seals, and gray seals, to gauge health (DiMolfetto-Landon et al., 1995; Erickson et al., 1995; Nielsen, 1995; Blanchard et al., 1999; Hall et al., 1999) (see Chapter 12, Immunology). Failure of lymphocytes to respond to mitogens can be an indicator of severe immune system deficiency, possibly as a result of stress. A young gray seal with elevated cortisol levels showed no response to intradermal PHA, and died 12 hours later of a respiratory infection (Hall et al., 1999). Additional research is needed to determine how reliable or sensitive such indicators might be in marine mammals. Cowan and Walker (1979) suggested that a variety of pathological changes in spinner and spotted dolphins killed in dolphin–fishery interactions were related to stress. They noted massive cardiac response to stress in some of the dolphins, and described the microscopic pathological lesions as consistent with those in laboratory animals injected with catecholamines and in humans with stress cardiomyopathy. Adrenal glands are an obvious site to examine for morphological evidence of chronic stimulation. Several cetacean species necropsied after stranding, including Atlantic white-sided dolphins (Lagenorhynchus acutus), harbor porpoises (Phocoena phocoena), belugas, and a common dolphin (Delphinus delphis), had adrenocortical cysts on necropsy exam (Geraci et al., 1978; Kuiken et al., 1993; Cartee et al., 1995) (see Chapter 23, Noninfectious Diseases). The adrenal glands from 95% of 90 spinner dolphins and 172 spotted dolphins chased during capture showed darkened adrenal cortices, which were interpreted as a consequence of continuous acute stress and/or vasogenic shock leading to death (Myrick and Perkins, 1995). Belugas from the St. Lawrence River have a high prevalence of adrenal lesions, including cortical hyperplasia, cortical and medullary nodular hyperplasia, and serous cysts, which were
0839_frame_13fm Page 264 Tuesday, May 22, 2001 11:10 AM
264
CRC Handbook of Marine Mammal Medicine
increasingly common in older whales (Lair et al., 1997). Chronic exposure to organohalogens was suggested as an underlying cause of adrenal hyperfunction in this species (De Guise et al., 1994) (see Chapter 22, Toxicology). These compounds are highly toxic in vitro to adrenal mitochondria from gray seals, inhibiting glucocorticoid-synthesizing enzymes and leading to adrenal hyperplasia (Lund, 1994). Associations are frequently made among overwhelming, but nonspecific, pathological changes in free-ranging marine mammals and the stresses imposed by a contaminated environment. Bergman (1999) described adrenocortical hyperplasia in gray and ringed seals found dead along the shores of the Baltic Sea. The animals also exhibited a variety of lesions, including claw and digit deformities, bone lesions, particularly around the teeth, overburdens of acanthocephalans (Corynosoma spp.) in the proximal colon, intestinal ulcers, arteriosclerosis of the aorta and its bifurcations, and uterine leiomyomas, stenosis, and occlusion. The adrenal changes may have been a consequence of exposure to endocrine-disrupting compounds and the stress of multisystemic disease. At the same time, adrenal hyperactivity might have further compromised an immune system already suppressed by environmental contaminants (de Swart et al., 1994; Ross et al., 1996). Zenteno-Savin et al. (1997) examined circulating levels of haptoglobins (Hp) as potential indicators of chronic stress in harbor seals and Steller sea lions (Eumetopias jubatus) from declining populations in Prince William Sound, Alaska. Elevated levels of these proteins had been demonstrated in river otters (Lutra canadensis) 1 year after the Exxon Valdez oil spill there, and were felt to be linked to that event (Duffy et al., 1993; 1994). Levels in the Prince William Sound harbor seals and Steller sea lions were higher than those from the more stable populations of southeast Alaska, and were associated with infection, inflammation, trauma, and tumors in the former groups. Recently, Southern (2000) identified a group of 30 stress-responsive proteins (SRP) with recognized roles in oxidative cell response, active cell death, cell growth and differentiation, cell adhesion, and immunological and neurological signaling. By using a multitarget antibody cocktail, the suite of SRPs can be simultaneously detected in tissues, including readily available epidermal biopsies. In a survey of seven species of cetaceans, tenfold or greater increases in SRP levels were noted in animals stressed by conditions such as ice entrapment, chronic illness, starvation, net capture, and coastal pollution (Southern, 2000). The SRP assay system shows great potential for monitoring the impacts of conservation and management strategies on marine mammals.
Future Research Marine mammal stress research has advanced considerably in recent years. The goals of stress research are twofold: First, to conduct interdisciplinary studies of the interactions among endocrine, immune, and neurological systems that maintain homeostasis, control acute stress, and respond to distress; and, second, to develop a broad database for indicators that will improve the ability to recognize and manage stress in the animals both in captivity and in the wild. To this end, further research is needed in the following areas: • • • • • •
Glucocorticoid metabolism; The effects of age and gender on the stress response; Differences among species with varying sensitivities to stress; New and creative diagnostic tests that can reliably detect stress; Rational prophylaxis and treatment for stressed marine mammals; The ways environmental pollutants act as stressors or interfere with the stress response;
0839_frame_13fm Page 265 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
265
• Reproductive physiology and stress; • Marine mammal population dynamics in relation to environmental stressors; • Correlations between pathological conditions and stressors.
Conclusion In virtually every clinical situation, stress and its consequences must be addressed, since disease itself is a stressor, and stress may be at the root of the illness in question. Nevertheless, the term is too often applied indiscriminately as a convenient “catch-all” when efforts to reach some other diagnosis fall short. Advancement of understanding of this important determinant of marine mammal health will depend on a focused, scientific approach to stress and the stress response.
Acknowledgments The authors thank Mona Haebler and Barbara Curry for reviewing an earlier version of this chapter and offering helpful suggestions for improving its content. Special thanks are due from the primary author (St. Aubin) to Joseph Geraci for the many long discussions about stress and what it means in marine mammals. Funding for studies on catecholamine research in belugas was provided by the Office of Naval Research to the primary author (St. Aubin) and to Joseph Geraci. This is contribution number 125 of the Sea Research Foundation.
References Bergman, A., 1999, Health condition of the Baltic grey seal (Halichoerus grypus) during two decades, APMIS (Denmark), 107: 270–282. Blanchard, M., Lepper, H., Ferrick, D., Stott, J., Aldridge, B., Beusse, D., Dover, S., Odell, D., Walsh, M., Dalton, L., Robeck, T., McBain, J., Reidarson, T., Yochem, P., Pledger, T., and Renner, M., 1999, Lymphocyte functional analysis as an indicator of killer whale (Orcinus orca) health, in Proceedings of the 30th International Association for Aquatic Animal Medicine Conference, Boston, MA, May 2–5, pp. 29–30. Breazile, J.E., 1987, Physiologic basis and consequences of distress in animals, J. Am. Vet. Med. Assoc., 191: 1212–1215. Breazile, J.E., 1988, The physiology of stress and its relationship to mechanisms of disease and therapeutics, Vet. Clin. North Am. Food Anim. Pract., 4(3): 441–480. Cartee, R.E., Tarpley, R., Mahoney, K, Ridgway, S.H., and Johnson, P., 1995, A case of cystic adrenal disease in a common dolphin (Delphinus delphis), J. Zoo Wildl. Med., 26: 293–297. Chrousos, G.P., 1988, The concept of stress and its historical development, in Adv. Exp. Med. Biol. Mech. Phys. Emot. Stress, Chrousos, G.P, Loriaux, D.L., and Gold, P.W. (Eds.), 245: 3–7. Chrousos, G.P., and Gold, P.W., 1992, The concepts of stress and stress system disorders, J. Am. Med. Assoc., 267: 1244–1252. Cowan, D.F., 2000, The histopathology of the alarm reaction in dolphins, in Proceedings of the American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine Conference, New Orleans, LA, Sept. 17–21, pp. 195–196. Cowan, D.F., and Walker, W.A., 1979, Disease factors in Stenella attenuata and Stenella longirostris taken in the eastern tropical Pacific yellowfin tuna purse seine fishery, Administrative Report #LJ-7932C, Southwest Fisheries Science Center, NMFS, NOAA, La Jolla, CA, 21 pp. Curry, B.E., 1999, Stress in mammals: The potential influence of fishery-induced stress on dolphins in the eastern tropical Pacific Ocean, NOAA Technical Memorandum NMFS, NOAA-TM-NMFSSWFSC-260, Southwest Fisheries Science Center, La Jolla, CA, 121 pp.
0839_frame_13fm Page 266 Tuesday, May 22, 2001 11:10 AM
266
CRC Handbook of Marine Mammal Medicine
Dantzer, R., and Mormede, P., 1995, Psychoneuroimmunology of stress, in Stress, the Immune System and Psychiatry, Leonard, B., and Miller, K. (Eds.), John Wiley & Sons, West Sussex, U.K., 47–53. De Guise, S., Legace, A., and Beland, P., 1994, Tumors in St. Lawrence beluga whales (Delphinapterus leucas), Vet. Pathol., 31: 441–449. de Swart, R.L., Ross, P.S., Vedder, L.J., Timmerman, H.H., Heister-Kamp, S.H., Van Lowren, H., Vos, J.G., Reijnders, P.J.F., and Osterhaus, A.D.M.E., 1994, Impairment of immune function in harbour seals (Phoca vitulina) feeding on fish from polluted waters, Ambio, 23: 15–59. DiMolfetto-Landon, L., Erickson, K.L., Blanchard-Channell, M., Jeffries, S.J., Harvey, J.T., Jessup, D.A., Ferrick, D.A., and Stott, J.L., 1995, Blastogenesis and interleukin-2 receptor expression assays in the harbor seal (Phoca vitulina), J. Wildl. Dis., 31(2): 150–158. Duffy, L.K., Bowyer, R.T., Testa, J.W., and Faro, J.B., 1993, Differences in blood haptoglobin and lengthmass relationships in river otters (Lutra canadensis) from oiled and non-oiled areas of Prince William Sound, Alaska, J. Wildl. Dis., 29(2): 353–359. Duffy, L.K., Bowyer, R.T., Testa, J.W., and Faro, J.B., 1994, Evidence for recovery of body mass and haptoglobin values of otters following the Exxon Valdez oil spill, J. Wildl. Dis., 30(3): 421–425. Dunn, A.J., 1995, Psychoneuroimmunology: Introduction and general perspectives, in Stress, the Immune System and Psychiatry, Leonard, B., and Miller, K. (Eds.), John Wiley & Sons, West Sussex, U.K., 1–17. Dunn, A.J., 1996, Psychoneuroimmunology, stress and infection, in Psychoneuroimmunology, Stress, and Infection, Friedman, H., Klein, T.W., and Friedman, A.L. (Eds.), CRC Press, Boca Raton, FL, 25–46. Engelhardt, F.R., and Geraci, J.R., 1978, Effects of experimental vitamin E deprivation in the harp seal, Phoca groenlandica, Can. J. Zool., 56: 2186–2193. Erickson, K.L., DiMolfetto-Landon, L., Wells, R.S., Reidarson, T., Stott, J.L., and Ferrick, D.A., 1995, Development of an interleukin-2 receptor expression assay and its use in evaluation of cellular immune responses in bottlenose dolphins (Tursiops truncatus), J. Wildl. Dis., 31(2): 142–149. Fothergill, M.B., Schwegman, P.A., Garratt, P.A., Govender, A., and Robertson, W.D., 1991, Serum alkaline phosphatase-changes in relation to state of health and age of dolphins, Aquat. Mammals, 17: 71–75. Geraci, J.R., 1972a, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618–623. Geraci, J.R., 1972b, Experimental thiamine deficiency in captive harp seals Phoca groenlandica induced by eating herring Clupea harrengus and smelts Osmerus mordax, Can. J. Zool., 50: 179–195. Geraci, J.R., and St. Aubin, D.J., 1987, Cetacean mass stranding: A study into stress and shock, in Proceedings of the 7th Biennial Conference on the Biology of Marine Mammals, Miami, FL, Dec. 5–9, p. 25. Geraci, J.R., and St. Aubin, D.J. (Eds.), 1990, Sea Mammals and Oil: Confronting the Risks, Academic Press, San Diego, CA, 282 pp. Geraci, J.R., and Smith, T.G., 1975, Functional hematology of ringed seals (Phoca hispida) in the Canadian Arctic, J. Fish. Res. Board Can., 32: 2559–2564. Geraci, J.R., and Smith, T.G., 1976, Direct and indirect effects of oil on ringed seals (Phoca hispida) of the Beaufort Sea, J. Fish. Res. Board Can., 33: 1976–1984. Geraci, J.R., Testaverde, S.A., St. Aubin, D.J., and Loop, T.H., 1978, A Mass Stranding of the Atlantic White-Sided Dolphin, Lagenorhynchus acutus: A Study into Pathobiology and Life History, National Technical Information Service, Springfield, VA, PB289-361, 141. Geraci, J.R., St. Aubin, D.J., and Smith, T.G., 1979, Influence of age, condition, sampling time, and method on plasma chemical constituents in free-ranging ringed seals, Phoca hispida, J. Fish. Res. Board Can., 36: 1278–1282. Goldstein, D.S., 1995, Stress and science, in Stress, Catecholamines and Cardiovascular Disease, Goldstein, D.S. (Ed.), Oxford University Press, Oxford, U.K., 3–55.
0839_frame_13fm Page 267 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
267
Gulland, F.M.D., Haulena, M., Lowenstine, L.J., Munro, C., Graham, P.A., Bauman, J., and Harvey, J., 1999, Adrenal function in wild and rehabilitated Pacific harbor seals (Phoca vitulina richardsi) and in seals with phocine herpesvirus-associated adrenal necrosis, Mar. Mammal Sci., 15(3): 810–827. Hall, A.J., Licence, S.T., and Pomeroy, P.P., 1999, The response of grey seal pups to intradermal phytohaemagglutinin injection, Aquat. Mammals, 25(1): 25–30. Keller, S.E., Schleifer, S.J., and Demetrikopoulos, M.K., 1991, Stress-induced changes in immune function in animals: Hypothalamic-pituitary-adrenal influences, in Psychoneuroimmunology, 2nd ed., Ader, R., Felten, D.L., and Cohen, N. (Eds.), Academic Press, San Diego, CA, 771–787. Kuiken, T., Hofle, U., Bennett, P.M., Allchin, C.R., Kirkwood, J.K., Baker, J.R., Appleby, E.C., Lockyer, C.H., Walton, M.J., and Sheldrick, M.C., 1993, Adrenocortical hyperplasia, disease, and chlorinated hydrocarbons in the harbour porpoise (Phocoena phocoena), Mar. Pollut. Bull., 26(8): 440–446. Lair, S., Beland, P., De Guise, S., and Martineau, D., 1997, Adrenal hyperplasia and degenerative changes in beluga whales, J. Wildl. Dis., 33(3): 430–437. Larsen, P.R., Davies, T.F., and Hay, I.D., 1998, The thyroid gland, in Williams Textbook of Endocrinology, Wilson, J.D., Foster, D.W., Kronenberg, H.M., and Larsen, P.R. (Eds.), W.B. Saunders, Philadelphia, 389–516. Leonard, B., and Miller, K. (Eds.), 1995, Stress, the Immune System, and Psychiatry, John Wiley & Sons, West Sussex, U.K., 238 pp. Levine, S., 1993, The influence of social factors on the response to stress, Psychother. Psychosom., 60: 33–38. Lovallo, W.R., 1997a, Physiological regulation during physical and psychological stress, in Stress and Health: Behavioral and Psychological Interactions, Lovallo, W.R. (Ed.), Sage Publications, Thousand Oaks, CA, 55–74. Lovallo, W.R., 1997b, Psychological stress response, in Stress and Health: Behavioral and Psychological Interactions, Lovallo, W.R. (Ed.), Sage Publications, Thousand Oaks, CA, 75–100. Lund, B.O., 1994, In vitro adrenal bioactivation and effects on steroid metabolism of DDT, PCBs and their metabolites in the gray seal (Halichoerus grypus), Environ. Toxicol. Chem., 13(6): 911–917. Marsh, H., and Anderson, P.K., 1983, Probable susceptibility of dugongs to capture stress, Biol. Conserv., 25: 1–3. McEwen, B.S., Biron, C.A., Brunson, K.W., Bullock, K., Chambers, W.H., Dhabhar, F.S., Goldfarb, R.H., Kitson, R.P., Miller, A.H., Spencer, R.L., and Weiss, J.M., 1997, The role of adrenocorticoids as modulators of immune function in health and disease: neural, endocrine and immune interactions, Brain Res. Rev., 23: 79–199. Medway, W., and Geraci, J.R., 1964, Hematology of the bottlenose dolphin (Tursiops truncatus), Am. J. Physiol., 207: 1367–1370. Medway, W., Geraci, J.R., and Klein, L.V., 1970, Hematologic response to administration of a corticosteroid in the bottle-nosed dolphin (Tursiops truncatus), J. Am. Vet. Med. Assoc., 157: 563–565. Moberg, G.P., 1985, Biological response to stress: Key to assessment of animal well-being, in Animal Stress, Moberg, G.P. (Ed.), American Physiological Society, Bethesda, MD, 27–49. Moberg, G.P., 1987a, Problems of defining stress and distress in animals, J. Am. Vet. Med. Assoc., 191: 1207–1211. Moberg, G.P., 1987b, Influence of the adrenal axis upon the gonads, Oxford Rev. Reprod. Biol., 6: 456–496. Munck, A., Guyre, P.M., and Holbrook, N.J., 1984, Physiological functions of glucocorticoids in stress and their relation to pharmacological actions, Endocr. Rev., 5: 25–44. Myrick, A.C., and Perkins, P.C., 1995, Adrenocortical color darkness and correlates as indicators of continuous acute postmortem stress in chased and purse-seined captured male dolphins, Pathophysiology, 2: 191–204. Neylan, T.C., 1998, Hans Selye and the field of stress research (includes a reprint of Selye’s original 1936 article), Neurophysiol. Classics, 10(2): 230–231.
0839_frame_13fm Page 268 Tuesday, May 22, 2001 11:10 AM
268
CRC Handbook of Marine Mammal Medicine
Nielsen, J., 1995, Immunological and hematological parameters in captive harbor seals (Phoca vitulina), Mar. Mammal Sci., 11(3): 314–323. Norris, K.E., and Dohl, T.P., 1980, The behavior of the Hawaiian spinner dolphin, Stenella longirostris, Fish. Bull., 77: 821–849. Norris, K.E., Stuntz, W.E., and Rogers, W., 1978, The behavior of porpoises and tuna in eastern tropical Pacific yellowfin tuna fishery-preliminary studies, Marine Mammal Commission, MMC-76-12, 86 pp. OED, 1999, Oxford English Dictionary, Oxford University Press, Oxford, U.K., 885–886. Orlov, M.V., Mukhlya, A.M., and Kulikov, N.A., 1988, Hormonal indices in the bottle-nosed dolphin Tursiops truncatus in the norm and in the dynamics of experimental stress, Sov. J. Evol. Biochem. Physiol., 24: 431–436. Ortiz, R.M., and Worthy, G.A.J., 2000, Effects of capture on plasma adrenal steroids and vasopressin levels in free-ranging bottlenose dolphins (Tursiops truncatus), Comp. Biochem. Physiol. A, 125: 317–324. Reidarson, T.H., and McBain, J., 1999, Hematologic, biochemical and endocrine effects of dexamethasone on bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med., 30(2): 310–312. Ridgway, S.H., and Patton, G.S., 1971, Dolphin thyroid: Some anatomical findings, Z. Vgl. Physiol., 71: 129–141. Rivier, C., 1991, Role of interleukins in the stress response, Methods Achiev. Exp. Pathol., 14: 63–79. Rivier, C., and Rivest, S., 1991, Review: Effect of stress on the activity of the hypothalamic-pituitarygonadal axis-periperal and central mechanisms, Biol. Reprod., 45: 523–532. Romano, T.A., Felten, S.Y., Olschowka, J.A., and Felten, D.L., 1994, Noradrenergic and peptidergic innervation of lymphoid organs in the beluga, Delphinapterus leucas: An anatomical link between the nervous and immune systems, J. Morphol., 221: 243–259. Ross, P.S., de Swart, R.L., Timmerman, H.H., Vedder, L.J., van Loveren, H., Vos, J.G., Riejnder, P.J.H., and Osterhaus, A.D.M.E., 1996, Suppression of natural killer cell activity in harbor seals (Phoca vitulina) fed Baltic Sea herring, Aquat. Biol., 34: 71–84. Rushen, J., 1986, Some problems with the physiological concept of “stress,” Aust. Vet. J., 63: 359–361. St. Aubin, D.J., 1990, Physiologic and Toxicologic Effects on Pinnipeds, in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, San Diego, CA, 103–127. St. Aubin, D.J., and Geraci, J.R., 1986, Adrenocortical function in pinniped hyponatremia, Mar. Mammal Sci., 2: 243–250. St. Aubin, D.J., and Geraci, J.R., 1988, Capture and handling stress suppresses circulating levels of thyroxine (T4) and triiodothyronine (T3) in beluga whales, Delphinapterus leucas, Physiol. Zool., 61(2): 170–175. St. Aubin, D.J., and Geraci, J.R., 1989, Adaptive changes in hematologic and plasma chemical constituents in captive beluga whales, Delphinapterus leucas, Can. J. Fish. Aquat. Sci., 46: 796–803. St. Aubin, D.J., and Geraci, J.R., 1990, Adrenal responsiveness to stimulation by adrenocorticotropic hormone (ACTH) in captive beluga whales, Delphinapterus leucas, in Advances in Research on the Beluga Whale, Delphinapterus leucas, Can. Bull. Fish. Aquat. Sci., 224: 149–157. St. Aubin, D.J., and Geraci, J.R., 1992, Thyroid hormone balance in beluga whales, Delphinapterus leucas: Dynamics after capture and influence of thyrotropin, Can. J. Vet. Res., 56: 1–5. St. Aubin, D.J., Austin, T.P., and Geraci, J.R., 1979, Effects of handling stress on plasma enzymes in harp seals, Phoca groenlandica, J. Wild. Dis., 15: 569–572. St. Aubin, D.J., Ridgway, S.H., Wells, R.S., and Rhinehart, H., 1996, Dolphin thyroid and adrenal hormones: Circulating levels in wild and semidomesticated Tursiops truncatus, and influence of sex, age, and season, Mar. Mammal Sci., 12(1): 1–13. Selye, H., 1936, A syndrome produced by diverse nocuous agents, Nature, 138: 32.
0839_frame_13fm Page 269 Tuesday, May 22, 2001 11:10 AM
Stress and Marine Mammals
269
Southern, S., 2000, Molecular analysis of stress-activated proteins and genes in dolphins and whales: A new technique for monitoring environmental stress, in Proceedings of the American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine Conference, New Orleans, LA, Sept. 17–21, pp. 240–243. Spraker, T.R., 1993, Stress and capture myopathy in artiodactylids, in Zoo and Wild Animal Medicine, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 481–488. Thomas, J.A., Kastelein, R.A., and Awbrey, F.T., 1990, Behavior and blood catecholamines of captive beluga whales during playbacks of noise from an oil drilling platform, Zoo Biol., 9: 393–402. Thomson, C.A., and Geraci, J.R., 1986, Cortisol, aldosterone, and leucocytes in the stress response of bottlenose dolphins, Tursiops truncatus, Can. J. Fish. Aquat. Sci., 43: 1010–1016. Turnbull, B.S., and Cowan, D.F., 1998, Myocardial contraction band necrosis in stranded cetaceans, J. Comp. Pathol., 118(4): 317–327. Zenteno-Savin, T., Castellini, M.A., Rea, L.D., and Fadely, B.S., 1997, Plasma haptoglobin levels in threatened Alaskan pinniped populations, J. Wildl. Dis., 3(1): 64–71.
0839_frame_13fm Page 270 Tuesday, May 22, 2001 11:10 AM
0839_frame_C14.fm Page 271 Tuesday, May 22, 2001 11:11 AM
14
Genetic Analyses Deborah A. Duffield and William Amos
Introduction This chapter explores how genetic techniques can contribute to understanding of marine mammals and their problems, with special emphasis on marine mammal strandings and maintenance and breeding of marine mammals in captivity. The chapter outlines genetic methodologies available, attempting to concentrate on those methods used most often. Also included are brief descriptions of the processes for sampling animals that strand.
Genetic Techniques The literature contains references to a wide and even bewildering range of genetic techniques, reflecting a restless search for greater resolution, robustness, comparability, and ease of use. A good review of the various techniques, including DNA analysis, is given in Hillis et al. (1996). It was as late as 1960 that starch gel electrophoresis was first used to reveal and quantify genetic variability in the form of protein polymorphisms. Bypassing the need to look at the genes themselves or their products, data were also collected from differences among chromosomes revealed by various staining techniques. In about 1970, with the discovery of restriction enzymes that recognize and cut particular DNA motifs, the world of DNA analysis opened. In its most basic form, the presence or absence of a cutting site yields either one long or two shorter fragments, known as restriction fragment length polymorphisms (RFLP). RFLP analysis is a generic method for revealing polymorphism that is still used widely in one form or another. More recently, two primary approaches have come to dominate the scene, and these appear to be poised to stay for most applications.
DNA Sequencing The first technique is DNA sequencing. Although once laborious and expensive, DNA sequencing is now rapid, accessible, and cheap. The most commonly sequenced genes are those with particularly attractive features, and at top of the list are two or three genes found on the DNA in mitochondria (Moritz, 1994). Mitochondria are the modern descendants of ancient bacteria that came to live inside the cells of higher organisms, and each mitochondrion carries its own degenerate, circular chromosome. Mitochondrial DNA (mtDNA) is a popular target for sequencing because of several unusual properties. First, mtDNA evolves very rapidly and, hence, even populations and closely related species tend to carry diagnostic differences. Second, every cells carries hundreds or even thousands of mitochondria, and therefore there are many more 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
271
0839_frame_C14.fm Page 272 Tuesday, May 22, 2001 11:11 AM
272
CRC Handbook of Marine Mammal Medicine
copies of an mtDNA gene compared with an equivalent gene present as a single copy in the cell nucleus. This high copy number can allow genetic tests even when most of the DNA has been degraded by putrefaction or antiquity (Hagelberg, 1994; Hagelberg et al., 1994). Third, in most higher organisms mitochondria are inherited strictly through the female line, giving a simple pattern of inheritance, which tends to reveal differences between populations more strongly than most other genetic markers. Fourth, even within the tiny mitochondrial genome there are regions that evolve at different rates. The fastest-evolving sequences are found in a region with little clear function known as the D-loop or control region. Evolving some five to ten times slower, are any one of a number of genes coding for mitochondrial proteins, of which a common target is called cytochrome oxidase, or CO1; another is cytochrome b.
“Tandem Repeats” and DNA Fingerprinting The second main technique involves an unusual class of DNA sequences called tandem repeats that show extreme levels of polymorphism. The term tandem repeat embraces any short DNA motif repeated head to tail from a few to hundreds or thousands of times, (e.g., ACCACCACCACCACCACC). Most exciting was the discovery in 1985 of medium-sized repeats called “minisatellites,” which show the greatest variability of all, and form the basis of the technique known popularly as DNA fingerprinting (Jeffreys et al., 1985a). DNA fingerprinting is a remarkably powerful technique, able to identify individuals uniquely and able to assign unambiguous parentage (Jeffreys et al., 1985b; Amos et al., 1993). Unfortunately, the technique also proved technically difficult to apply and has since been almost completely replaced by an alternative approach based on the smaller repeats, christened with simple logic microsatellites. Since its potential was discovered in 1989 (Litt and Luty, 1989; Tautz, 1989), microsatellite analysis has grown to a dominant position in the literature, being both accessible and powerful. Microsatellites are the shortest possible tandem repeats, with the repeating unit usually two to five DNA letters long, for example, ACACACACAC (see reviews in Bruford and Wayne, 1993; Bruford et al., 1996; Goldstein and Schlötterer, 1999). Microsatellites are attractive markers for several reasons. First, they are highly polymorphic, typically carrying 5 to 10 alleles/locus. By combining information from several loci, this is sufficient to allow a range of analyses, from the identification of individuals (Jeffreys et al., 1992) and parentage testing (Worthington Wilmer et al., 1999) to the detection of differences between populations and species (Paetkau et al., 1995). Second, microsatellites are assayed by the polymerase chain reaction (PCR). In PCR, an enzyme is used repeatedly to make copies of a target piece of DNA, identified by its sequence. The result is an immensely powerful tool that can analyze as little as a single molecule, making the technique ideal for dealing with the sort of DNA one can extract from decaying or degraded tissue (Reed et al., 1997; Taberlet et al., 1997). Third, microsatellite analysis is easy to use and yields data that are ideally suited to inclusion in databases (each allele is recorded as a fragment with a discrete length). A slight drawback is that some preparatory work is needed to develop microsatellite markers for each new species, although with more and more studies appearing and with many markers working on related species (Valsecchi and Amos, 1996; Gemmell et al., 1997), this problem is resolving.
Genetic Analyses Applied to Stranded Marine Mammals A stranded marine mammal can provide material for genetic analysis that can elucidate many aspects of a species biology. For marine mammals, many of which live in inaccessible regions and spend much of their lives out of view below the sea surface, information gained through genetics can play an even greater role than for more easily studied terrestrial species. The sorts
0839_frame_C14.fm Page 273 Tuesday, May 22, 2001 11:11 AM
Genetic Analyses
273
of questions that can be addressed include identification, from species through population down to individual identity; studies of social organization, based on determining the relationships among individuals in a group; and diagnosing the cause of death by, for example, using gene sequencing to identify a particular pathogen.
Species Identification Genetic analysis is most obviously useful for determining the species from which a sample has been collected, and the primary technique used here is gene sequencing. Much progress has been made, largely as a result of pioneering studies by Baker and others aimed at identifying the origin of whale products sold in food markets in Japan and Korea (Baker et al., 1996). Results showed that much of the meat was from minke whales (Balaenoptera acutorostrata) taken under license for scientific whaling, but that significant numbers of samples could be attributed to protected species, including at least two different individual blue whales (B. musculus). There is now an almost complete catalog of mtDNA sequences available, with all but a handful of extant species represented, along with most of the major populations. Any new, unidentified specimen can be matched with great precision, essentially always to species, and often to the ocean basin it came from. Many fresh strandings provide material that can be identified with high confidence based on morphological traits. In such cases, DNA sequencing can have two functions. First, it provides a useful double-check for field misidentification. Cetacean coloration can change rapidly after death, making identification difficult. Even when a nominal species has been accurately determined, several instances have emerged where genetic analysis has revealed the presence of cryptic species, subspecies, or races. Second, the more sequences that can be added to the database, the more complete the database becomes, thereby facilitating future matches. This is particularly important for rarer species whose distribution may be poorly understood, and for species with strong population structure, where a more complete database can be used to pinpoint an animal’s origin. It is important not to forget that a dead marine mammal may contain more than one species. Parasites, bacteria, and viruses also contain DNA, which can be used for their identification. In 1988, large numbers of harbor seals (Phoca vitulina) were found washed up dead and dying, first around Denmark and then spreading up around the North Sea coasts to Scotland and Ireland (Swinton et al., 1998). In some areas, more than 50% of all seals died (see Chapter 15, Viral Diseases). The cause was initially a mystery, although the acute respiratory distress and secondary infections were suggestive of canine distemper. DNA sequences obtained from viral isolates later proved to be from a new pathogen known as phocine distemper virus. Similar, although less spectacular, mortality events in porpoises, dolphins, and (possibly) monk seals yielded further members of this viral family (Barrett et al., 1993); morbilliviruses are now prime suspects when marine mammals start dying in large numbers (see Chapter 2, Emerging Diseases).
Population Identification Below the level of species, one is interested in identification of the population or stock from which a given individual derives. Such questions are an ongoing concern with marine mammals because of their great capacity for movement (Dizon et al., 1997); threats posed in one area can exert strong influences elsewhere. Until the patterns of movement of each species, and how to recognize where one population ends and the next one begins, are better understood, any attempts at management or conservation will be difficult.
0839_frame_C14.fm Page 274 Tuesday, May 22, 2001 11:11 AM
274
CRC Handbook of Marine Mammal Medicine
Population studies usually involve either mtDNA sequencing (mainly of the fast-evolving D-loop or control region) or microsatellite analysis (Allen et al., 1995), although protein polymorphisms historically played an important part. Together these techniques have helped to elucidate patterns of movement of many species, from great whales (Baker et al., 1990; Palsbøll et al., 1995) and belugas (Delphinapterus leucas) (O’Corry-Crowe et al., 1997) to manatees, seals (Burg et al., 1999), and sea otters (Cronin et al., 1996). Size is no predictor of where divisions will exist. Thus, while sperm whales (Physeter macrocephalus) show little evidence of population structure throughout the world oceans (Lyrholm and Gyllensten, 1998), humpback whales (Megaptera novaeangliae) exhibit very strong structure, because of the way offspring learn their mothers’ patterns of movements (Baker et al., 1990). Other interesting examples include the harbor seal, in which great individual mobility belies strong genetic isolation between most breeding colonies (Goodman, 1998), and killer whales (Orcinus orca), in which two behaviorally and genetically distinct groups of the same nominal species coexist in the same area off the Washington coast (Hoelzel and Dover, 1991). Although most studies looking for evidence of population substructure use either mtDNA or microsatellites, a few use both mitochondrial and nuclear markers (Burg et al., 1999). The advantage of using both markers together is that their contrasting modes of inheritance can indicate sex-specific patterns of gene flow. In many mammals, females tend to stay to breed near their natal site, whereas males disperse to avoid inbreeding. Here, mtDNA sequences, inherited solely through the female line, will show a pattern of strong substructure, reflecting the lack of movement by females between sites. However, microsatellites are nuclear markers and alleles are inherited from both parents. Consequently, even though only males move between sites, this will provide sufficient mixing to reduce or even eliminate evidence of substructure. By using the two markers together, it becomes possible to deduce these sex-based differences in dispersal behavior; whenever mtDNA shows strong substructure while microsatellites do not, this is good evidence that females return to breed where they were born, whereas males tend to disperse (Palumbi and Baker, 1994).
Social Organization The primary tool for examining questions about relatedness and social organization is microsatellite analysis. These markers are eminently suitable for identifying individuals, calculating indices of relatedness, and conducting parentage analysis, and they have the particular advantage that they can be genotyped in older, more degraded samples, including museum specimens. Genetic identity can be used to track individual movements in just the same way that early workers implanted “discovery tags,” large metal projectiles that lodged inside a whale’s body and were later recovered and recorded when that whale was subsequently killed (Palsbøll et al., 1997). With the advent of biopsy darting, genetic “tagging” is now a practical way to follow an individual throughout its life. As the number of studies increases, the chances improve that a sample from a meat market, stranding, or net entanglement will provide an informative last data point. Studies of parentage and relatedness using genetic analysis provide vital information that allows reconstruction of the social organization and breeding behavior of any organism, but they are particularly important for inaccessible marine mammals. Individual projects can be based on as few as two or three animals that strand together, but range in size to long-term studies with directed sampling, where databases of thousands of individuals now exist. Many revelations are emerging. These include the relative lack of polygynous behavior in gray seals (Halichoerus grypus) (Ambs et al., 1999; Worthington Wilmer et al., 1999), with some individuals even showing partner fidelity (Amos et al., 1995) and the dissection of social groups of cetaceans, such as pilot whales (Amos et al., 1993).
0839_frame_C14.fm Page 275 Tuesday, May 22, 2001 11:11 AM
Genetic Analyses
275
There is a further use of microsatellites that is of particular interest to veterinarians. Since every individual inherits one allele from each parent, the similarity of alleles at a locus provides a measure of the degree of parental similarity. Thus, highly inbred individuals will tend to carry pairs of alleles that are very similar to each other, whereas animals born to genetically dissimilar parents will tend to carry dissimilar alleles. By using this logic, studies on red deer (Cervus elaphus) (Coulson et al., 1998), harbor seals (Coltman et al., 1998), and Soay sheep (Aries aries) (Coltman et al., 1999) have used molecular estimates of parental similarity to show that the level of inbreeding has a significant impact on fitness. Juvenile survival is greater in individuals born to more genetically dissimilar parents. It has even been shown that individuals born to dissimilar parents tend to carry lower parasite burdens as adults (Coltman et al., 1999). The possibility of a genetic explanation for at least some of the observed variation in susceptibility to infection is an exciting one that may well blossom in the near future.
Genetic Analysis Applied to Captive Maintenance and Breeding Programs Zoos and aquaria play an important role in species conservation and propagation. As wild populations dwindle, it often falls on captive breeding programs, not only to maintain captive populations, but also to reintroduce individuals to the wild (Kleiman et al., 1996). For marine mammals, successful captive breeding has been well documented with births reported in 17 species (Asper et al., 1990), including cetaceans, pinnipeds, sea otters, and manatees (see Chapter 11, Reproduction). In commonly held species, such as bottlenose dolphins (Tursiops truncatus), California sea lions (Zalophus californianus), and harbor seals, breeding groups have had second- and third-generation offspring.
Paternity Testing Maintaining genetic diversity is a primary population goal for long-term management of captive populations (Ballou and Foose, 1996). Genetic variation is important to the ability of a captive population to adapt to changing environments, as well as to help prevent loss of individual fitness due to the deleterious effects of inbreeding (Ralls et al., 1988; Lacy et al., 1993). Tracking parentage in captive propagation programs by genetic monitoring ensures that a balanced gene pool is maintained and that breeding programs avoid inbreeding. Documentation of the relationships between individuals provides valuable information for use when setting up breeding colonies and when exchanging animals or sperm for breeding or artificial insemination purposes. In most instances, a mother–offspring relationship is known, so that the evaluation of parentage usually rests on determination of paternity. Among group-living animals, paternity cannot always be reliably assigned based on social dominance or observed copulatory behavior, hence, the importance of genetic discrimination of paternity in colonies with multiple males. In the past decade, there have been significant technological advances influencing the range of molecular genetic analyses that are being used to aid breeding programs in zoos (Bruford et al., 1996; Ryder and Fleischer, 1996). Methodologies currently in use with marine mammals include protein electrophoresis; in particular, hemoglobin electrophoresis; fluorescent R-band chromosome analysis and DNA microsatellite analysis. Hemoglobin electrophoresis is inexpensive and has been useful for establishing paternity in cases where the potential sires were of different hemoglobin types (Duffield and Chamber-Lea, 1990). Similarly, cetacean chromosomes are excellent discriminators for paternity testing, because they have numerous variable regions, referred to as heteromorphisms, in their karyotypes. These are readily visualized by fluorescent R-banding (Duffield
0839_frame_C14.fm Page 276 Tuesday, May 22, 2001 11:11 AM
276
CRC Handbook of Marine Mammal Medicine
FIGURE 1 Fluorescent R-band chromosome heteromorphism analysis for bottlenose dolphin chromosome pair 19. An example of the use of fluorescent R-band chromosome heteromorphism analysis for paternity testing in bottlenose dolphins. (A) Calf. The karyotype of the offspring is screened for chromosome pairs with heteromorphic variants. (B) Mother. The heteromorphic pairs of the offspring are compared with those same pairs in the female to establish which variants were inherited from the mother. This defines the “required paternal match.” (C) Potential fathers. The karyotypes of all possible sires are compared with the offspring to determine which male has the paternal match. Given the number of heteromorphic chromosome pairs in cetacean karyotypes, each paternal discrimination is made on the basis of matching several such variants.
and Chamberlin-Lea, 1990; Duffield and Wells, 1991; Duffield et al., 1991). In contrast, the chromosomes of pinnipeds, the sea otter, and the manatee do not exhibit the degree of chromosomal heteromorphism seen in cetaceans. An example of how chromosome heteromorphism analysis is used in paternity testing in cetaceans is presented in Figure 1. With the advent of microsatellite analysis, this latter technique is becoming the DNA methodology of choice for paternity testing in captive breeding colonies. Microsatellite primer sequences have now been reported for a broad range of cetacean and pinniped species (Buchanan et al., 1996; 1998; Valsecchi and Amos, 1996; Gemmell et al., 1997; Shinohara et al., 1997). An example of paternal assignment using the microsatellite primer EV-37 (Valsecchi and Amos, 1996) in bottlenose dolphins is given in Figure 2. More than 20 different alleles for this single locus have been identified in the North American captive bottlenose dolphin population, and this amount of variability has made paternal discrimination very effective in these breeding groups.
Hybrid Detection Genetic analysis is also useful for identifying interspecies hybrids. For odontocete cetaceans, hybrids have occurred between Tursiops truncatus and Grampus griseus, T. truncatus and Steno
0839_frame_C14.fm Page 277 Tuesday, May 22, 2001 11:11 AM
Genetic Analyses
277
FIGURE 2 Example of microsatellite paternity testing in bottlenose dolphins. An example of assigning paternity with microsatellites, using primer EV-37 (Valsecchi and Amos, 1996). Alleles are represented by a top darker band, followed by fainter stutter (shadow) bands, which decrease in intensity. Microsatellite alleles in calves (C) are compared with those in the dam (D) to identify the allele given by the sire (arrows). The photograph was cropped to eliminate lanes that were not pertinent to this assignment. The sire (S) for both calves (C1 and C2) is in lane three from the left. No other animal shares his top allele (given to C1), and the animals sharing his bottom allele (given to C2) are an unrelated female and a male that was not present at the time of conception.
bredanensis, T. truncatus and Globicephala macrorhynchus, T. truncatus and Pseudorca crassidens, T. truncatus and Delphinus delphis, Phocoena phocoena and Phocoenoides dalli, and possibly between D. capensis and Lagenorhynchus obscurus (Sylvestre and Tanaka, 1985; Reyes, 1996; Baird et al., 1998; Sea World, pers. comm.). One T. truncatus and Pseudorca crassidens hybrid has had two live-born offspring, sired by bottlenose dolphin males. One of these secondgeneration hybrids survived for nearly 8 years (North American Bottlenose Dolphin Studbook). The ability to have offspring proves that these particular hybrids are fertile, an important confirmation for evaluation of naturally occurring hybrids. Live-born pinniped hybrids have been reported between Zalophus californianus and Callorhinus ursinus, Z. californianus and Arctocephalus pusillus, Z. californianus and Eumetopius jubatus, and Phoca kurilensis and P. largha (King, 1983; Kamogawa SeaWorld, pers. comm.; DeLong, pers. comm.). One of the crosses between Z. californianus and C. ursinus gave birth to two pups, sired by California sea lions, again indicating the fertility of this hybrid. One pup was live-born, but died within a few days; the second pup was stillborn. Breeding and recognition of cetacean and pinniped hybrids in captivity affords a rare opportunity to develop anatomical and genetic profiles for these hybrids, which one hopes will further recognition of interspecific crosses in the wild.
Sampling To perform genetic analysis, it is first necessary to obtain tissue samples from which to extract DNA. For live animals, there are several possible sampling routes, including blood sampling, direct tissue sampling with a biopsy device, and collection of body by-products such as sloughed skin or feces. For most stranding situations, the problem simplifies to which part of a dead animal is best to plunder, and this depends largely on how fresh the body is (see Chapter 21, Necropsy).
0839_frame_C14.fm Page 278 Tuesday, May 22, 2001 11:11 AM
278
CRC Handbook of Marine Mammal Medicine
From live animals, blood extraction provides an excellent source of DNA, both nuclear and mitochondrial. For stranded animals, probably the best all-around tissue for DNA extraction is skin. Skin has the immediate and obvious advantage of being on the outside. It is also metabolically sluggish, its cells potentially living for days after death. When fresh, skin yields sufficient good-quality DNA for any application. However, when the tissue is not fresh, skin usually offers the best chance to obtain a usable DNA sample, often weeks, months, or even years after death. When decay is advanced to the point where the skin has been lost, it may be still possible to take a useful sample by concentrating on drier parts of the body and metabolically inactive tissues such as connective tissue. Extraction of DNA from bones is often used in forensic studies, although the process is time-consuming and fallible and is thus not likely to be of use unless the specimen is particularly important. Further DNA extraction information is available in Davis et al. (1986) and Escorza et al. (1997). Sample preservation can be achieved by a variety of methods. For euthanized live stranded or zoo animals, fresh blood can be taken and is best treated by refrigeration and immediate (within 3 to 4 days) DNA extraction, or it can be frozen to −20°C (−4°F). Volumes can be reduced significantly if a low-speed centrifuge is available, allowing all but the white cells (buffy coat) to be discarded. For solid tissues, the simplest method of preservation is to freeze to −20°C. When a freezer is not available, room-temperature preservation can be achieved by use of agents that neutralize the process of degradation. Of several options, two of the most widely used are a saturated solution of table salt in 20% DMSO (dimethylsulfoxide, added to aid rapid penetration into the tissue) or alcohol (70% or above). In either case, the preservative should be added to a level of one part sample to two to three parts preservative. Less robust, although perfectly practicable, alternatives include rapid drying of the tissue and packing in an excess of dry table salt. Formalin should be avoided, since in most cases this causes irrevocable damage to the DNA.
Conclusion The field of marine mammal genetics, including technologies, methodologies, and utilities, has grown considerably over the last decade, since the publication of the first edition of this book. The updated references and discussions presented here are meant to lead the reader to a more detailed investigation of the literature for future research in genetic analyses in marine mammal medicine and husbandry.
Acknowledgments The authors thank Andy Dizon for reviewing this chapter and making recommendations that helped add depth and additional references for the reader’s use.
References Allen, P.J., Amos, W., Pomeroy, P.P., and Twiss, S.D., 1995, Microsatellite variation in grey seals (Halichoerus grypus) shows evidence of genetic differentiation between two British breeding colonies, Mol. Ecol., 4: 653–662. Ambs, S.M., Boness, D., Bowen, W.D., Perry, E.A., and Fleischer, R.C., 1999, Proximate factors associated with high levels of extraconsort fertilization in polygynous grey seals, Anim. Behav., 58: 527–535. Amos, W., Schlötterer, C., and Tautz, D., 1993, Social structure of pilot whales revealed by analytical DNA typing, Science, 260: 670–672.
0839_frame_C14.fm Page 279 Tuesday, May 22, 2001 11:11 AM
Genetic Analyses
279
Amos, W., Pomeroy, P.P., Twiss, S.D., and Anderson, S.S., 1995, Evidence for mate fidelity in the gray seal, Science, 268: 1897–1899. Asper, E.D., Duffield, D.A., Dimeo-Ediger, N., and Shell, D., 1990, Marine mammals in zoos, aquaria and marine zoological parks in North America: 1990 census report, Int. Zoo Yearb., 29: 179–187. Baird, R.W., Willis, P.M., Guenther, T.J., Wilson, P.J., and White, B.N., 1998, An intergeneric hybrid in the family Phocoenidae, Can. J. Zool., 76: 198–204. Baker, C.S., Palumbi, S.R., Lambertsen, R.H., Weinrich, M.T., Calambokidis, J., and O’Brien, S.J., 1990, Influence of seasonal migration on geographic distribution of mitochondrial DNA haplotypes in humpback whales, Nature, 344: 238–240. Baker, C.S., Cipriano, F., and Palumbi, S.R., 1996, Molecular genetic identification of whale and dolphin products from commercial markets in Korea and Japan, Mol. Ecol., 5: 671–685. Ballou, J.D., and Foose, T.J., 1996, Demographic and genetic management of captive populations, in Wild Mammals in Captivity, Principles and Techniques, Kleiman, D.G., Allen, M.E., Thompson, K.V., and Lumpkin, S. (Eds.), University of Chicago Press, Chicago, 263–283. 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. Bruford, M.W., and Wayne, R.K., 1993, Microsatellites and their application to population genetic studies, Curr. Opin. Genet. Dev., 3: 939–943. Bruford, M.W., Cheesman, D.J., Coote, T., Green, H.A.A., Haines, S.A., O’Ryan, C., and Williams, T.R., 1996, Microsatellites and their application to conservation genetics, in Molecular Genetic Approaches in Conservation, Smith, T.B., and Wayne, R.K. (Eds.), Oxford University Press, New York, 278–297. Buchanan, F.C., Friesen, M.K., Littlejohn, R.P., and Clayton, J.W., 1996, Microsatellites from the beluga whale Delphinapterus leucas, Mol. Ecol., 5, 571–575. Buchanan, F.C., Maiers, L.D., Thue, T.D., De March, B.G.E., and Stewart, C.E.A., 1998, Microsatellites from the Atlantic walrus Odobenus rosmarus rosmarus, Mol. Ecol., 7: 1083–1085. Burg, T.M., Trites, A.W., and Smith, M.J., 1999, Mitochondrial and microsatellite DNA analyses of harbour seal population structure in the northeastern Pacific Ocean, Can. J. Zool., 77: 930–943. Coltman, D.W., Bowen, W.D., and Wright, J.M., 1998, Birth weight and neonatal survival of harbour seal pups are positively correlated with genetic variation measured by microsatellites, Proc. R. Soc. London B, 265: 803–809. Coltman, D.W., Pilkington, J.G., Smith, J.A., and Pemberton, J.M., 1999, Parasite mediated selection against inbred Soay sheep in a free-living island population, Evolution, 53: 1259–1267. Coulson, T.N., Pemberton, J.M., Albon, S.D., Beaumont, M., Marshall, T.C., Slate, J., Guiness, F.E., and Clutton-Brock, T.H., 1998, Microsatellites reveal heterosis in red deer, Proc. R. Soc. London B, 265: 489–495. Cronin, M.A., Bodkin, J., Ballachey, B., Estes, J., and Patton, J.C., 1996, Mitochondrial DNA variation among subspecies and populations of sea otters (Enhydra lutris), J. Mammal., 77: 546–557. Davis, L.G., Dibner, M.D., and Battey, J.F., 1986, Basic Methods in Molecular Biology, Elsevier, New York, 388 pp. Dizon, A.E., Chivers, S.J., and Perrin, W.F., 1997, Molecular Genetics of Marine Mammals, Special Publication 3, Society for Marine Mammalogy, Lawrence, KS, 388 pp. Duffield, D.A., and Chamberlin-Lea, J., 1990, Use of chromosome heteromorphisms and hemoglobins in studies of bottlenose dolphin populations and paternities, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 609–622. Duffield, D.A., and Wells, R.S., 1991, The combined application of chromosome, protein and molecular data in the determination of social unit structure and dynamics in Tursiops truncatus, in Genetic Ecology of Whales and Dolphins, Hoelzel, A.R. (Ed.), Report of the International Whaling Commission, Special Issue 13, Cambridge, U.K., 155–169.
0839_frame_C14.fm Page 280 Tuesday, May 22, 2001 11:11 AM
280
CRC Handbook of Marine Mammal Medicine
Duffield, D.A., Chamberlin-Lea, J., Sweeney, J., Odell, D.K., Asper, E.D., and Gilmartin, W.G., 1991, Use of corneal cell culture for R-band chromosome studies on stranded cetaceans, in Marine Mammal Strandings, Reynolds, J.E., and Odell, D.K. (Eds.), NOAA Technical Report, NMFS 88: 91–100. Escorza, S., Lux, C.A., and Costa, A.S., 1997, Methods of DNA extraction: From initial tissue preservation to purified DNA storage, in Molecular Genetics of Marine Mammals, Dizon, A.E., Chivers, S.J., and Perrin, W.F. (Eds.), Special Publication 3, Society for Marine Mammalogy, Lawrence, KS, 87–106. Gemmell, N.J., Allen, P.J., Goodman, S.J., and Reed, J.Z., 1997, Interspecific microsatellite markers for the study of pinniped populations, Mol. Ecol., 6: 661–666. Goldstein, D.B., and Schlötterer, C. (Eds.), 1999, Microsatellites: Evolution and Applications, Oxford University Press, New York, 352 pp. Goodman, S.J., 1998, Patterns of extensive genetic differentiation and variation among European harbor seals (Phoca vitulina vitulina) revealed using microsatellite DNA polymorphisms, Mol. Biol. Evol., 15: 104–118. Hagelberg, E., 1994, Ancient DNA studies, Evol. Anthropol., 2: 199–207. Hagelberg, E., Thomas, M.G., Cook, C.E., Jr., Sher, A.V., Baryshnikov, G.F., and Lister, A.M., 1994, DNA from ancient mammoth bones, Nature, 370: 333–334. Hillis, D.M., Moritz, C., and Mable, B.K. (Eds.), 1996, Molecular Systematics, 2nd ed., Sinauer Association, Sunderland, MA, 655 pp. Hoelzel, A.R., and Dover, G.A., 1991, Genetic differentiation between sympatric killer whale populations, Heredity, 66: 191–195. Jeffreys, A.J., Allen, M.J., Hagelberg, E., and Sonnberg, A., 1992, Identification of the skeletal remains of Josef Mengele by DNA analysis, Forens. Sci. Int., 56: 65–76. Jeffreys, A.J., Wilson, V., and Thein, S.L., 1985a, Hypervariable “minisatellite” regions in human DNA, Nature, 314: 67–73. Jeffreys, A.J., Brookfield, J.F.Y., and Semeonoff, R., 1985b, Positive identification of an immigration testcase using DNA fingerprints, Nature, 317: 818–819. King, J.E., 1983, Seals of the World, British Museum (Natural History), London, 154 pp. Kleiman, D.G., Allen, M.E., Thompson, K.V., Lumpkin, S., and Harris, H. (Eds.), 1996, Wild Mammals in Captivity: Principles and Techniques, University of Chicago Press, Chicago, 639 pp. Lacy, R.C., Petric, A., and Warneke, M., 1993, Inbreeding and outbreeding in captive populations of wild animal species, in The Natural History of Inbreeding and Outbreeding, Thornhill, N.W. (Ed.), University of Chicago Press, Chicago, 352–374. Litt, M., and Luty, J.A., 1989, A hypervariable microsatellite revealed by in vitro amplification of a dinucleotide repeat within the cardiac muscle actin gene, Am. J. Hum. Genet., 44: 397–401. Lyrholm, T., and Gyllensten, U., 1998, Global matrilineal population structure in sperm whales as indicated by mitochondrial DNA sequences, Proc. R. Soc. London B, 265: 1679–1684. Moritz, C., 1994, Applications of mitochondrial DNA analysis in conservation: A critical review, Mol. Ecol., 3: 401–411. O’Corry-Crowe, G.M., Suydam, R.S., Rosenberg, A., Frost, K.J., and Dizon, A.E., 1997, Phylogeography, population structure and dispersal patterns of the beluga whale Delphinapterus leucas in the western Nearctic revealed by mitochondrial DNA, Mol. Ecol., 6: 955–970. Paetkau, D., Calvert, W., Stirling, I., and Strobeck, C., 1995, Microsatellite analysis of population structure in Canadian polar bears, Mol. Ecol., 4: 347–354. Palsbøll, P.J., Clapham, P.J., Mattila, D.K., Larsen, F., Sears, R., Siegismund, H.R., Sigurjónsson, J., Vasquez, O., and Arctander, P., 1995, Distribution of mtDNA haplotypes in North Atlantic humpback whales: The influence of behaviour on population structure, Mar. Ecol. Prog. Ser., 116: 1–10. Palsbøll, P.J., Allen, J., Bérubé, M., Clapham, P.J., Feddersen, T.P., Hammond, P.S., Hudson, R.R., Jørgensen, H., Katona, S., Larsen, F., Lien, J., Mattila, D.K., Sigurjónsson, J., Sears, R., Smith, T., Sponer, R., Stevick, P., and Øien, N., 1997, Genetic tagging of humpback whales, Nature, 388: 767–769.
0839_frame_C14.fm Page 281 Tuesday, May 22, 2001 11:11 AM
Genetic Analyses
281
Palumbi, S.R., and Baker, C.S., 1994, Contrasting population structure from nuclear intron sequences and mtDNA of humpback whales, Mol. Biol. Evol., 11: 426–435. Ralls, K., Ballou, J.D., and Templeton, A.R., 1988, Estimates of lethal equivalents and the cost of inbreeding in mammals, Conserv. Biol., 2: 185–193. Reed, J.Z., Tollit, D.J., Thompson, P.M., and Amos, W., 1997, Molecular scatology: The use of molecular genetic analysis to assign species, sex and individual identity to seal faeces, Mol. Ecol., 6: 225–234. Reyes, J.C., 1996, A possible case of hybridism in wild dolphins, Mar. Mammal Sci., 12: 301–307. Ryder, O.A., and Fleischer, R.C., 1996, Genetic research and its application to zoos, in Wild Mammals in Captivity, Kleiman, D.G., Allen, M.E., Thompson, K.V., and Lumpkin S. (Eds.), University of Chicago Press, Chicago, 255–262. Shinohara, M., Domingo Roura, X., and Takenaka, D., 1997, Microsatellites in the bottlenose dolphin Tursiops truncatus, Mol. Ecol., 6: 695–696. Swinton, J., Harwood, J., Grenfell, B.T., and Gilligan, C.A., 1998, Persistence thresholds for phocine distemper virus infection in harbour seal Phoca vitulina metapopulations, J. Anim. Ecol., 67: 54–68. Syvestre, J.P., and Tanaka, S., 1985, On the intergeneric hybrids in cetaceans, Aquat. Mammals, 11: 101–108. Taberlet, P., Camarra, J.-J., Griffin, S., Uhrès, E., Hanotte, O., Waits, L.P., Dubois Paganon, C., Burke, T.A., and Bouvet, J., 1997, Noninvasive tracking of the endangered Pyrenean brown bear population, Mol. Ecol., 6: 869–876. Tautz, D., 1989, Hypervariability of simple sequences as a general source of polymorphic DNA markers, Nucl. Acids Res., 17: 6462–6471. Valsecchi, E., and Amos, W., 1996, Microsatellite markers for the study of cetacean populations, Mol. Ecol., 5: 151–156. Worthington Wilmer, J., Allen, P.J., Pomeroy, P.P., Twiss, S.D., and Amos, W., 1999, Where have all the fathers gone? An extensive microsatellite analysis of paternity in the grey seal (Halichoerus grypus), Mol. Ecol., 8: 1417–1430.
0839_frame_C14.fm Page 282 Tuesday, May 22, 2001 11:11 AM
0839_frame_C15 Page 283 Tuesday, May 22, 2001 11:11 AM
III Infectious Diseases of Marine Mammals
0839_frame_C15 Page 284 Tuesday, May 22, 2001 11:11 AM
0839_frame_C15 Page 285 Tuesday, May 22, 2001 11:11 AM
15 Viral Diseases Suzanne Kennedy-Stoskopf
Introduction The last decade of the 20th century saw an explosion in the recognition and characterization of viruses in marine mammals. In part, this increase can be attributed to heightened public concern with repeated morbillivirus epizootics in cetaceans and pinnipeds throughout the waters of the world. Success in studying marine mammals viruses has been dependent on laboratories maintaining primary pinniped and cetacean cell cultures for initial isolation of marine mammal viruses. Advances in biotechnology now allow amplification of viral genomes from tissues and detection of viral antigens with cross-reactive antibodies developed for use in better-characterized human and animal viruses. Undoubtedly, the more one looks, the more viruses will be found in marine mammals. For comprehensive reviews of marine mammal morbilliviruses, see Kennedy (1998) and Duignan (1999). For a comprehensive review of cetacean viruses, see Van Bressem et al. (1999b).
Virus Isolation—An Overview Large numbers of animals dying acutely, repeated seasonal mortality events, and recurring clinical signs in individuals suggest a virus etiology. To diagnose a viral infection requires proper handling and preservation of tissue samples. Although virus isolation is preferred, it may not always be possible. Amplification of viral gene products from tissue DNA samples by polymerase chain reaction (PCR), detection of viral antigen by immunocytochemistry, and demonstration of viral particles by electron microscopy can be used to confirm the presence of a virus. Whether the virus actually caused disease is subject to interpretation and is best resolved by recreating the clinical and pathological signs through experimental inoculation. However, this is not always practical and sometimes not even conclusive, since other factors, not duplicated in a laboratory setting, can affect the pathogenicity of a virus. Samples collected for potential virus isolation, such as skin lesions and focal areas of necrosis in organs, may be obvious. Take samples as aseptically as possible and be sure to include normal tissue with the abnormal tissue. In the absence of overt pathology, take representative samples from lung, liver, kidney, spleen, lymph nodes, and brain. Individual specimens should be frozen in plastic bags, preferably sterile, at −70°C (−94°F). Admittedly, this is not always feasible in field settings, unless one is well prepared. A container of ice can keep samples cool for 6 to 12 hours 3 until they can be transferred to an ultralow freezer. Samples should be small enough (<1 cm ) that they freeze quickly. Multiple small samples are preferable to one large chunk of tissue.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
285
0839_frame_C15 Page 286 Tuesday, May 22, 2001 11:11 AM
286
CRC Handbook of Marine Mammal Medicine
Samples frozen in this manner are suitable for primary virus isolation and identification of viral nucleotide sequences by PCR. Sometimes it is better to attempt virus isolation from fresh tissue. This requires coordination with a laboratory that can provide transport media for the samples. Tissue culture medium and balanced salt solutions, both fortified with antibiotics and antifungal agents, with or without fetal bovine serum, have been used to maintain specimens for as long as 24 hours on ice. In the absence of tissue culture medium, sterile physiological saline may be used. Investigators have developed primary cell cultures from a variety of tissues of different marine mammals that have greatly facilitated the isolation of viruses from these species. If virus cannot be isolated, then specimens preserved for electron microscopy can be examined for viral particles. Although ultrastructural studies can be performed on formalin-fixed tissues, morphology is usually better if a fixative such as gluteraldehyde is used. Again, the limiting factor is ice, since most fixatives for electron microscopy must be refrigerated. If the desired fixative is not available in the field, then tissue samples stored in plastic bags or wrapped in aluminum foil can be stored in ice for 4 to 6 hours before transferring to 3 to 5% buffered 3 gluteraldehyde. Samples should be no larger that 1 mm to allow adequate fixation with gluteraldehyde. If samples cannot be kept cold or transferred quickly enough, then fix them in 10% buffered formalin. Formalin-fixed tissue, in some instances, may be suitable for immunocytochemistry. However, formalin can adversely affect the viral proteins present in tissues so that the virus-specific antibodies used for immunocytochemical detection do not recognize their target. Snap freezing of tissue can preserve both the tissue morphology as well as the viral proteins, but requires access to freezing compounds, and either dry ice or liquid nitrogen, which makes it difficult to perform in field settings. Finally, collect serum samples to archive in a non-defrosting freezer at −20°C (−4°F). In the event of an animal’s recovery, collect convalescent serum samples. Serological assays to detect virus-specific antibodies indicate exposure (see Chapter 12, Immunology). With paired serum samples, a significant rise in antibody titer, usually a fourfold or greater increase, suggests a recent infection. Paired serum samples are rarely available from free-ranging animals or those found dead. However, a positive IgM titer in single serum samples also indicates recent infection by agents such as morbilliviruses.
Poxviruses Host Range Poxvirus is associated with skin lesions in Atlantic bottlenose dolphins (Tursiops truncatus), a stranded Atlantic white-sided dolphin (Lagenorynchus acutus), a killer whale (Orcina orca), dusky dolphins (Lagenorynchus obscurus), long-beaked common dolphins (Delphinus capensis), Hector’s dolphin (Cephalorhynchus hectori), and Burmeister’s porpoises (Phocoena spinipinnis) (Geraci et al., 1979; Flom and Houk, 1979; Van Bressem et al., 1993; Van Bressem and Van Waerebeek, 1996; Duignan, 2000). Affected populations have been reported in the North Atlantic, North Sea, and the southeast Pacific, as well as individuals in captivity. Poxvirus has been identified morphologically in skin lesions from numerous species of pinnipeds, including California sea lions (Zalophus californianus) (Wilson et al., 1969; 1972a,b), South American sea lions (Otaria byronia) (Wilson and Poglayen-Neuwall, 1971), harbor seals (Phoca vitulina) (Wilson et al., 1972a), gray seals (Halichoerus grypus) (Hicks and Worthy, 1987), and a northern fur seal (Callorhinus ursinus) (Hadlow et al., 1980). Both captive and free-ranging pinnipeds are affected.
0839_frame_C15 Page 287 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
287
Clinical Signs Cutaneous manifestations of associated poxvirus infections in cetaceans range from ring or pinhole lesions to black, punctiform, stippled patterns. The latter are referred to as “tattoo” lesions. Ring or pinhole lesions appear as solitary, 0.5 to 3 cm, round or elliptical blemishes, which sometimes coalesce. These lesions are usually light gray in color and may have a dark gray border, although the reverse color pattern is also seen. Geraci et al. (1979) suggested that these blemishes progress to tattoo lesions. Smith et al. (1983a) reported the development of raised, blanched, edematous lesions in connection with previously existing tattoo lesions. Poxvirus was demonstrated in both sets of lesions. Lesions may occur on any part of a cetacean’s body, but are most frequently observed on the skin of the dorsal body surface, particularly the head, and on the pectoral flippers, dorsal fin, and tail fluke. Lesions may persist for months or years without any apparent ill effect to the animal. The distribution of cutaneous lesions in pinnipeds is similar, and includes the head, neck, and flippers. In harbor seals and gray seals, initial lesions begin as small, raised nodules, 0.5 to 1 cm in diameter which, over a period of approximately 1 week, may increase to 1.5 to 3 cm in diameter. During the second week, these lesions ulcerate and, in the case of harbor seals, may suppurate. Also during the second week, rapidly spreading satellite lesions develop in harbor seals around the initial nodules. After the fourth week, lesions begin to regress in both species of seal, although nodules have persisted as long as 15 to 18 weeks in harbor seals. Areas of alopecia and scar tissue may remain following resolution. In the northern fur seal and South American sea lions, nodules are nonulcerative, whereas in California sea lions, nodules are both ulcerative and nonulcerative. The case involving two South American sea lions was unusual in that the nodules were irregularly distributed over the entire body surface.
Therapy Poxviruses of marine mammals do not appear to cause systemic infections. Although animals have died with cutaneous poxvirus lesions, other factors were responsible for their deaths. Therapy to control secondary bacterial infections is indicated only when skin suppurates. Otherwise, lesions usually resolve, given time.
Pathology In cetaceans, cells of the stratum intermedium contain small, spherical or irregularly shaped, pale eosinophilic, intracytoplasmic inclusion bodies. Cells with inclusion bodies are in a so-called transition zone and mark the edges of the ring lesion. Stratum intermedium cells in the central zone of the lesion show marked cytoplasmic vacuolation and a prominent reticular pattern of keratinaceous fibers. The longitudinal axis of the uninfected tissue adjacent to the lesion is displaced from its normal orientation to one perpendicular to the skin surface. The outermost layer of the epidermis in cetaceans, known as the stratum externum or parakeratotic layer, is not keratinized as in most mammals, but consists of 10 to 15 layers of viable flattened cells with elongated nuclei and mitochondria. These cells, which have an extraordinarily high mitotic rate, continually slough, leaving a smooth epidermal surface. In poxvirus infections, the number of cells in the stratum externum increases up to threefold, compressing the cells downward so there is no elevation of the skin surface associated with the lesion. As the lesions progress to form the dark stippled patterns associated with tattooing, breaks appear in the stratum externum, creating pits that extend to the stratum intermedium. Histological features of formalin-fixed biopsy specimens from harbor seals, gray seals, and California sea lions are similar. Variable amounts of hyperkeratosis and parakeratosis are present
0839_frame_C15 Page 288 Tuesday, May 22, 2001 11:11 AM
288
CRC Handbook of Marine Mammal Medicine
in the stratum corneum. In some areas, ulceration, neutrophilic infiltration, and bacteria are observed. Cytoplasmic vacuolation and nuclear degeneration are most prominent in cells of the stratum spinosum. Large eosinophilic intracytoplasmic inclusion bodies (2 to 15 µm) are found in these cells of the epidermis. In the dermis, a mixed inflammatory cell infiltrate is present. In the northern fur seal and South American sea lion, nodules are composed of compact lobules of polygonal epithelial cells, which proliferate downward toward the dermis. These cells have abundant, finely granular, eosinophilic cytoplasm and a round vesicular nucleus containing a prominent nucleolus. A single, large, round eosinophilic intracytoplasmic inclusion body, 8 to 26 µm in diameter, may be located in these cells.
Diagnosis A presumptive diagnosis based on the presence of eosinophilic, intracytoplasmic inclusion bodies histologically is confirmed by the identification of typical poxvirus particles in skin biopsies prepared as ultrathin sections for electron microscopy. Two genera of poxviruses have been reported in marine mammals, Parapoxvirus and Orthopoxvirus. Morphologically, viruses in the genus Parapoxvirus are distinct from the other genera. Virions in this genus appear more ovoid, show a crossed helical surface pattern in negatively stained preparations, and average about 150 × 200 nm in size (Palmer and Martin, 1988). Based on these distinguishable characteristics, poxviruses identified in California sea lions, harbor seals, and gray seals are assigned to the genus Parapoxvirus. The poxviruses responsible for reported lesions from South American sea lions and the northern fur seal may belong to genera other than Parapoxvirus, because the target cell for virus replication in vivo appears to be epithelial rather than fibroblastic. Unfortunately, there were no negatively stained preparations made of these isolates so it is impossible to determine if they possess the characteristic morphology of parapoxviruses. Cetacean poxviruses are brick-shaped, have a surface pattern resembling whorled filaments, and average 225 × 300 nm in size. This morphology is consistent for Orthopoxvirus, as well as for other more species-restricted poxvirus genera (Palmer and Martin, 1988). Virus isolation and further characterization will be necessary to confirm the genus of cetacean poxviruses. Although poxvirus morphology is diagnostic, viral isolation or determination of appropriate nucleotide sequences is necessary to compare isolates from different species of pinnipeds or between isolates from pinnipeds and cetaceans. Poxvirus isolation from marine mammals is difficult and has not succeeded without the use of primary cell cultures from target species. Parapoxvirus has been isolated from gray seals using either gray seal or harbor seal kidney cells (Osterhaus et al., 1994; Nettleton et al., 1995). After initial isolation in gray seal kidney cells, that particular isolate replicates in fetal lamb muscle, skin, and cornea and is antigenically related to orf, a sheep parapoxvirus, based on monoclonal antibody reactivity (Housawi et al., 1998). An Orthopoxvirus has been isolated from a gray seal with lesions that contained viral particles morphologically consistent for both Parapoxvirus and Orthopoxvirus. (Osterhaus et al., 1990). Only viral particles consistent with orthopoxviruses were recovered from primary gray seal skin explants inoculated with tissue suspensions of the poxlike lesions. This isolate could be passaged with cell-free culture medium, and was recognized by antibodies against mousepox virus in the genus Orthopoxvirus. The significance of the one Orthopoxvirus isolation from a gray seal remains to be determined.
Differentials Cutaneous streptothricosis causes nodular lesions in pinnipeds, similar to those caused by poxvirus (Wilson et al., 1972b). The causative agent, Dermatophilus congolensis, can be isolated on blood agar, provided there is no overgrowth with other bacteria. Alternatively, numerous
0839_frame_C15 Page 289 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
289
branching filaments with longitudinal and transverse divisions can be detected in histological sections of lesions. Caliciviruses cause vesicular lesions in pinnipeds that may be confused with eroding poxvirus lesions. However, vesicular lesions due to caliciviruses usually involve only the flippers, not the head and neck of the animal, although oral lesions do occur.
Epidemiology Van Bressem and Van Waerebeek (1996) note that the prevalence of poxvirus infection correlates with the body length class of dusky dolphins, long-beaked common dolphins, and Burmeister’s porpoises taken from the eastern South Pacific. Prevalence of tattoo lesions peaks around weaning, presumably due to loss of maternal antibodies, and gradually decreases in the older delphinids. Prevalence, however, remains high in the adult Burmeister’s porpoise and is greater in males than in females. Regression of lesions probably correlates with immunity. Formation of detectable virus-specific antibody coincides with resolution of both acute and chronic skin lesions in bottlenose dolphins (Smith et al., 1983a). Lesions also resolve locally in areas from which biopsies are taken. Geraci et al. (1979) speculates that the appearance and regression of pox-associated lesions in dolphins is related to their general health and environmental conditions. Poor water quality, drastic drops in water temperature, and the development of gastric ulcers have all coincided with the onset of poxvirus lesions. Parapoxvirus lesions in pinnipeds are frequently seen in neonates. Parapoxviruses are readily transmissible in such susceptible populations. Outbreaks typically occur in postweanling pinnipeds recently introduced into captivity. The incubation period ranges from 3 to 5 weeks. In the case of gray seals, only animals with access to cement pools developed lesions, suggesting that a breach in the epithelial surface is required to start an infection. Osterhaus et al. (1994) have suggested that concurrent infection with phocine distemper virus can predispose animals to outbreaks. Parapoxvirus lesions can recur in ruminants, so the possibility of relapses exists for pinnipeds.
Public Health Significance The parapoxviruses of pinnipeds can cause isolated lesions on the hands of humans that come in contact with infected animals. The assumption should be made that an animal can shed virus as long as lesions are present. In one report, two of three people handling infected gray seals developed solitary skin lesions 19 days after initial contact (Hicks and Worthy, 1987). Between the time the lesion was first noted and the scab fell off, 35 days elapsed. In one case, the lesion completely healed over a period of 3 to 4 months. However, the other person had repeated relapses, and it took nearly a year for the lesion to heal completely. Although not life-threatening, pinniped parapoxviruses are clearly a potential nuisance to humans. People who come in contact with infected animals should wear gloves. Most poxviruses are relatively resistant to such environmental factors as drying and cold temperatures, as well as most common disinfectants.
Papillomaviruses Host Range Viral particles consistent with Papillomavirus, a genus in the family Papovaviridae, have been identified in sperm whale (Physeter macrocephalus) genital papillomas (Lambertson et al., 1987), harbor porpoise (Phocoena phocoena) cutaneous papillomas (Geraci et al., 1987; Van Bressem et al., 1999a), beluga (Delphinapterus leucas) gastric papillomas (De Guise et al., 1994),
0839_frame_C15 Page 290 Tuesday, May 22, 2001 11:11 AM
290
CRC Handbook of Marine Mammal Medicine
killer whale cutaneous papillomas (Bossart et al., 1996), and genital warts from dusky dolphins and Burmeister’s porpoises (Van Bressem et al., 1996). Lesions morphologically consistent with papillomas and fibropapillomas have been described in Atlantic white-sided dolphins, common dolphins, bottlenose dolphins, narwhales (Monodon monoceros), and blue whales (Balaenoptera musculus) (Geraci et al., 1987; Van Bressem et al., 1996).
Clinical Signs Areas of localized epithelial hyperplasia with a defined boundary and an intact basement membrane define papillomas and warts. These growths, sometimes referred to as benign tumors, have been seen on the skin, penile mucosa, vaginal mucosa, gastric mucosa, and tongue of both free-ranging and captive odontocetes and on the tongue of a free-ranging mysticete. In general, lesions are focal and randomly distributed. There is one report of bilaterally symmetrical lesions occurring in the axillae and over the peduncle of a killer whale (Bossart et al., 1996). The cutaneous lesions appear as raised, often smooth, plaques that are the color of the underlying skin. The mucosal lesions vary in color and often have an irregular surface. Size is quite variable, ranging from a few millimeters to in excess of 20 cm (8 in.).
Therapy No therapy is available. Lesions are generally self-limiting and will regress. In most domestic animal species, complete immunity follows regression of lesions, but not enough is known to make this statement about cetacean papillomas. In one instance, a killer whale experienced a 10-year cyclical pattern of proliferation and regression (Bossart et al., 1996). This animal had an inverted CD4 : CD8 lymphocyte ratio and abnormal cytokine expression, suggesting that immunosuppression may have been responsible for relapses.
Pathology Characteristic histological lesions include marked epidermal hyperplasia and hydropic degeneration or koilocytosis (the hollow appearance of a cell due to large perinuclear vacuoles). The presence of lymphocytic infiltrates varies, and intranuclear inclusions are not necessarily seen. Van Bressem et al. (1996) make the distinction between warts and papillomas, with the latter characterized by hyperplasia of the papillae.
Diagnosis Papillomaviruses do not usually grow in tissue cultures and cannot always be detected microscopically. When detected by electron microscopy, papilloma viral particles are naked icosahedrons about 55 nm in diameter. In the absence of viral particles, papillomavirus group–specific antigens were detected immunocytochemically in formalin-fixed warts from dusky dolphins and Burmeister’s porpoises (Van Bressem et al., 1996).
Differentials A plaquelike thickening on the surface of the penis from a stranded dead harbor porpoise was characterized by epithelial hyperplasia and numerous amphophilic to eosinophilic intranuclear inclusion bodies (Lipscomb et al., 1996). The inclusions contained viral particles consistent with herpesvirus when examined by electron microscopy and were recognized immunocytochemically by antibodies to human herpesvirus-1 and -2.
0839_frame_C15 Page 291 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
291
Epidemiology The genital location of warts and their high prevalence in Burmeister’s porpoises and dusky dolphins off the central coast of Peru suggest venereal transmission (Van Bressem et al., 1996). Lesions were sufficiently severe in two Burmeister’s porpoises possibly to impede copulation, prompting the observation that this viral infection could have an indirect impact on population dynamics. Papillomas in the first gastric compartment of 8 of 24 belugas from the St. Lawrence River necropsied between 1988 and 1990 suggest an epizootic during this period, as the lesions have not been seen as frequently since (De Guise et al., 1994). The transmission mode for cutaneous papillomas and warts is not known. Atlantic bottlenose dolphins housed with an affected killer whale did not develop lesions (Bossart et al., 1996), which is consistent with the species specificity of papillomaviruses. However, harbor porpoises housed with an affected harbor porpoise also did not develop lesions, suggesting the unaffected animals were either immune or experienced inapparant infections, or that the virus was not highly contagious (Van Bressem et al., 1999a).
Public Health Significance Papillomaviruses are very species specific and should pose no human health risk.
Adenoviruses Host Range Adenoviruses have been isolated from rectal swabs of a sei whale (Balaenoptera borealis) taken in the Antarctic (Smith and Skilling, 1979), from colon samples of two bowhead whales (Balaena mysticetus) collected in Barrow, Alaska (Smith et al., 1987), and from the intestines of a beluga from the St. Lawrence estuary (De Guise et al., 1995). The pathogenicity of these viruses and their relationship to each other are not known. Viral particles characteristic of adenoviruses have been observed in livers from California sea lions with hepatitis (Britt et al., 1979; Dierauf et al., 1981).
Clinical Signs There are two published case reports of hepatitis associated with adenovirus in California sea lions, in which a total of six animals were affected. All were stranded along the coast of California in the spring, and all but one were estimated to be less than a year old. Animals died between 1 and 28 days following rescue. Early clinical signs in the animal that died at 28 days included weakness, emaciation, and photophobia (Dierauf et al., 1981). Polydypsia was also noted. Gastrointestinal signs included abdominal splinting and intermittent blood-tinged diarrhea that gradually worsened despite supportive therapy. Hematology showed a relative lymphopenia and monocytosis. These clinical signs and hemograms are consistent with those observed in dogs with infectious canine hepatitis (ICH), a disease caused by canine adenovirus type 1 (CAV-1).
Therapy To date, adenovirus-associated hepatitis in California sea lions is presumably fatal. However, the sample size is small, and undiagnosed animals with milder clinical signs may have recovered. As with all viral infections, therapy is supportive. Vaccination with a killed CAV-1 vaccine protected American black bears during an epizootic of adenovirus infection (Collins et al., 1984),
0839_frame_C15 Page 292 Tuesday, May 22, 2001 11:11 AM
292
CRC Handbook of Marine Mammal Medicine
and perhaps should be considered if a similar outbreak is recognized in sea lions (see Diagnosis, below).
Pathology Livers ranged from pale yellow to congested and from normal size to greatly enlarged. The most prominent histopathological finding in all cases was hepatic necrosis. Lymphoid cells were in the walls of hepatic arteries and central veins. In the first report, basophilic intranuclear inclusions were observed in hepatocytes (Britt et al., 1979). In the second report, granular amphophilic intranuclear inclusions were observed in Kupffer’s cells (Dierauf et al., 1981). Pneumonia was present in all animals. Although adenoviruses are pneumotropic, no inclusion bodies were reported in sections of lung, and the pneumonias were attributed to other causes, such as parasitism and aspiration.
Diagnosis Electron microscopy was performed with formalin-fixed tissues. In all cases, the diagnosis was based on the detection of viral particles with typical adenovirus morphology in the nuclei of cells with inclusions. A few particles were found in the cytoplasm near nuclear pores. Virions were hexagonal to round and had diameters ranging from 70 to 75 nm. Particles were not enveloped. Attempts to isolate the sea lion adenovirus from liver homogenates failed using the following cells: green monkey kidney, rhesus monkey kidney, WI36 fibroblasts, HeLa, and human amnion. CAV-1 replicates in Madin Darby canine kidney (MDCK) cells, porcine kidney and testicle cells, and raccoon and ferret kidney cells (Pursell et al., 1983; Collins et al., 1984). Future attempts to isolate adenovirus from sea lions should use one or more of these cells if primary sea lion kidney cells are unavailable.
Epidemiology The source of adenovirus in California sea lions is not known. Insufficient information is available to determine the relationship of the sea lion adenovirus to CAV-1. However, considering CAV-1 is infectious to black bears and otariid seals are phylogenetically related to ursids, it is tempting to speculate that canines are the source of infection. All adenovirus-infected sea lions were brought to marine mammal centers where numerous other pinnipeds were housed, yet there was no clinical evidence that any of these other animals became infected.
Public Health Significance Adenovirus from California sea lions and CAV-1 are not known to cause any human disease.
Herpesviruses Host Range Herpesviruses and herpeslike viruses have been detected in belugas (Martineau et al., 1988; Barr et al., 1989), harbor porpoises (Kennedy et al., 1992; Lipscomb et al., 1996), and dusky dolphins (Van Bressem et al., 1994). Herpesviruses have been isolated from harbor seals (Osterhaus et al., 1985; Harder et al., 1996; King et al., 1998), gray seals (Kennedy-Stoskopf, 1988; Lebich et al., 1994), and a California sea lion (Kennedy-Stoskopf et al., 1986). Serological surveys indicate that herpesvirus infections occur in a variety of pinnipeds including Weddell
0839_frame_C15 Page 293 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
293
seals (Leptonychotes weddellii) from the Antarctic (Stenvers et al., 1992); harp (Pagophilus groenlandicus) and hooded (Cystophora cristata) seals from the Arctic (Daoust et al., 1994; Stuen et al., 1994); and walruses (Odobenus rosmarus), northern fur seals, harbor seals, spotted seals (Phoca largha), ribbon seals (Histriophoca fasciata), Steller sea lions (Eumetopias jubatus), bearded seals (Erignathus barbatus), and ringed seals (Phoca hispida) from waters between Alaska and Russia (Zarnke et al., 1997).
Virology Two distinct herpesviruses have been characterized from harbor seals and gray seals. Phocid herpesvirus type-1 (PhHV-1) has been isolated from harbor seals and gray seals in northern European waters and from harbor seals in waters from both coasts of the United States (Harder et al., 1996; King et al., 1998). Sequence analysis of PCR products shows greatest similarity to members of the genus Varicellovirus of the subfamily α-herpesvirinae and in particular to canine herpesvirus (Harder et al., 1996; Harder and Osterhaus, 1997; King et al., 1998). The Pacific harbor seal isolate has nine nucleotide substitutions over a short segment of the glycoprotein D compared with the European and western Atlantic isolates. This parallels the divergence of harbor seal subspecies and suggests evolution of the virus with its host (King et al., 1998). Phocid herpesvirus-2 has been isolated from harbor seals in the western Atlantic and the Wadden Sea (Lebich et al., 1994; Harder et al., 1996). Antibody cross-neutralization is negligible between PhHV-1 and -2. Sequence analysis of PhHV-2 PCR products is most consistent with herpesviruses in the subfamily γ-herpesvirinae. The putative γ-herpesvirus from a California sea lion was in fact an isolate from a captive gray seal that was mislabeled. The original isolate from the lung of a California sea lion (Kennedy-Stoskopf et al., 1986) has not been successfully recultured to permit further characterization. DNA sequences detected by consensus primer PCR in metastatic carcinomas from California sea lions were most consistent with a γ-herpesvirus in the genus Rhabdinovirus (Lipscomb et al., 2000b).
Clinical Signs With the exception of a herpesviral encephalitis in a harbor porpoise found dead (Kennedy et al., 1992), all reports of herpesviruses in cetaceans are restricted to skin lesions and, in one case, penile mucosa (Lipscomb et al., 1996). Esophageal ulcers have been seen in belugas and a harbor porpoise with herpesvirus infections, but no viral particles were detected in the esophageal lesions (Kennedy et al., 1992; Mikaelian et al., 1999). A generalized dermatitis was observed in two stranded, juvenile beluga females that died from multisystemic diseases (Martineau et al., 1988). Lesions were circular, measuring up to 2 cm in diameter, and appeared slightly depressed. A narrow dark rim outlined paler-than-normal skin that contained a small dark center. The centers of some lesions were necrotic. Similar lesions developed on a subadult female 3.5 months after capture (Barr et al., 1989). Over a 7-month period, a lesion in the area of the dorsal ridge grew to cover an area 20 to 30 cm in diameter. The animal was otherwise apparently healthy. The lesions became quiescent but remained visible. Lesions in the free-ranging dusky dolphins appeared as a few black dots on the beak, but in one animal were widely dispersed (Van Bressem et al., 1994). All affected cetaceans were subadults, except for the harbor porpoise with a penile lesion. Fatal generalized PhHV-1 infections of pinnipeds usually occur only in neonates (Borst et al., 1986; Gulland et al., 1997; Harder et al., 1997). During the phocid distemper virus (PDV) epizootic in 1988, generalized PhHV-1 was detected in adult seals and was concluded to be secondary to the PDV (Osterhaus and Vedder, 1988). Clinical signs differ between European and Pacific harbor seals infected with PhHV-1. In European seals, the respiratory system is affected. Early in the course of fatal infections, nasal discharge, inflammation of the oral mucosa,
0839_frame_C15 Page 294 Tuesday, May 22, 2001 11:11 AM
294
CRC Handbook of Marine Mammal Medicine
vomiting, diarrhea, and fever as high as 40°C (36.5 to 37.8°C is normal) may be observed. Coughing, anorexia, and lethargy occur in later stages. In Pacific harbor seals, the adrenal glands and the liver are the target organs. Clinical signs may vary because concurrent bacterial infections are frequently present (Gulland et al., 1997). Marked lymphopenia occurs prior to death, but death is not caused by adrenal insufficiency (Gulland et al., 1999). Serum chemistries remain within normal ranges. Phocid herpesvirus-2 has not been associated with clinical disease. However, the gray seal PhHV-2, previously confused for a California sea lion virus, was isolated from skin lesions on an adult female. The lesions were circumscribed areas of alopecia, approximately 0.5 cm in diameter, distributed primarily along the abdomen. These recurring lesions were seasonal and coincided with the male gray seal’s breeding season.
Therapy In the cases of systemic infection, therapy is supportive. In the face of a documented epizootic, oral acyclovir (9-(2) hydroxyethoxymethyl guanine), an antiviral agent used to treat herpesvirus infections in humans, may be tried, although its use has not yet been documented in marine mammals. This drug does not eliminate the infection, but significantly shortens local symptoms in primary infections. Its effectiveness during recurrent episodes and against herpesviruses other than Herpes simplex is variable. Therefore, its therapeutic value against pinniped or cetacean herpesvirus is highly speculative. Harder et al. (1997) suggests that convalescent seal sera may be useful to provide passive protection in newborn seals at risk. Similarly, considering the close antigenic relationship with herpesviruses of terrestrial carnivores (Lebich et al., 1994), inactivated or subunit vaccines derived from canid or felid herpesviruses might be useful.
Pathology In belugas, epidermal necrosis and intracellular edema characterize skin lesions. Necrotic keratinocytes are pyknotic and display variable degrees of cytoplasmic vacuolation. Eosinophilic intranuclear inclusions are prominent. The one reported case of herpesvirus encephalitis in a harbor porpoise featured neuronal degeneration and necrosis, neurophagia, and diffuse microglial infiltration limited to the cerebral cortex (Kennedy et al., 1992). Many neurons contained intranuclear acidophilic inclusions. In European harbor seals, prominent gross lesions include pneumonia, hepatomegaly, and small erosions of the oral mucosa. In some cases in Europe, not all lesions observed were conclusively linked to herpesvirus. Microscopically, the liver shows severe to massive coagulation necrosis with no specific zonal pattern. Interstitial pneumonia with mononuclear infiltrates is also found. No intranuclear inclusions are observed, unlike the presence of amphophilic intranuclear inclusions consistently seen in the adrenal tissues of Pacific harbor seals. Adrenal cortical necrosis varies in severity and is located mostly within the zona fasciculata. Mild hepatic necrosis may also be seen.
Diagnosis Herpesviruses have yet to be isolated from cetaceans. All diagnoses are based on the detection of characteristic viral particles by electron microscopy or immunocytochemical staining. Two types of herpesvirus particles are seen in both cetaceans and pinnipeds. One is a naked nucleocapsid ranging from 60 to 110 nm in diameter found in the nucleus. The other is an enveloped nucleocapsid ranging from 115 to 250 nm found in the cytoplasm. Dark nucleocapsids surrounded by
0839_frame_C15 Page 295 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
295
a clear halo and a dark concentric ring produce the so-called target configuration typical of herpesviruses. Antisera to human herpesvirus-1 and bovine herpesvirus-1, both α-herpesviruses, recognize harbor porpoise herpesviral antigens in fixed tissues (Kennedy et al., 1992; Lipscomb et al., 1996). Antiserum to the latent membrane protein of Epstein–Barr virus, a γ-herpesvirus, reacts with an intraepithelial neoplasia in a California sea lion (Lipscomb et al., 2000b). Phocid herpesvirus-1 and -2 were initially isolated from primary host tissues (lung, kidney, urinary bladder, and skin), leukocytes, or nasal swabs. Primary pinniped cell cultures were used initially but many of these isolates now replicate in Crandell feline kidney (CrFK) cells. Virus isolation may not always be feasible because of lack of appropriate pinniped cell cultures and degraded specimen quality. A PCR assay overcomes both of these drawbacks, although Harder et al. (1997) suggest that testing paired serum samples for specific antibodies is probably the method of choice when screening a population.
Differentials See morbillivirus differentials.
Epidemiology Little is known about the epidemiology of herpesvirus infections in cetaceans. Mikaelian et al. (1999) demonstrated that at least 50% of the serum samples from 13 belugas found dead from various causes along the St. Lawrence estuary between 1995 and 1997 had antibodies that crossreacted with bovine herpesvirus-1. Two calves that had not nursed had no detectable titers. Infections in cetaceans probably increase with age and become latent, similar to α-herpesvirus infections in other species. Serosurveys indicate that herpesviruses are widespread in pinnipeds. Phocid herpesvirus-1 appears most common in European seals, whereas a limited study sample suggests PhHV-2 is more common in seals in the western Atlantic (Harder et al., 1996). A closely related PhHV-1 is present in Pacific harbor seals off the coast of California (King et al., 1998), whereas a variety of pinniped species have antibodies to both PhHV-1 and -2 in the seas between Alaska and Russia (Zarnke et al., 1997). The relationship between PhHV-2, a γ-herpesvirus of phocid seals, and the putative γ-herpesvirus of California sea lions is not known. Phocid herpesvirus-1 can cause mortality in neonates similar to the effect of α-herpesviruses in other mammals. Although the pathology is different in European and Pacific harbor seals infected with PhHV-1, the available epidemiological evidence suggests that pups are probably infected shortly after birth or perhaps in utero, and lateral transmission is possible in rehabilitation settings. Based on the course of the 1984 epizootic at the Seal Nursery Station in the Netherlands, the incubation period appears to be 10 to 14 days (Borst et al., 1986). Since herpesviruses cause persistent infections in their hosts, latent virus can be reactivated so that recurrent manifestations of infection appear. Mechanisms responsible for triggering new rounds of viral replication are not understood, but such factors as stress and immunosuppression are associated with recrudescence. Whether or not PhHV-1 can cause abortions like other α-herpesviruses remains to be determined, but the ramifications on populations could be considerable. The presence of a putative γ-herpesvirus in intraepithelial neoplasia of the genital tracts of California sea lions implies sexual transmission (Lipscomb et al., 2000b).
Public Health Significance Based on the biology of better-characterized herpesviruses in animals other than primates, the herpesviruses of marine mammals appear not to be zoonotic.
0839_frame_C15 Page 296 Tuesday, May 22, 2001 11:11 AM
296
CRC Handbook of Marine Mammal Medicine
Morbilliviruses For review and comprehensive primary references, see Kennedy (1998) and Duignan (1999).
Host Range Since 1987, morbilliviruses have caused several major epizootics with high mortalities in marine mammals including European harbor seals and gray seals in 1988, Baikal seals (Phoca sibirica) in Siberia from 1987 to 1988, Caspian seals (P. caspica) in the Caspian Sea in 2000 (Kennedy et al., 2000), striped dolphins (Stenella ceoruleoalba) in the Mediterranean Sea from 1990 to 1992, Black Sea common dolphins (Delphinus delphis ponticus), and bottlenose dolphins along the eastern coast of the United States from 1987 to 1988 and in the Gulf of Mexico from 1993 to 1994. Morbillivirus-associated mortalities have also occurred in harbor porpoises, pilot whale (Globicephalus sp.) calves, a white-beaked dolphin, a harp seal, two hooded seals, and Mediterranean monk seals (Monachus monachus). Serosurveys have demonstrated morbillivirus antibodies in 14 of 18 odontocete species in the western Atlantic ranging from Arctic Canada to the Gulf of Mexico. Antibodies have also been detected in dusky dolphins, long-beaked common dolphins, and bottlenose dolphins from the Southeast Pacific (Van Bressem et al., 1998; Duignan, 2000). Pinniped species seropositive for morbilliviruses include harp seals, ringed seals, harbor seals, and gray seals in waters where they range between northwestern Europe and Arctic Canada south to the New England coastline of the United States. Antibodies have also been detected in leopard seals (Hydrurga leptonyx) and crabeater seals (Lobodon carcinophagus) in Antarctica, walrus from northwestern Canada, and New Zealand sea lions (Phocarctos hookeri) and fur seals (Arctocephalus fosteri) (Duignan, 2000). Manatees (Trichecus manatus) from Florida waters and polar bears (Ursus maritimus) in Alaska and Russia also have antibodies to morbilliviruses (Garner et al., 2000).
Virology The genus Morbillivirus is in the family Paramyxoviridae. Until 1988, there were four members in this genus: measles virus (MV), canine distemper virus (CDV), rinderpest virus (RPV), and peste-des-petits ruminants virus (PPRV). Phocine distemper virus (PDV), responsible for the 1988 epizootic in northern European harbor seals, is more closely related to CDV than other morbilliviruses, but sequence differences are such that PDV is a separate viral species. CDV, however, was responsible for the epizootic in the Baikal and Caspian seals and the antibody titers in Antarctic seals. During the 1988 PDV epizootic, harbor porpoises also stranded with distemper-like lesions. A morbillivirus was isolated from these animals and called porpoise morbillivirus (PMV). Subsequently, a morbillivirus was isolated from striped dolphins in the Mediterranean during the 1990 epizootic and called dolphin morbillivirus (DMV). Both PMV and DMV are distinct from PDV and are more closely related to the ruminant morbilliviruses RPV and PPRV. However, PMV and DMV differ from each other only at a few epitopes and are now generally considered strains of the same viral species, cetacean morbillivirus (CMV).
Clinical Signs Clinical signs are infrequently observed in free-ranging cetaceans. Body condition is poor and there is often a heavy accumulation of ectoparasites. Abnormal behavior and respiratory distress have also been reported. In pinnipeds, clinical signs resemble canine distemper in dogs and include fever, serous or mucopurulent oculonasal discharge, conjunctivitis, keratitis, coughing, difficulty breathing,
0839_frame_C15 Page 297 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
297
diarrhea, and abortion. Muscle twitching, abnormal posture, and increased tolerance to humans have been reported. Subcutaneous emphysema of the cervical and thoracic regions causes increased buoyancy and interferes with diving. Skin lesions characterized by alopecia and crusting that affected the extremities of a stranded harp seal contained PDV. Dermatitis associated with PDV was also observed in a hooded seal (Lipscomb et al., 2000a).
Therapy Treatment is supportive, but mortality is high in susceptible populations. Morbilliviruses infect lymphoid tissues, causing immunosuppression and increased susceptibility to secondary infections. Vaccinations with a commercially available modified live virus canine distemper (CD) vaccine, a killed CD vaccine, and a subunit CD vaccine have been used in Europe in rehabilitation centers. Antibody responses are elicited, and appear to be protective, although vaccination does not necessarily prevent replication of PDV in vaccinated seals and subsequent shedding to susceptible animals. Vaccination has not been performed in North America, in part because there is currently no readily available safe and efficacious vaccine other than modified live virus CD, and there has been no perceived need to vaccinate. Vaccination of freeranging marine mammals remains controversial and is difficult to implement effectively.
Pathology Bronchial pneumonia and alveolitis are the most common findings in both cetaceans and pinnipeds. Lungs are edematous with areas of emphysema and consolidation. Bronchointerstitial pneumonia with congestion, edema, serofibrinous exudation into alveoli, proliferation of type II pneumocytes, and syncytia are seen histologically. Syncytia and inclusions (intracytoplasmic and intranuclear) are more common in cetacean than pinniped lung tissue. The central nervous system, most commonly the cerebrum, may also be affected. Neuronal necrosis, gliosis, perivascular cuffing, and demyelination with astrocytosis and syncytia characterize the encephalitis. Lymphoid depletion is marked in acute infection. In the two pinniped cases of morbilliviral dermatitis, syncytia were prominent in the epidermis, follicular epithelium, and sebaceous glands, as were eosinophilic intracytoplasmic inclusions (Lipscomb et al., 2000a).
Diagnosis Diagnosis is based on the presence of characteristic histopathological lesions and is supported by immunocytochemistry and electron microscopy. Viral RNA can be detected in fixed or fresh tissue using a PCR-based assay. An antigen capture enzyme-linked immunosorbent assay (ELISA) for use on tissue homogenates is also available. Virus isolation in primary or secondary kidney cells can be difficult unless the carcass is fresh. Paired serum samples demonstrating an increasing antibody titer, or a single serum sample with a high IgM titer, may also be used to confirm morbillivirus infection.
Differentials PDV, PhHV-1, and influenza all cause pneumonia in pinnipeds, but the histopathology is somewhat distinct for each. PDV causes a bronchointerstitial pneumonia, PhHV-1 causes an interstitial pneumonia, and influenza A virus causes a bronchial pneumonia. Although other infectious organisms such as bacteria and parasites can cause pneumonia, that does not necessarily eliminate the possibility of a concurrent viral infection. Proper diagnosis is essential to confirm a viral etiology in any case.
0839_frame_C15 Page 298 Tuesday, May 22, 2001 11:11 AM
298
CRC Handbook of Marine Mammal Medicine
Epidemiology The apparent reservoirs for PDV and CMV are harp seals and pilot whales, respectively. Both these marine mammals are gregarious, migratory, and sufficiently numerous to sustain a morbillivirus within their populations. Morbilliviruses, in general, are readily transmitted from the respiratory tract via aerosol contamination. Prior to 1988, there is no serological evidence that European harbor seals had been exposed to PDV, so they were a naive population. Evidence does suggest that harbor seals may be more susceptible to PDV than gray seals, in part because of differential immune responses to viral antigens. Morbilliviruses, particularly CDV, can infect numerous species without necessarily causing clinical disease (Kennedy-Stoskopf, 1999). CDV was isolated from a clinically healthy captive gray seal in Canada (Lyons et al., 1993). CDV, however, has caused high mortalities in Baikal and Caspian seals. Interspecies transmission of marine morbilliviruses also occurs. Although the role morbilliviruses played in the Mediterranean monk seal deaths in 1997 remains controversial (Osterhaus et al., 1998; Chapter 2, Emerging and Resurging Diseases), virus isolated from three seals most closely resembled CMV sequences (van de Bildt et al., 1999). Manatees also had neutralizing antibodies to CMV, suggesting interspecies transmission occurred without overt clinical signs. Clinical disease from morbilliviruses in aberrant hosts has been attributed to predisposing factors, such as environmental contaminants, contributing to immunosuppression (see Chapter 22, Toxicology).
Public Health Significance No human health risk from the marine morbilliviruses is known.
Influenza Viruses Host Range Two influenza A viruses, subtypes H13N9 and H13N2, were isolated from a pilot whale (Globicephala melas) that stranded off the coast of Maine in 1984 (Hinshaw et al., 1986). An influenza A virus (H1N3) was isolated from minke whales caught in the South Pacific in 1975 to 1976 (Lvov et al., 1978). Four influenza A viruses have been isolated from harbor seals. The first, A/Seal/Mass/1/80 (H7N7), was isolated from harbor seals dying off the New England Coast in 1980 (Geraci et al., 1982). A mycoplasma was isolated concurrently from these animals. The second, A/Seal/Mass/ 133/82 (H4N5), was isolated from animals dying off the New England Coast between June 1982 and March 1983 (Hinshaw et al., 1984). In winter 1991–1992 two further isolates of subtypes H4N6 and H3N3 were cultured (Callan et al., 1995). Influenza B, B/Seal/Netherlands/1/99, was isolated from a throat swab of a juvenile seal with respiratory signs (Osterhaus et al., 2000).
Clinical Signs Clinical signs in the one pilot whale were nonspecific. The animal had difficulty maneuvering, was extremely emaciated, and was sloughing skin. Clinical signs during the original epizootic in harbor seals were dramatic. Well-nourished animals were weak, uncoordinated, dyspneic, and had conjunctivitis. Occasionally, a frothy white or bloody nasal discharge was observed. Common findings were swollen necks, due to air escaping through the thoracic inlet and becoming trapped within fascia and muscles. The harbor seal with influenza B recovered.
0839_frame_C15 Page 299 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
299
Therapy The epizootics have been so virulent that any attempts at serious supportive care were precluded.
Pathology In the pilot whale, the lungs were hemorrhagic, and a hilar node was greatly enlarged. In the seals, pneumonia was characterized by necrotizing bronchitis and bronchiolitis, and hemorrhagic alveolitis.
Diagnosis In both the pilot whale and seals with influenza A, virus was isolated from tissue homogenates of lung and hilar lymph nodes inoculated allantoically into embryonated chicken eggs (Lang et al., 1981; Hinshaw et al., 1984; 1986). Virus was also isolated from seal brains. Reference antisera to influenza hemagglutinin (H) and neuraminidase (N) antigens were used to characterize the isolates serologically. The influenza B isolate was initially cultured in MDCK cells and could be propagated in primary seal kidney cells. The diagnosis was confirmed by reverse-transcription PCR of RNA isolated from culture supernatant.
Differentials See morbillivirus differentials.
Epidemiology The influenza A viruses recovered from marine mammals off the northeastern coast of the United States are antigenically and genetically related to avian influenza virus strains. Both isolates from the pilot whale are related to H13 influenza viruses from gulls. The original seal isolate is antigenically related to some strains of influenza virus responsible for fowl plague; however, the seal isolate is nonpathogenic to poultry. The incubation period of the virus during epizootics seems to be 3 days or less. The isolates from harbor seals replicate in ferrets, cats, pigs, and phocid seals, including harbor, ringed, and harp seals inoculated experimentally (Webster et al., 1981a; Geraci et al., 1984; Hinshaw et al., 1984). Virus fails to replicate in gray seals experimentally inoculated even though antibodies to A/Seal/Mass/1/80 influenza virus have been detected in free-ranging animals (Geraci et al., 1984). In an attempt to reproduce clinical signs seen during the epizootics, seals have been inoculated with both influenza A and the mycoplasma isolated from symptomatic seals. Harp seals develop no overt signs of pneumonia even though a necrotizing bronchitis and alveolitis are present microscopically. Many interacting factors probably contribute to the explosive nature of the epizootics in the harbor seals, such as high population densities and unseasonably warm temperatures. Sequence analysis of the influenza B isolate showed it was closely related to strains present in the human population in 1995. This marks the first time that influenza B has been isolated from a species other than humans. Retrospective serosurveys demonstrated no antibodies to influenza B in the seal population around the Netherlands prior to 1995. Osterhaus et al. (2000) estimated that between 0.5 and 2% of the animals have subsequently been infected.
0839_frame_C15 Page 300 Tuesday, May 22, 2001 11:11 AM
300
CRC Handbook of Marine Mammal Medicine
Public Health Significance Four individuals performing post-mortem studies on seals during the initial epizootic developed a keratoconjunctivitis within 2 to 3 days of known eye contamination. An investigator developed a severe conjunctivitis 40 hours after an experimentally infected seal sneezed in his face (Webster et al., 1981a,b). An influenza virus identical to the one from the seal was recovered from conjunctival swabs of this person. All affected humans recovered completely within 7 days. None of these individuals developed any antibody titers to the virus, suggesting that the reaction was local, as occurs with Newcastle disease virus in the eyes of humans. However, the seal virus can replicate and cause systemic disease in squirrel monkeys inoculated intratracheally, indicating once again the zoonotic potential of these suspected avian viruses adapted to mammalian hosts (Murphy et al., 1983). Osterhaus et al. (2000) speculate that seals may serve as a reservoir for influenza B viruses that have circulated previously in the human population.
Caliciviruses (San Miguel Sea Lion Virus) Host Range Since 1972, more than 20 serotypes of caliciviruses have been isolated from a variety of marine mammals, including California sea lions, northern fur seals, northern elephant seals (Mirounga angustirostris), Pacific walruses (Odobenus rosmarus divergens), Steller sea lions, and bottlenose dolphins (for review, see Smith and Boyt, 1990; Barlough et al., 1998). A serotype is defined as an antigenic variant that is not neutralized by antiserum containing 20 times the antibody concentration needed to neutralize the homologous virus type. Additional marine mammals that have antibodies to marine calicivirus serotypes include Hawaiian monk seals (Monachus schauinslandi) and California gray (Eschrichtius robustus), sperm, fin (Balaenoptera physalus), sei, and bowhead whales.
Clinical Signs The most consistent lesion caused by caliciviruses in marine mammals is skin vesicles. In dolphins, vesicular lesions have been observed in association with “tattoo” lesions and old scars (Smith et al., 1983a). In pinnipeds, the vesicles are most prevalent on the dorsal surfaces of the flippers. Vesicles range from 1 mm to 3 cm in diameter, but may coalesce to form bullae. They usually erode leaving shallow, fast-healing ulcers, but occasionally vesicles regress leaving plaquelike lesions. Depending on the severity, lesions resolve between 1 and 9 weeks (Gage et al., 1990). Nonvesicular lesions at the mucocutaneous junction of the lips, around the nose, and on the chin have been observed in California sea lions with flipper lesions. The oral and facial lesions are firm, raised nodules that resemble pox lesions except the latter more commonly occur on the neck and chest and do not affect mucosal surfaces (Gage et al., 1990). Oral vesicles are also seen (Van Bonn et al., 2000). Premature parturition has been observed in association with the presence of caliciviruses in California sea lions. However, Leptospira pomona was also isolated, so it is not certain whether the virus or spirochete was responsible for early parturition (Gilmartin et al., 1976). Premature sea lion pups exhibited respiratory and locomotor difficulties and did not survive. Experimentally infected northern fur seal pups developed an interstitial pneumonitis and mild encephalitis, but no virus was recovered from lung or nervous tissue (Smith et al., 1980).
Therapy Skin lesions usually resolve without any supportive treatment.
0839_frame_C15 Page 301 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
301
Pathology Early vesicles form in the stratum spinosum, and spread by extension between the stratum lucidum and the basal layers of the epidermis.
Diagnosis Virus can be isolated from throat and rectal swabs or aspirated vesicular fluid placed in vials of phosphate buffered glycerol (pH 7.2) or culture medium supplemented with fetal bovine serum and then frozen. The virus forms plaques, 1 to 4 mm in diameter, on Vero cells (African green monkey kidney cell line) after several blind passages. Infected Vero cells prepared for thin-section electron microscopy show viral particles in tubular configurations. Individual virions have a distinctive morphology characterized by 22 calyces on their surface and measure 36 nm in diameter. The Laboratory for Calicivirus Studies, College of Veterinary Medicine, Oregon State University (Corvallis, OR) and Plum Island Animal Disease Center (Greenport, Long Island, NY) can serotype viral isolates and serum samples.
Epidemiology For a review, see Smith and Boyt (1990). The discovery that the original viral isolates from sea lions on San Miguel Island off the coast of California were indistinguishable from viruses responsible for vesicular exanthema of swine, a disease last seen in the United States in 1959, led to the hypothesis that marine mammals might have been the original source of the virus in swine in California in the 1930s, where it was endemic for 20 years before spreading to the rest of the country. Clinical disease was eliminated in swine by slaughtering affected animals and prohibiting the feeding of uncooked garbage. Considering that marine products have been used as a source of feed for swine, it was easy to postulate that sea lions were the original source of virus. However, the observation that the population of sea lions was too small to serve as a reservoir for so many different serotypes prompted a search for more likely candidates. This led to the discovery that opaleye perch (Girella nigricans), which overlap the southern range of California sea lions, were also infected with calicivirus serotypes that have been isolated from marine mammals. Opaleye can live for 10 years, and the virus can remain viable for at least 32 days. The suggestion has also been made that lungworm (Parafilaroides decorus) larvae that remain viable in the guts of fish for long periods of time may also harbor virus. Opaleye are probably responsible for the enzootic status of caliciviruses in marine mammals that inhabit the coastal waters of California. Serological studies have shown that by the time most sea lions are 4 months old, they have neutralizing antibodies to one or more serotypes. The reservoir of caliciviruses in the Arctic seas is not known. Originally, infections in Arctic marine mammals were considered transient and occurred only in species that migrated through endemic waters. However, the isolation of calicivirus from walrus negates the theory. So far, no caliciviruses or antibodies to them have been isolated from marine mammals in the Atlantic Ocean. This is not to imply they are not susceptible, but that a serotype has yet to establish itself in an appropriate reservoir in these waters. Multiple serotypes may infect one animal, and certain serotypes appear more pathogenic than others. A carrier state has been described in northern fur seals. Marine mammals have antibodies to terrestrial caliciviruses isolated from mink, dairy cattle, and reptiles housed in a California zoo. The reptilian calicivirus Crotalus type1 has been isolated from California sea lion, Steller sea lion, and northern fur seal (Barlough et al., 1998). Thus, the marine/terrestrial transmission of caliciviruses appears to remain ongoing.
0839_frame_C15 Page 302 Tuesday, May 22, 2001 11:11 AM
302
CRC Handbook of Marine Mammal Medicine
Public Health Significance There is no confirmation that marine caliciviruses cause clinical disease in humans. Yet the ability of these viruses to infect a wide variety of species argues that these agents should not be taken lightly. Researchers have developed neutralizing antibodies to two serotypes, suggesting either they were exposed to large, antigenic concentrations of virus or the virus actually replicated in their bodies (Smith et al., 1987). A “blisters on the eyes” syndrome was experienced by a biologist after working with fur seals exhibiting vesicular lesions on their flippers (Smith, 1987). The isolation of a calicivirus from primates with vesicular lesions and encephalitis is further proof that these viruses should be handled carefully (Smith et al., 1983b).
Other Viruses Hepadnavirus A hepatitis B-like infection has been diagnosed in a Pacific white-sided dolphin (Bossart et al., 1990). A 22-year-old female residing for 14 years in an oceanarium developed a sporadic cyclical pattern of lethargy, anorexia, and icterus with increased alanine transaminase (ALT), aspartate transaminase (AST), and γ-glutamyltransferase (GGT) activity. Complete blood counts ranged from normal to a leukocytosis characterized by a neutrophilia during clinical episodes. Initial presenting signs were multifocal, light brown skin lesions, 1 to 3 cm in diameter, over the dorsum that would slough with light digital pressure. The animal appeared photophobic. Treatment was supportive. Sera obtained from blood obtained on days 9, 20, 39, and 92 after clinical onset contained hepatitis B viral DNA, antihepatitis B virus core antigen, and hepatitis B surface antibody (anti-HBs). The clinical signs, serum biochemical findings, and serological data were consistent with chronic persistent hepatitis caused by a hepatitis B-like virus. Five years earlier, a Pacific white-sided dolphin in the same pool died after a short nonspecific illness from chronic–active hepatitis of unknown origin. A killer whale that shared the pool with the dolphins had only anti-HBs antibodies, indicating this animal had developed immunity. No serological markers were detected in sera from either other cetaceans in the park or humans in contact with the affected animal. Although there is antigen cross-reactivity between mammalian hepadnaviruses, host range is fairly restricted, suggesting cetaceans with a hepadnavirus hepatitis would not pose a public health risk.
Coronavirus Two adult harbor seals died without clinical signs and a third died acutely following a brief period of anorexia and abnormal behavior. They had been in the current exhibit with three California sea lions for 3 years. Acute necrotizing enteritis and pulmonary edema were consistent pathological findings in all three harbor seals (Bossart and Schwartz, 1990). Fluorescent antibody staining of small intestinal tissue from two of the affected seals was positive with antisera to porcine transmissible gastroenteritis virus, feline infectious peritonitis virus, and canine enteric coronavirus. No virus was isolated, but the gross, microscopic, and immunocytochemical findings were consistent with coronavirus infections in other mammals. The California sea lions remained asymptomatic.
Retrovirus The first retrovirus in a marine mammal was a Spumavirus isolated from a California sea lion that subsequently died with a pneumonia from which both Pasteurella multocida and an uncharacterized herpesvirus were isolated (Kennedy-Stoskopf et al., 1986). The Spumavirus
0839_frame_C15 Page 303 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
303
was consistently isolated from recurring skin lesions and adherent cells derived from peripheral blood mononuclear cells. Whether the virus actually caused the skin lesions is not known. Spumaviruses can cause persistent infections in numerous species, although they have not been linked to any particular disease.
Rhabdoviruses Osterhaus et al. (1993) isolated a rhabdo-like virus from the lungs and kidneys of a stranded white-beaked dolphin. This isolate was identified as a rhabdovirus based on ultrastructure by electron microscopy but was not recognized by antisera to three recognized genera of animal rhabdoviruses: rabies virus (genus Lyssavirus), bovine ephemeral fever virus (genus Ephemerovirus), and vesicular stomatitis virus (genus Vesiculovirus). Neutralizing antibodies have been detected in numerous cetacean species stranded on the coasts of northwestern Europe and the Mediterranean Sea between 1988 and 1992, but the significance remains to be determined. Rabies has been diagnosed in a ringed seal in Norway (Odegaard and Krogsrud, 1981). The animal was found wounded on the posterior part of the body and appeared confused. Its overall condition deteriorated during the next 5 days, and it became aggressive. Rabies was confirmed by immunofluorescent examination of the brain. At the time, there was an epizootic of rabies in foxes. Considering the ability of rabies to infect numerous mammalian species, it should be an added differential in areas where rabies is present in terrestrial mammals that can interact with marine mammals.
Acknowledgments The author thanks Carol House and Padraig Duignan for reviewing this chapter.
References Barlough, J.E., Matson, D.O., Skilling, D.E., Berke, T., Berry, E.S., Brown, R.F., and Smith, A.W., 1998, Isolation of reptilian calicivirus Crotalus type 1 from feral pinnipeds, J. Wildl. Dis., 34: 451–456. Barr, B., Dunn, L., Daniel, M.D., and Banford, A., 1989, Herpes-like viral dermatitis in a beluga whale (Delphinapterus leucas), J. Wildl. Dis., 25: 608–613. Borst, G.H.A., Walvoort, H.C., Reijnders, P.J.H., van der Kamp, J.S., and Osterhaus, A.D.M.E., 1986, An outbreak of a herpesvirus in harbor seals (Phoca vitulina), J. Wildl. Dis., 22: 1–6. Bossart, G.D., and Schwartz, J.C., 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., Cabal, C., Kuhns, M., Eimstad, E.A., Caron, J., Trimm, M., and Bradley, P., 1990, Hepatitis B-like infection in a Pacific white-sided dolphin (Lagenorhynchus obliquidens), J. Am. Vet. Med. Assoc., 196: 127–130. Bossart, G.D., Solorzano, J.L., Decker, S.J., and Cronell, L.H., 1996, Cutaneous papillomaviral-like papillomatosis in a killer whale (Orcinus orca), Mar. Mammal Sci., 12: 274–281. Britt, J.O., Nagy, A.Z., and Howard, E.B., 1979, Acute viral hepatitis in California sea lions, J. Am. Vet. Med. Assoc., 175: 921–923. Callan, R.J., Early, G., Kida, H., and Hinshaw, V.S., 1995, The appearance of H3N3 influenza viruses in seals, J. Gen. Virol., 76: 199–203. Collins, J.E., Leslie, P., Johnson, D., Nelson, D., Peden, W., Boswell, R., and Draayer, H., 1984, Epizootic of adenovirus infection in American black bears, J. Am. Vet. Med. Assoc., 185: 1430–1432. Daoust, P.Y., Taylor, R.G., and Greenlaw, B.L., 1994, Herpesvirus in botriomycotic lesions from a harp seal (Phoca groenlandica), Vet. Pathol., 31: 385–389.
0839_frame_C15 Page 304 Tuesday, May 22, 2001 11:11 AM
304
CRC Handbook of Marine Mammal Medicine
De Guise, S., Lagace, A., and Béland, P., 1994, Gastric papillomas in eight St. Lawrence beluga whales (Delphinapterus leucas), J. Vet. Diagn. Invest., 6: 385–388. De Guise, S., Lagace, A., Béland, P., Girard, C., and Higgins, R., 1995, Non-neoplastic lesions in beluga whales (Delphinapterus leucas) and other marine mammals from the St. Lawrence estuary, J. Comp. Pathol., 112: 257–271. Dierauf, L.A., Lowenstine, L.J., and Jerome, C., 1981, Viral hepatitis (adenovirus) in a California sea lion, J. Am. Vet. Med. Assoc., 179: 1194–1197. Duignan, P.J., 1999, Morbillivirus infections of marine mammals, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 497–501. Duignan, P.J., 2000, Diseases of New Zealand sea mammals, Surveillance, 27: 9–15. Flom, J.O., and Houk, E.J., 1979, Morphologic evidence of poxvirus in “tatoo” lesions from captive bottlenose dolphins, J. Wildl. Dis., 15: 593–596. Gage, L.J., Amaya-Sherman, L., Roletto, J., and Bently, S., 1990, Clinical signs of San Miguel sea lion virus in debilitated California sea lions, J. Zoo Wildl. Med., 21: 79–83. Garner, G.W., Evermann, J.F., Saliki, J.T., Follmann, E.H., and McKeirnan, A.J., 2000, Morbillivirus ecology in polar bears, Polar Biol., 23: 474–478. Geraci, J.R., Hicks, B.D., and St. Aubin, D.J., 1979, Dolphin pox: A skin disease of cetaceans, Can. J. Comp. Med., 43: 399–404. Geraci, J.R., St. Aubin, D.J., Barker, I.K., Webster, R.G., Hinshaw, V.S., Bean, W.J., Ruhnke, H.L., Prescott, J.H., Early, G., Baker, A.S., Madoff, S., and Schooley, R.T., 1982, Mass mortality of harbor seals: Pneumonia associated with influenza A virus, Science, 215: 1129–1131. Geraci, J.R., St. Aubin, D.J., Barker, I.K., Hinshaw, V.S., Webster, R.G., and Ruhnke, H.L., 1984, Susceptibility of grey (Halichoerus grypus) and harp (Phoca groenlandica) seals to the influenza virus and mycoplasma of epizootic pneumonia of harbor seals (Phoca vitulina), Can. J. Fish. Aquat. Sci., 41: 151–156. Geraci, J.R., Palmer, N.C., and St. Aubin, D.J., 1987, Tumors in cetaceans: Analysis and new findings, Can. J. Fish. Aquat. Sci., 44: 1289–1300. Gilmartin, W.G., Delong, R.L., Smith, A.W., Sweeney, J.C., De Lappe, B.W., Risebrough, R.W., Griner, L.A., Dailey, M.D., and Peakall, D.B., 1976, Premature parturition in the California sea lion, J. Wildl. Dis., 12: 104–115. Gulland, F.M.D., Lowenstine, L.J., Lapointe, J.M., Sparker, T., and King, D.P., 1997, Herpesvirus infection in stranded harbor seals of coastal California, J. Wildl. Dis., 33: 450–458. Gulland, F.M.D., Haulena, M., Lowenstine, L.J., Munro, C., Graham, P.A., Baulman, J., and Harvey, J., 1999, Adrenal function in wild and rehabilitated Pacific harbor seals (Phoca vitulina richardii) and in seals with phocine herpesvirus-associated adrenal necrosis, Mar. Mammal Sci., 15: 810–827. Hadlow, W.J., Cheville, N.F., and Jellison, W.L., 1980, Occurrence of pox in a northern fur seal on the Pribilof Islands in 1951, J. Wildl. Dis., 16: 305–312. Harder, T.C., and Osterhaus, A.D.M.E., 1997, Molecular characterization and baculovirus expression of the glycoprotein B of a seal herpesvirus (phocid herpesvirus-1), Virology, 227: 343–352. Harder, T.C., Harder, M., Vos, H., Kulonen, K., Kennedy-Stoskopf, S., Liess, B., Appel, M.J.G., and Osterhaus, A.D.M.E., 1996, Characterization of phocid herpesvirus-1 and -2 as putative alphaand gammaherpesviruses of North American and European pinnipeds, J. Gen. Virol., 77: 27–35. Harder, T.C., Vos, H.W., de Swart, R.L., and Osterhaus, A.D.M.E., 1997, Age-related disease in recurrent outbreaks of phocid herpesvirus type-1 infections in a seal rehabilitation center: Evaluation of diagnostic methods, Vet. Rec., 140: 500–503. Hicks, B.D., and Worthy, G.A., 1987, Sealpox in captive grey seals (Halichoerus grypus) and their handlers, J. Wildl. Dis., 23: 1–6. Hinshaw, V.S., Bean, W.J., Webster, R.G., Rehg, J.E., Fiorelli, P., Early, G., Geraci, J.R., and St. Aubin, D.J., 1984, Are seals frequently infected with avian influenza viruses? J. Virol., 51: 863–865. Hinshaw, V.S., Bean, W.J., Geraci, J., Fiorelli, P., Early, G., and Webster, R.G., 1986, Characterization of two influenza A viruses from a pilot whale, J. Virol., 58: 655–656.
0839_frame_C15 Page 305 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
305
Housawi, F.M.T., Roberts, G.M., Gilray, J.A., Pow, I., Reid, H.W., Nettleton, P.F., Sumption, K.J., Hibma, M.H., and Mercer, A.A., 1998, The reactivity of monoclonal antibodies against orf virus with other parapoxviruses and the identification of a 39 kDa immunodominant protein, Arch. Virol., 143: 2289–2303. Kennedy, S., 1998, Morbillivirus infections in aquatic mammals, J. Comp. Pathol., 119: 201–225. Kennedy, S., Lindstedt, I.J., McAliskey, M.M., McConnell, S.A., and McCullough, S.J., 1992, Herpesviral encephalitis in a harbor porpoise (Phocoena phocoena), J. Zoo Wildl. Med., 23: 374–379. Kennedy, S., Kuiken, T., Jepson, P.D., Deaville, R., Forsyth, M., Barrett, T., van de Bildt, M.W.G., Osterhaus, A.D.M.E., Eybatov, T., Duck, C., Kydyrmanov, A., Mitrofanov, I., and Wilson, S., 2000, Mass die-off of Caspian seals caused by canine distemper virus, Emerging Infect. Dis., 6: 637–639. Kennedy-Stoskopf, S., 1999, Emerging viral infections in large cats, in Zoo and Wild Animal Medicine Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 401–410. Kennedy-Stoskopf, S., Stoskopf, M.K., Eckhaus, M.A., and Strandberg, J.D., 1986, Isolation of a retrovirus and a herpesvirus from a California sea lion, J. Wildl. Dis., 22: 156–164. Kennedy-Stoskopf, S., Stoskopf, M.K., and Bunton, T.E., 1988, Herpesvirus isolation from a grey seal (Halichoerus grypus), in Proceedings of the American Association of Zoo Veterinarians and American Association of Wildlife Veterinarians, Toronto, Ontario, 84. King, D.P., Parselles, R., Gulland, F.M.D., Lowenstine, L.J., Ferrick, D.A., and Stott, J.L., 1998, Antigenic and nucleotide characterization of a herpesvirus isolated from Pacific harbor seals (Phoca vitulina richardsii), Arch. Virol., 143: 2021–2027. Lambertsen, R.H., Kohn, B.A., Sundberg, J.P., and Buergelt, C.D., 1987, Genital papillomatosis in sperm whale bulls, J. Wildl. Dis., 23: 361–367. Lang, G., Gagnon, A., and Geraci, J.R., 1981, Isolation of an influenza A virus from seals, Arch. Virol., 68: 189–195. Lebich, M., Harder, T.C., Frey, H.-R., Visser, I.K.G., Osterhaus, A.D.M.E., and Liess, B., 1994, Comparative immunological characterization of type-specific and conserved B-cell epitopes of pinniped, felid and canid herpesviruses, Arch. Virol., 136: 335–347. Lipscomb, T.P., Habecker, P.L., Damback, D.M., and Schoelkopf, R., 1996, Genital herpes infection in a male harbor porpoise, in Proceedings of the International Association for Aquatic Animal Medicine, 17. Lipscomb, T.P., Mense, M.G., Habecker, P.L., Taubenberger, J.K., and Schoelkopf, R., 2000a, Morbilliviral dermatitis in seals, in Proceedings of the Joint Conference of American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine, New Orleans, LA, 337–338. Lipscomb, T.P., Scott, D.P., Garber, R.L., Krafft, A.E., Tsai, M.M., Lichy, J.H., Taubenberger, J.K., Schulman, F.Y., and Gulland, F.M.D., 2000b, Common metastatic carcinoma of California sea lions (Zalophus californianus): Evidence of genital origin and association with novel gammaherpesvirus, Vet. Pathol., 37: 609–617. Lvov, D.K., Zdanov, V.M., Sazonov, A.A., Braude, N.A., Vladimirtceva, E.A., Agafonova, L.V., Skljanskaja, E.I., Kaverin, N.V., Reznik, V.I., Pysina, T.V., Oserovic, A.M., Berzin, A.A., Mjasnikova, I.A., Podcernjaeva, R.Y., Klimenko, S.M., Andrejev, V.P., and Yakhno, M.A., 1978, Comparison of influenza viruses isolated from man and from whales, Bull. World Health Organ., 56: 923–930. Lyons, C., Welsh, M.J., Thorsen, J., Ronald, K., and Rima, B.K., 1993, Canine distemper virus isolated from a captive seal, Vet. Rec., 132, 487–488. 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. Mikaelian, I., Tremblay, M.P., Montipetit, C., Tessaro, S.V., Cho, H.J., House, C., Measures, L., and Martineau, D., 1999, Seroprevalence of selected viral infections in a population of beluga whales (Delphinapterus leucas) in Canada, Vet. Rec., 144: 50–51. Murphy, B.R., Harper, J., Lewis Sly, D., London, W.T., Miller, N.T., and Webster, R.G., 1983, Evaluation of the A/Seal/Mass/1/80 virus in squirrel monkeys, Infect. Immunity, 42: 424–426.
0839_frame_C15 Page 306 Tuesday, May 22, 2001 11:11 AM
306
CRC Handbook of Marine Mammal Medicine
Nettleton, P.F., Munro, R., Pow, I., Gilray, J., Gray, E.W., and Reid, H.W., 1995, Isolation of a parapoxvirus from a grey seal (Halichoerus grypus), Vet. Rec., 137: 562–564. Odegaard, O.A., and Krogsrud, J., 1981, Rabies in Svalbard: Infection diagnosed in arctic fox, reindeer and seal, Vet. Rec., 109: 141–142. Osterhaus, A.D.M.E., and Vedder, E.J., 1988, Identification of virus causing recent seal deaths, Nature, 335: 20. Osterhaus, A.D.M.E., Yang, H., Spijkers, H.E.M., Groen, J., Teppema, J.S., and van Steenis, G., 1985, The isolation and partial characterization of a highly pathogenic herpesvirus from the harbor seal (Phoca vitulina), Arch. Virol., 86: 239–251. Osterhaus, A.D.M.E., Broeders, H.W.J., Visser, I.K.G., Teppema, J.S., and Vedder, E.J., 1990, Isolations of an orthopoxvirus from pox-like lesions of grey seal (Halichoerus grypus), Vet. Rec., 127: 91–92. Osterhaus, A.D.M.E., Broeders, H.W.J., Teppema, J.S., Kuiken, T., House, J.A., Vos, H.W., and Visser, I.K.G., 1993, Isolation of virus with rhabdovirus morphology from a white-beaked dolphin (Lagenorhynchus albirostris), Arch. Virol., 133: 189–193. Osterhaus, A.D.M.E., Broeders, H.W.J., Visser, I.K.G., Teppema, J.S., and Kuiken, T., 1994, Isolation of a parapoxivirus from pox-like lesions in grey seals, Vet. Rec., 135: 601–602. Osterhaus, A., van de Bildt, M., Vedder, L., Martina, B., Niesters, H., Vos, J., van Egmond, H., Liem, D., Baumann, R., Androukaki, E., Kotomatas, S., Komnenou, A., Sidi, B.A., Jiddou, A.B., and Barham, M.E.O., 1998, Monk seal mortality: Virus or toxin? Vaccine, 16: 979–981. Osterhaus, A.D.M.E., Rimmelzwaan, G.F., Martina, B.E., Bestebroer, T.M., and Fouchier, R.A, 2000, Influenza B virus in seals, Science, 288: 1051–1053. Palmer, E.L., and Martin, M.J., 1988, Poxviridae, in Electron Microscopy in Viral Diagnosis, CRC Press, Boca Raton, FL, chap. 23. Pursell, A.R., Stuart, B.P., Styer, E., and Case, J.L., 1983, Isolation of an adenovirus from black bear cubs, J. Wildl. Dis., 19: 269–271. Smith, A.W., 1987, San Miguel sea lion virus, in Virus Infections of Carnivores, Appel, M. (Ed.), Elsevier Science Publication, Amsterdam, 481–489. Smith, A.W., and Boyt, P.M., 1990, Calciviruses of ocean origin: A review, J. Zoo Wildl. Med., 21: 3–23. Smith, A.W., and Skilling, D.E., 1979, Viruses and virus diseases of marine mammals, J. Am. Vet. Med. Assoc., 175: 918–920. Smith, A.W., Skilling, D.E., and Brown, R.J., 1980, Preliminary investigation of a possible lung worm (Parafilaroides decorus), fish (Girella nigricans), and marine mammals (Callorhinus ursinus) cycle for San Miguel sea lion virus type 5, Am. J. Vet. Res., 41: 1846–1850. Smith, A.W., Skilling, D.E., Ridgway, S.H., and Fenner, C.A., 1983a, Regression of cetacean tattoo lesions concurrent with conversion of precipitin antibody against a poxvirus, J. Am. Vet. Med. Assoc., 183: 1219–1222. Smith, A.W., Skilling, D.E., Ensley, P.K., Benirschke, K., and Lester, T.L., 1983b, Calicivirus isolation and persistence in a pygmy chimpanzee (Pan paniscus), Science, 221: 79–81. Smith, A.W., Prato, C., and Skilling, D.E., 1987, Caliciviruses infecting monkeys and possibly man, Am. J. Vet. Res., 39: 287–289. Stenvers, O., Plötz, J., and Ludwig, H., 1992, Antarctic seals carry antibodies against seal herpesvirus, Arch. Virol., 123: 421–424. Stuen, S., Have, P., Osterhaus, A.D.M.E., Ardemo, J.M., and Moustgard, A., 1994, Serological investigations of virus infections in harp seals (Phoca groenlandica) and hooded seals (Cystophora cristata), Vet. Rec., 134: 502–505. Van Bonn, W., Jensen, E.D., House, C., House, J.A., Burrage, T., and Gregg, D.A., 2000, Epizootic vesicular disease in captive California sea lions, J. Wildl. Dis., 36: 500–507. Van Bressem, M.-F., and Van Waerebeek, K., 1996, Epidemiology of poxvirus in small cetaceans from the eastern South Pacific, Mar. Mammal Sci., 12: 371–382.
0839_frame_C15 Page 307 Tuesday, May 22, 2001 11:11 AM
Viral Diseases
307
Van Bressem, M.-F., Van Waerebeek, K., Reyes, J.C., Dekegel, D., and Pastoret, P.-P., 1993, Evidence of poxvirus in dusky dolphin (Lagenorhynchus obscurus) and Burmeister’s (Phocoena spinipinnis) from coastal Peru, J. Wildl. Dis., 29: 109–113. Van Bressem, M.-F., Van Waerebeek, K., Garcia-Godos, A., Dekegel, D., and Pastoret, P.-P., 1994, Herpes-like virus in dusky dolphins Lagenorrhynchus obscurus, from coastal Peru, Mar. Mammal Sci., 10: 354–359. Van Bressem, M.-F., Van Waerebeek, K., Pierard, G.E., and Desaintes, C., 1996, Genital and lingual warts in small cetaceans from coastal Peru, Dis. Aquat. Organisms, 26: 1–10. Van Bressem, M.-F., Waerebeek, K.V., Fleming, M., and Barrett, T., 1998, Serological evidence of morbillivirus infection in small cetaceans from the Southeast Pacific, Vet. Microbiol., 59: 89–98. Van Bressem, M.-F., Kastelein, R.A., Flamant, P., and Orth, G., 1999a, Cutaneous papillomavirus infection in a harbor porpoise (Phocoena phocoena) from the North Sea, Vet. Rec., 144: 592–593. Van Bressem, M.-F., Van Waerebeek, K., and Raga, J.A., 1999b, A review of virus infections of cetaceans and the potential impact of morbilliviruses, poxviruses and papillomaviruses on host population dynamics, Dis. Aquat. Organisms, 38: 53–65. van de Bildt, M.W.G., Vedder, E.J., Martina, B.E.E., Sidi, B.A., Jiddou, A.B., Barham, M.E.O., Androukaki, E., Komnenou, A., Niesters, H.G.M., and Osterhaus, A.D.M.E., 1999, Morbilliviruses in Mediterranean monk seals, Vet. Microbiol., 69: 19–21. Webster, R.G., Hinshaw, V.S., Bean, W.J., Van Wyke, K.L., Geraci, J.R., St. Aubin, D.J., and Petursson, G., 1981a, Characterization of an influenza A virus from seals, Virology, 113: 712–724. Webster, R.G., Geraci, J., Petursson, G., and Skirnisson, K., 1981b, Conjunctivitis in human beings caused by influenza A virus of seals, N. Engl. J. Med., 304: 911. Wilson, T.M., and Poglayen-Neuwall, I., 1971, Pox in South American sea lions, Otaria byronia, Can. J. Comp. Med., 35: 174–177. Wilson, T.M., Cheville, N.F., and Karstad, L., 1969, Seal pox, Bull. Wildl. Dis. Assoc., 5: 412–418. Wilson, T.M., Boothe, A.D., and Cheville, N.F., 1972a, Sealpox field survey, J.Wildl. Dis., 8: 158–160. Wilson, T.M., Dykes, R.W., and Tsai, K.S., 1972b, Pox in young, captive harbor seals, J. Am. Vet. Med. Assoc., 161: 611–617. Zarnke, R.L., Harder, T.C., Vos, H.W., Ver Hoef, J.M., and Osterhaus, A.D.M.E., 1997, Serologic survey for phocid herpesvirus-1 and -2 in marine mammals from Alaska and Russia, J. Wildl. Dis., 33: 459–465.
0839_frame_C15 Page 308 Tuesday, May 22, 2001 11:11 AM
0839_frame_C16 Page 309 Tuesday, May 22, 2001 11:12 AM
16
Bacterial Diseases of Cetaceans and Pinnipeds J. Lawrence Dunn, John D. Buck, and Todd R. Robeck
Introduction In the decade since the preparation of the chapter on bacterial and mycotic diseases for the first edition of this book, innumerable advances have improved our ability to diagnose, treat, and prevent many infectious diseases. Unfortunately, despite this significant progress, new challenges have arisen to test the ability of practitioners to deal with serious bacterial infections. Bacterial disease secondary to primary conditions, such as those caused by morbilliviruses and phytotoxins, are now well recognized as major causes of mortality in wild marine mammal populations. In addition, other bacterial pathogens are infecting new or previously unrecognized hosts (Hernandez et al., 1998; Harvell et al., 1999). Diseases (such as those caused by mycobacterial species), which were seldom referenced in the earlier marine mammal literature, have emerged as significant causes of morbidity and mortality in both wild and captive populations of marine mammals (see Chapter 2, Emerging Diseases). Brucellosis, long thought to be a disease confined to terrestrial mammals, has been documented in marine mammals in several areas of the world. Increasingly, there are reports of altered immune response and a decrease in natural resistance to bacterial and viral infection in marine mammals exposed to high levels of anthropogenic substances such as organohalogens (Thompson and Hall, 1993; De Guise et al., 1995a; Parsons and Jefferson, 2000). Dissenting positions regarding this relationship also exist, and it is unfortunate that, although details of tissue contaminant levels in marine mammals abound, the ultimate consequences of these elevated levels remain speculative (Wilson et al., 1999; see Chapter 22, Toxicology). In marine mammals held in aquaria or oceanaria, as well as those cared for as stranded animals, antibiotic resistance is now recognized much more frequently than in the past (Johnson et al., 1998). This is occurring, at least in part, as a result of misuse of these pharmaceuticals by all segments of the health-care community. Many factors make it difficult to quantify the role of bacterial disease in wild marine mammal mortalities. Debilitating bacterial disease increases the likelihood of predation of affected individuals, and this may result in an underrepresentation of such mortalities in surveys of stranded marine mammals. Alternatively, post-mortem colonization of organ systems of dead, beached animals with gastrointestinal, environmental, or surface microflora might 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
309
0839_frame_C16 Page 310 Tuesday, May 22, 2001 11:12 AM
310
CRC Handbook of Marine Mammal Medicine
result in positive bacterial cultures, possible misdiagnoses, and subsequent overrepresentations of bacteria-associated mortalities. The recovery of pathogenic bacteria from a dead animal is in no way proof that those recovered organisms are the direct cause of the animal’s death (Howard et al., 1983). This is demonstrated by the fact that recognized bacterial pathogens are routinely recovered from the upper respiratory or lower gastrointestinal systems of healthy captive marine mammals. Thornton et al. (1998) suggest that there are host species differences in the relative frequency of bacterial isolations from stranded pinnipeds, although age classes and immunity status may also contribute to the differences. Recent advances in diagnostic identification of viral diseases and phytointoxications have resulted in a significant increase in reports of stranded marine mammals diagnosed with one or more of these conditions as a primary problem (Hernandez et al., 1998). Bacterial or fungal infections frequently complicate the interpretation of the pathological findings in these cases (Lipscomb et al., 1994). Demographic and geographic changes in the populations of several species that have been encountered in the northeastern United States as stranded animals have altered the relative rankings of the causes of death in these taxa. Despite these findings and changes in ranking, bacterial disease, whether primary or secondary, persists as one of the leading causes of death in both wild and captive marine mammals. Buck et al. (1988; 1991) described the bacterial microflora of stranded cetaceans from the northeastern United States and southwestern Florida; earlier, Stroud and Jaffe (1979) reported that bacterial infection accounted for the deaths of 27% of the stranded marine mammals they examined. Johnson et al. (1998) cataloged 19 bacterial genera from three species of stranded pinnipeds, and Parsons and Jefferson (2000) implicated preexisting disease or bacterial infection in the deaths of 29% of the cetaceans they examined from Hong Kong waters. Higgins (2000), in his review, gives a comprehensive listing of bacteria and fungi isolated from marine mammals and notes the host and the organ system from which the isolate was obtained. One also must consider the possible interplay between microorganisms and the development of neoplastic disease, as has been reported in human medicine. The goal of this chapter is to acquaint the reader briefly with the more common, clinically important bacterial diseases of both wild and captive pinnipeds and cetaceans. This will provide a starting point from which the interested reader may begin a more vigorous literature review in areas of special interest. A number of older reports are cited for historical interest, or because knowledge of some conditions has not changed dramatically from the dates of their publication. Newer, as-yet-unpublished information from the authors’ experiences is also included here.
Microbial Sampling Techniques Diagnostic techniques and necropsy protocols are covered in detail elsewhere in this volume (see Chapter 19, Clinical Pathology; Chapter 21, Necropsy), but a brief overview of common sampling and specimen-processing techniques is worthwhile. Microbial sampling is greatly facilitated when the animals being examined have been trained to perform husbandry behaviors, such as exhaling onto an agar plate, opening the mouth for pharyngeal swabbing, swallowing a nasogastric tube for gastric fluid cytology or pH, urinating on command, and allowing urinary catheterization without struggle. Unfortunately, not all animals have had such training, or they are unwilling to perform these behaviors when ill; hence, sampling techniques that involve physical or chemical restraint may be required.
0839_frame_C16 Page 311 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
311
At Mystic Aquarium one of author’s (J.L.D.) routine microbial sampling procedure for cetaceans include anal swabs, skin swabs, deep blowhole swabs, vaginal or preputial swabs, pharyngeal swabs, and blowhole expiration plates. Any wounds or drainage tracts are sampled. In necropsy specimens, thoracic and abdominal cavities are swabbed, and major organs are superficially flamed and then sampled deeply. In pinnipeds, a similar sampling scheme is followed, except that nasal swabs are taken, and investigators are usually forced to settle for oral swabs rather than pharyngeal swabs in living, nonanesthetized animals. Swabs or other samples for microbial examination must be processed immediately, if possible. However, isolated strandings and other events may preclude prompt laboratory examination. If expedient processing is not possible, the authors recommend that swab transport systems using Stuart’s or Amies diluent be maintained at 20 to 25°C until laboratory examination, rather than being refrigerated. Culture techniques employ methods discussed in the literature (Buck, 1982; Buck and Spotte, 1986a; Buck et al., 1988; 1991). The National Animal Disease Laboratory at Ames, Iowa, is very helpful in assisting in the identification of species that are difficult to type or culture. Local Animal and Plant Health Inspection Service (APHIS) agents from the U.S. Department of Agriculture are helpful in securing this service. Briefly, these techniques are as follows. Swabs for aerobic bacterial cultures are streaked onto blood agar (Columbia agar base plus 5% sheep blood), Hektoen agar, MacConkey agar, mannitol salt agar, and thiosulfate citrate bile salt agar (TCBS), and then incubated aerobically at 35 to 37°C for 24 to 48 hours. Brucella isolations may be attempted using Brucella agar plates incubated under CO2 at 35 to 37°C. Anaerobes are cultured on blood agar in Gas Pak Jars at 35 to 37°C, and individual culture isolates are maintained in thioglycolate broth. Gram-negative, cytochrome-oxidase-negative bacteria are identified by means of API-20E (BioMérieux, Hazelwood, MO) strips. These are also useful in identifying sucrose-positive or -negative vibrios from TCBS plates, but additional tests, such as growth in varying sodium chloride (NaCl) concentrations, may be required. West and Colwell (1984) provide another overview of identification schemes for Vibrionaceae. Most Gram-positive species are identified by standard colonial morphology and microscopic characteristics. Staphylococci can be identified using the API Staph-Ident system (BioMérieux) and the coagulase reaction. The API Strep system (BioMérieux) is helpful in separating streptococci and enterococci, as are a variety of rapid enzyme kits. These, as well as rapid and inexpensive traditional culture methods, have become important in identifying enterococci, which have emerged as important nosocomial pathogens in humans and presumably, in marine mammals, as well. Howard et al. (1983) recommended PPLO media for mycoplasma, but until the pinniped viral epidemics of the 1970s and 1980s, when comprehensive sampling for pathogens of all types was conducted, the agent was infrequently diagnosed in marine mammals (Geraci et al., 1982; Baker et al., 1998; Measures, 1998). Mycoplasma has been isolated from human patients after seal bites (Baker et al., 1998) and now is widely considered to be the agent responsible for “seal finger” (see Chapter 34, Public Health). Tuberculous, atypical mycobacteria, and other “higher” bacteria may offer culture and diagnostic challenges. Prolonged incubation in media such as Lowenstein Jensen agar at room temperature, as well as at body temperature, in reduced and normal room light, may be required before mycobacterial identification is possible. Impression smears of lesions and acid-fast staining, or the use of molecular biological techniques, will often help speed a presumptive diagnosis of these and other bacterial pathogens and permit a more rapid employment of appropriate treatment regimens.
0839_frame_C16 Page 312 Tuesday, May 22, 2001 11:12 AM
312
CRC Handbook of Marine Mammal Medicine
Specific Bacterial Diseases of Cetaceans and Pinnipeds Septicemia Recognition of bacterial disease in marine mammals frequently presents a diagnostic challenge. Marine mammals, like many other wild animals, often fail to show obvious signs of disease until their condition has deteriorated to the point that available veterinary assistance may be ineffective. Nowhere is this failure to show obvious signs of illness more frustrating than in the case of septicemia. Septicemic crises may develop so rapidly that it is not unusual to have an animal die within hours of eating well and acting in a manner not dissimilar to that of its pool mates. The problems involved in trying to recognize disease in animals that spend all, or most, of their lives in and under the water are manifold. Close observation by experienced personnel is extremely useful, but sometimes impossible, as some facilities may lack good underwater viewing. Husbandry behaviors taught to animals that assist in clinical examinations are useful, but unfortunately, are often not performed if an animal is seriously ill. In cases where clinical signs indicate a problem, even prompt initiation of appropriate therapy is often not successful. The increased prevalence of Gram-negative bacterial infections in marine mammals increases the odds that endotoxemia will develop. Use of antibiotics with less-than-rapid bactericidal activity may exacerbate the situation and actually promote the release of endotoxin and the induction of septic shock. Fortunately, the quinolones, aminoglycosides, and later-generation cephalosporins commonly employed in marine mammal medicine do not exhibit this propensity. In cases of non-peracute death, significant necropsy findings may involve many organ systems and may include petechial and ecchymotic hemorrhage and diffuse intravascular coagulation. In human medicine, multiple organ system dysfunction subsequent to septic shock is the most common cause of death in noncoronary intensive care units. With severe bacterial sepsis, especially that associated with shock, mortalities are close to 50%, despite appropriate antimicrobial therapy and optimum supportive care (Munford, 1994). At Mystic Aquarium, Gram-negative bacteria predominate in isolates from diagnosed cases of marine mammal septicemias, but Streptococcus, Staphylococcus, and Erysipelothrix have also been recovered. Streptococcus zooepidemicus septicemia has been associated with mortalities at a number of facilities holding Tursiops spp. Gram-negative isolates associated with fatal septicemia at Mystic have included bacteria in the following genera: Vibrio, Edwardsiella, Aeromonas, Pseudomonas, Pasteurella, and Klebsiella. These pathogens are generally cultured from most, or all, major organ systems sampled. Howard et al. (1983) described gross and microscopic lesions associated with one form of marine mammal septicemia. Other authors (Cusick and Bullock, 1973; Medway and Schryver, 1973; Coles et al., 1978; Sweeney, 1978; Cohen et al., 1993) have reported septicemias in marine mammal that have implicated the bacterial species listed above, as well as additional species, including Salmonella spp., Corynebacterium spp., Pseudomonas spp., P. pseudomallei, Klebsiella spp., Pasteurella hemolytica, and V. alginolyticus. Cowan et al. (1998) stressed that the high pathogenicity of many of the bacteria isolated from bacteremic stranded cetaceans contributes to the high mortality seen in these animals.
Brucellosis Beginning in 1994, reports from the United Kingdom, the United States, Canada, and Norway described either the isolation of a Brucella spp. from marine mammals or serological evidence of current or prior exposure to organisms serologically cross-reactive with Brucella (Foster et al., 1996; Nielsen et al., 1996; Ross et al., 1996; Garner et al., 1997; Jepson et al., 1997; Clavareau et al., 1998; Forbes et al., 2000). Recently, an item in the popular press reported the
0839_frame_C16 Page 313 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
313
presence of antibodies against Brucella in 5 of 16 fur seals (Arctocephalus gazella) and one Weddell seal (Leptonychotes weddellii) from the Antarctic Continent. Although initially these reports dealt only with infection in wild populations, Miller et al. (1999) documented cases in captive marine mammals. Subsequently, there was a report of exposure, illness, seroconversion, and eventual recovery in a human researcher working with Brucella strains recovered from marine mammals (Brew et al., 1999). The patient in this case responded positively to a 6-week course of rifampin and doxycycline. Analyses of genetic relatedness among Brucella isolates from marine mammals show considerable differences from the profiles of recognized terrestrial Brucella species. Brucella cultured from dolphins in the Atlantic and Pacific Oceans showed a close relationship to each other, but differed significantly from isolates from porpoises from the same areas. Brucella recovered from seals showed similar profiles for isolates from both Atlantic and Pacific Ocean pinnipeds. No doubt, it will be some time before microbial taxonomists attach identifying labels to these isolates and it is possible to determine whether one is dealing with one or several biovars or species of Brucella (Ewalt et al., 1994; Jahans et al., 1997; Clavareau et al., 1998; Jensen et al., 1999; Bricker et al., 2000). Diagnosis of Brucella infection in marine mammals is fraught with obstacles. Isolation and subsequent culture of this pathogen may require specialized media and culture conditions. Depending on the culture conditions and the media employed, the incubation time of some strains may be so lengthy that culture plates and tubes might be discarded before growth is visible (Miller et al., 1999). Serological diagnosis of exposure of marine mammals to Brucella spp. is also potentially problematic, as there is controversy about the degree of agreement between newer enzyme-linked immunosorbent assays (ELISA) techniques and some of the classical techniques used in agricultural species. The older techniques may produce an increased number of false positives when employed on samples from marine mammals. In domestic animals, infection with any of several species of bacteria may result in Brucella-positive agglutination and complement fixation reactions (Weynants et al., 1995). The standard tube agglutination and rivanol tests lack sensitivity in detecting exposure to some marine-origin Brucella spp., so meaningful interpretation of ELISA data will require additional research. Miller et al. (1999) caution that, as yet, no “definitive positive correlation of Brucella infection in marine mammals with available Brucella serologic tests developed and approved for cattle and swine has been established.” Despite the present uncertainty about diagnostic tests for marine-origin Brucella, it seems likely that the success seen with fluorescence polarization assay, indirect immunoassay, and competitive immunoassay in diagnosing B. abortus will prove useful in these endeavors (Gall et al., 2000). At present, there is a dearth of information regarding the pathophysiology of infection with marine-origin Brucella. The limited clinical experience with this pathogen and knowledge of the pathobiology of other Brucella species suggest that infection with this organism presents an increased possibility of the development of, at least, transitory reproductive disorders. This could adversely impact efforts at increasing genetic diversity in sparsely populated captive collections of some marine mammals and could especially impact populations of severely endangered species such as the vaquita (Phocoena sinus). In addition, those dealing with marine mammals or their tissues should not discount the potential zoonotic implications of infection with this organism. Cetaceans
Serological evidence of exposure to Brucella has been documented in harbor porpoises (Phocoena phocoena), bottlenose dolphins (Tursiops truncatus), orcas (Orcinus orca), common dolphins (Delphinus delphis), pilot whales (Globicephala spp.), and striped dolphins (Stenella
0839_frame_C16 Page 314 Tuesday, May 22, 2001 11:12 AM
314
CRC Handbook of Marine Mammal Medicine
coeruleoalba). Thus far, of the baleen whales sampled—minke (Balaenoptera acutorostrata), fin (B. physalus), and sei (B. borealis) whales—only a single minke whale has shown a positive titer. Isolations of Brucella spp. have also been made from harbor porpoises, common, Atlantic white-sided (Lagenorhynchus acutus), and bottlenose dolphins (Clavareau et al., 1998). In addition to the recovery of Brucella spp. from wild cetaceans, there have been isolations from captive animals. Specimens from aborted premature bottlenose dolphin fetuses and from the lung of an adult bottlenose dolphin maintained in the same facility were culture-positive for Brucella. At least one of the aborting females has subsequently become pregnant and delivered a healthy calf (Miller et al., 1999). Isolation attempts from blood, milk, and placental samples from most of the eight dolphins exposed to one of the aborting females were negative. The later delivery by these animals of either a premature fetus or a weak calf that survived only a few minutes is problematic, but not diagnostic. Serum from a juvenile stranded bottlenose dolphin diagnosed with osteomyelitis of one of its caudal vertebrae tested positive for Brucella, using all standard serological testing modalities, as well as the newer ELISA technique. Attempts at isolation of a causative agent were initially unsuccessful, but positive at necropsy. Temporary regression of clinical signs followed treatment with third-generation cephalosporins after a failure of response to intralesional and oral doxycycline (Dunn, unpubl. data). Pinnipeds
Thus far, Brucella spp. have been isolated from several species of phocid seals—harbor (Phoca vitulina), hooded (Cystophora cristata), gray (Halichoerus grypus), ringed (P. hispida), and harp (Pagophilus groenlandicus) seals—and serological evidence of exposure exists in Atlantic walruses (Odobenus rosmarus) and Antarctic fur, harp, hooded, ringed, and Weddell seals. Pinnipeds from the North Atlantic, North Pacific, and Antarctic Oceans have manifested positive titers or have had Brucella isolated from their tissues. In southern New England, 4 of 90 stranded phocid seals showed positive titers to Brucella (Maratea, pers. comm.). Recently, Brucella was isolated from tissues of two of six stranded southern New England phocid seals sampled at necropsy. Unlike previously reported cases, the biochemical characteristics of these isolates differed significantly from isolates from Pacific phocids (Maratea, pers. comm.). Histopathological analysis of tissues from a Pacific harbor seal from which a Brucella spp. was isolated showed the presence of small coccobacilli, consistent with Brucella spp., within and attached to the lining of the uterus of lungworms (Parafilaroides spp.). These coccobacilli showed positive immunolabeling with antibody to B. abortus. The authors of the report suggested that exposure to infected Parafilaroides spp. might be one source of Brucella infections in harbor seals (Garner et al., 1997).
Vibriosis Vibrios are Gram-negative bacteria usually found in marine and brackish water environments. The most common form of Vibrio disease in marine mammals involves the contamination of wounds, but deaths, presumably due to a Vibrio septicemia, have been reported (Tangredi and Medway, 1980; Martineau et al., 1988). In human medicine, experts stress the need to utilize TCBS agar in Vibrio isolation attempts. A similar recommendation can be made for attempted recoveries from marine animals. Increasing resistance of a number of Vibrio spp. isolates to many of the commonly employed antibiotics has been noted. Early treatment with aminoglycosides, quinolones, or thirdgeneration cephalosporins may be necessary to prevent development of fatal septicemias (Buck et al., 1984; Greco et al., 1986; Fujioka et al., 1988).
0839_frame_C16 Page 315 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
315
Cetaceans
Disease in cetaceans caused by bacteria of the genus Vibrio has been reported on several occasions. Vibrio spp. are regularly isolated from the blowhole and anus of healthy cetaceans, so care must be exercised in the interpretation of such isolations. Commonly isolated species include the following: V. alginolyticus, V. damsela, V. fluvialis, V. parahemolyticus. Several other species have been less frequently isolated, including V. cholerae (Tangredi and Medway, 1980; Dailey, 1985; Greco et al., 1985; Buck and Spotte, 1986b; Fujioka et al., 1988; Martineau et al., 1988; DeGuise et al., 1995b; Parsons and Jefferson, 2000). Pinnipeds
Reports of Vibrio spp. isolations from pinnipeds are scanty (Buck and Spotte, 1986b). This could be due to resistance in this group to Vibrio-caused disease, failure to employ appropriate culture media or conditions in isolation attempts, or the fact that many of the institutions that exhibit pinnipeds maintain them in fresh water where vibrios would be less likely to flourish. Vedros (pers. comm.) regularly found antibodies to V. alginolyticus in many pinniped species, but never isolated a Vibrio from a pinniped. Sugita et al. (1996) recovered Vibrio spp. from only two of nine California sea lions (Zalophus californianus) studied. Visser et al. (1999) reported the isolation of a non-01 V. cholerae from the stool of a “young seal” with diarrhea.
Pasteurellosis Pasteurellosis most often manifests as an acute or peracute septicemia. Death often occurs either without obvious clinical signs or only a few hours after the development of anorexia or other behavioral signs, such as lethargy, decreased swimming, or failure to interact normally with pool mates. Sweeney (1978) reported hemorrhage, enteritis, and necrotic peritonitis; however, the majority of cases this author (J.L.D.) has seen (14) have had very few gross lesions, which is consistent with their peracute nature. Lesions when present included areas of fat necrosis in the blubber near the cervical esophagus, cervical swelling, epicardial and pericardial petechiation, and heavy, wet lungs. Histopathological findings include splenitis, hepatitis, interstitial and bronchial pneumonia, myocarditis, and nephritis. Cetaceans
Pasteurella multocida has been cultured from cetaceans at Mystic Aquarium on only a few occasions, with most of these isolations as incidental findings not associated with disease. A previously stranded bottlenose dolphin, which had been successfully rehabilitated over a period of several months, died peracutely while showing no signs of abnormal behavior. Routine blood samples collected the day prior to death showed a marked neutrophilia, a lymphopenia, and an eosinopenia. The necropsy findings were consistent with bacteremia/septicemia, and included diffusely edematous lungs and a diffuse acute lymphadenitis. Pasteurella multocida was isolated from several organs. Sweeney (1978) mentioned a P. multocida-caused enteritis, leading to death due to bacteremia and intestinal hemorrhage. He suggested a nearby bird rookery as the source of the organism. He also mentioned an outbreak of P. hemolytica infection in a dolphin colony that was effectively controlled with chloramphenicol after one animal had died with hemorrhagic tracheitis. Medway and Schryver (1973) recovered Pasteurella spp. from the lungs of a bottlenose dolphin with acute hemorrhagic bronchopneumonia. Pinnipeds
Pasteurellosis has been diagnosed in both phocids and otariids, and in three of six pinniped species maintained at Mystic Aquarium, suggesting the possibility that all pinniped species are susceptible to P. multocida septicemia. At Mystic Aquarium, most cases occurred in the fall and
0839_frame_C16 Page 316 Tuesday, May 22, 2001 11:12 AM
316
CRC Handbook of Marine Mammal Medicine
late summer, and were thought to be associated with the presence of large numbers of migrating waterfowl in a nearby lake. No cases developed during an extended period of time when the pinniped population was treated prophylactically with low levels of oral tetracycline (250 mg per animal BID PO). A polysaccharide vaccine developed from a necropsy isolate at Mystic appears to have provided long-term immunity after administration of appropriate primary and booster vaccinations (Vedros, 1982). Since the initiation of vaccination of the Mystic pinniped population with the Pasteurella vaccine there has been only a single Pasteurellaassociated death (an aged, female northern fur seal). Prior to this, losses averaged two to three animals per year. Earlier use of commercial P. multocida vaccines developed for use in cattle failed to provide any protection. Sensitivity tests and the success with the use of prophylactic tetracycline show that P. multocida is easily controlled with antibiotics. Unfortunately, the peracute nature of the disease has thus far resulted in a situation where treatment has been of no value once the disease is clinically obvious. Kennedy-Stoskopf et al. (1986) discussed a P. multocida-associated pericarditis and septicemia in a California sea lion. They speculated that the Pasteurella may require the presence of a concurrent immunosuppressive virus to cause clinical disease.
Erysipelothrix Erysipelothrix rhusiopathiae is a Gram-positive or Gram-variable (Patterson, 2000) bacillus most commonly associated with disease in swine and turkeys. Marine mammals are presumed to contract the disease by ingesting fish contaminated with the organism (Geraci et al., 1966). The pathogen has been isolated from freshwater and marine fishes and marine crabs (Lauckner, 1985). A recent survey of Australasian seafood revealed the presence of the pathogen in 19 species of marine fish and invertebrates, and repeated isolations have been made from several species of food fish routinely offered to marine mammals (Harris, pers. comm.). The potential for transmission of Erysipelothrix pathogens to humans should not be underestimated (see Chapter 34, Public Health). Diagnosis is based on clinical signs, isolation of the organism, or serological means (Jones et al., 1995; 1996; Patterson et al., 1997). Molecular techniques developed for diagnostic use in humans and food animals (Makino et al., 1994) will permit direct, rapid detection of Erysipelothrix DNA in infected marine mammals. The use of ionizing irradiation, such as the γ-irradiation emitted by cobalt-60, to decrease the pathogen load of food fish has been suggested as one means of reducing potential exposure to Erysipelothrix and other significant food-borne pathogens (Kilgen, 2000). Although this may provide an interim solution, the logistical and fiscal impacts associated with employment of this technology limit its immediate usefulness and provide an impetus for the development of a safe and efficacious Erysipelothrix vaccine. A recent workshop at the Shedd Aquarium in Chicago, Illinois brought together researchers, veterinarians, and marine mammal care staff members from facilities in North America, Europe, and Asia. The group conducted an intensive review of the state of knowledge as it relates to the epidemiology, diagnosis, and prevention of Erysipelothrix infections in cetaceans. Many of the findings of this group are mentioned throughout the following discussion of this disease in cetaceans. Cetaceans
In cetaceans (unlike the situation in pinnipeds), disease caused by Erysipelothrix has been frequently diagnosed in many species (Siebold and Neal, 1956; Geraci et al., 1966; Medway and Schryver, 1973; Sweeney and Ridgway, 1975; Howard et al., 1983; Buck and Spotte, 1986a; Bossart and Eimstad, 1988; Kinsel et al., 1997). The condition is not unique to captive cetaceans, as stranded cetaceans have been diagnosed with the disease (Young et al., 1997). Geraci and co-workers (1966) discussed the difficulties in ante-mortem diagnosis. Nonspecific clinical signs,
0839_frame_C16 Page 317 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
317
such as weakness and anorexia, were of little diagnostic value. Only an increase in circulating neutrophils hinted at severe infection. Two variants of the disease have been reported, an acute septicemic form and a dermatological form. The dermatological form in bottlenose dolphins is characterized by gray rhomboid dermal plaques over the entire body trunk (Sweeney, 1978). In belugas (Delphinapterus leucas), the dermatological variant may take the form of indurated lesions that may eventually scar over after appropriate treatment, and then may manifest as a differently pigmented area of skin. In the more common cutaneous form of the disease, the animal develops a leukocytosis, and dermal plaques appear shortly after anorexia is seen. Prompt initiation of treatment with appropriate antibiotics is usually curative. Failure to recognize and treat the condition may result in death, however. The septicemic variant quite often results in death, since early clinical signs may be absent or, if present, are nonspecific. Anorexia, lethargy, and an initial leukocytosis may be followed by a severe leukopenia just prior to death. A high index of suspicion for Erysipelothrix infection is always necessary, and usually treatment must be begun before confirmation of one’s diagnosis can be made. Indeed, many diagnoses of this infection are made retrospectively (Calle et al., 1993). Necropsy findings in cetaceans with the septicemic form of the disease have included serosanguinous ascitic fluid, multifocal intestinal petechial and ecchymotic hemorrhage, sloughing skin, swollen lymph nodes, and splenomegaly. Neuraminidase, a potent enzyme that removes neuraminic acid from cell membranes, is thought to be a major factor in mediating the characteristic vascular damage associated with E. rhusiopathiae. Isolation of the pathogen can often be made from all organ systems. One formerly stranded, unvaccinated, Atlantic white-beaked dolphin (Lagenorhynchus albirostris) died within a few hours of eating normally. Other cetaceans maintained in the same water system and fed the same food fish by the same handlers showed no evidence of the septicemic form of the disease. However, one of several belugas maintained in this system later developed chronic ulcerative suppurative multifocal dermatitis due to Erysipelothrix spp. Another beluga with grossly similar dermatological lesions showed a return to normal skin, although it was differently pigmented, after a 30-day course of oral cephalosporins. Not all cases of Erysipelothrix-induced septicemia end fatally. Seroconversion after an acute onset of illness gave presumptive evidence of active infection in an adult beluga that recovered uneventfully after treatment with ciprofloxacin (Calle et al., 1993). A precipitous change in behavior and appetite in an adult male beluga at Mystic was followed by a similar recovery and increase in titer to the organism after prompt initiation of treatment with ceftiofur. A year earlier, another adult beluga at the same institution died within minutes of a bloodsampling procedure initiated after the animal’s appetite had decreased during the previous 12 hours. A pronounced leukopenia was noted and E. rhusiopathiae was cultured from multiple sites in the body. Only about one third of the 75 institutions replying to a recent survey (Lacave, pers. comm.) vaccinate against Erysipelothrix infections. Most institutions in North America have discontinued the practice because of side effects associated with the administration of commercially available bacterins. Reports of a live culture vaccine producing the disease prompted the U.S. Navy to discontinue the use of such vaccines. On several occasions, the authors have noted swelling at the site of killed bacterin injections administered in an aseptic manner. Two cases of anaphylaxis in belugas after booster vaccinations with killed vaccine have been observed by one of the authors (J.L.D.). Intravenous administration of epinephrine reversed the condition. Recognizing that there was a distinct likelihood that continued vaccinations might be followed by additional anaphylactic episodes, the decision to discontinue vaccinating against the disease with that particular bacterin came without hesitation. Sweeney (1978) also reported anaphylactic reactions in animals receiving a second or later exposure to the bacterin.
0839_frame_C16 Page 318 Tuesday, May 22, 2001 11:12 AM
318
CRC Handbook of Marine Mammal Medicine
Even when vaccination was commonly practiced, considerable controversy existed regarding the relative efficacy and appropriateness of various vaccination regimens to prevent infection with Erysipelothrix. Gilmartin and co-workers (1971) recommended that bottlenose dolphins be vaccinated initially with a killed bacterin. At 7 weeks after the primary vaccination, the animals received booster injections of a live culture vaccine. Regular revaccination with live culture vaccine at 6-month intervals produced substantial antibody levels. Suer et al. (1988) reported that approximately 10% of the wild bottlenose dolphins and false killer whales (Pseudorca crassidens) they sampled had received sufficient challenge to induce significant antibody levels to E. rhusiopathiae. Approximately 25% of the captive dolphins sampled in their study, all of which had received many vaccinations, showed negligible antibody titers. Hermans et al. (1997) demonstrated the efficacy of a commercial vaccine in mice against cetacean-origin Erysipelothrix isolates. Lacave and Cox (pers. comm.) reported that 49 of 58 cases of the disease, diagnosed in a population of 1373 captive cetaceans in the period 1989 through 2000, occurred in nonvaccinated animals. No cases were reported in animals vaccinated on a regular basis. These data compellingly manifest the benefits of the vaccination protocol they employ. Lacave and Cox (pers. comm.) outline a vaccination program in young bottlenose dolphins, which uses a European-developed killed Erysipelothrix vaccine perfected for use in swine. They have not experienced any cases of significant side effects such as anaphylaxis or the development of the disease in any of the vaccinated animals. The titers produced are variable, but in most instances show an early increase in IgM followed by a later increase in IgG. As yet, there are no data regarding what constitutes a protective titer. Lacave and Cox have not utilized this vaccination regimen on belugas, so it is unknown whether this protocol eliminates the anaphylaxis problem sometimes seen in this species following vaccination. Progress in the development of a safe and broadly efficacious vaccine against Erysipelothrix infections in marine mammals has lagged behind the advances in developing vaccines for economically important terrestrial species (Bricker et al., 2000). Recent developments in vaccine technology have shown promise that safer, more efficacious vaccines may soon become available and, one hopes, these improved vaccines can be adapted for use in marine mammals. Development of a DNA vaccine employing the gene for the production of the 64-kDa antigen of E. rhusiopathiae might eliminate many of the adverse effects seen in older, whole-cell bacterins. Administration of monoclonal antibodies, which have shown efficacy in protecting other species against challenge with E. rhusiopathiae, could become an important adjunct in early treatment (Ziesenis et al., 1992), but, at present, prompt initiation of antibiotic therapy remains the treatment of choice. Pinnipeds
Lauckner (1985) states that Erysipelothrix infections are rarely encountered in pinnipeds. Nevertheless, the organism can infect pinnipeds. A report of isolation of E. insidiosa from multiple organs of two dead fur seals probably represents the septicemic form of the disease (Benkovsky and Golovina, 1971). Suer and Vedros (1988) isolated E. rhusiopathiae from the teeth and/or gums of two elephant seals (Mirounga angustirostris) and two northern fur seals (Callorhinus ursinus). They also detail modifications to culturing techniques that increase the likelihood of a successful isolation of the organism. Suer and Vedros (1988) also documented the presence of significant antibody levels to E. rhusiopathiae in many adults of the pinniped species they sampled; northern fur seal and stranded Pacific harbor seal pups had no significant antibodies. At Mystic Aquarium, one case of the septicemic form of the disease was seen in a young hooded seal 2 days after it arrived from Florida, where it had beach-stranded 2 months earlier. Upon clinical examination, the animal exhibited shallow, rapid respiration, cyanosis, unilateral
0839_frame_C16 Page 319 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
319
crackling rales, diarrhea, dehydration, and a marked leukopenia. Necropsy findings included pulmonary hemorrhage, lymphoid depletion, and diffuse multifocal congestion and fibrosis. Erysipelothrix rhusiopathiae was isolated from multiple organs. Sweeney (1974) commented that infection with E. rhusiopathiae was not an important clinical problem in captive pinnipeds and that vaccination with a commercial bacterin, while not harmful, did not appear necessary. Since the authors have not documented any cases in collection pinnipeds in the last 25 years, and have never vaccinated pinnipeds against the disease and feed the same food fish as fed to their cetaceans, they concur with his observation.
Mycobacterial Disease Scanty, anecdotal, and incomplete reports of mycobacterial infections in marine mammals have been in the literature since the early 1900s (Blair, 1913; Ehlers, 1965). Beginning in the 1970s and throughout the 1980s, more complete reports of mycobacterial disease in captive pinnipeds and sirenians began to surface (Boever et al., 1976; Morales et al., 1985; Gutter et al., 1987; Stoskopf et al., 1987; Bernadelli et al., 1990; Wells et al., 1990; Castro Ramos et al., 1998). All of these cases involved atypical, nontuberculous mycobacterial species. In the early 1990s, tuberculous mycobacterial infections were diagnosed in a number of Southern Hemisphere captive and wild pinniped populations (Forshaw and Phelps 1991; Cousins et al., 1993; Gales, 1993; Thorel and Moutou, 1994; Hunter et al., 1998; Thorel et al., 1998). The source of the pathogen in the tuberculosis cases, reported in both wild and captive seals (and in a seal trainer), remains a matter of conjecture. None of the many hypotheses that have been postulated has been proved. These have included infection passed from staff or visitors to captive seals, or infection of wild and captive seals by infected cattle or other zoological park collection species, or transmission via ingestion of the corpse(s) of infected humans buried at sea, or the disease may have been present in wild seal populations for thousands of years. At Mystic, the authors have repeatedly been able to isolate atypical mycobacteria from sediments in fish and invertebrate holding tanks, but have never isolated tuberculous mycobacterial species. Cetaceans
Bernadelli et al. (1990) employed a number of techniques in attempting to diagnose mycobacteriosis in marine mammals and birds. Using the intradermal tuberculin test, ELISA tests, and isolation attempts, they were unable to document infection or exposure in any of the dolphins and orcas they sampled. At Mystic Aquarium, the authors isolated Mycobacterium marinum, in very low numbers, from deep in a skin lesion of a beluga diagnosed with the cutaneous form of the disease. The animal was treated with minocycline and cephalexin for ∼30 days and the lesion healed well. Nearly 12 years later this animal was diagnosed with severe deep pyogranulomatous panniculitis from which M. marinum was readily isolated. The animal died of other causes before specific antimycobacterial therapy was instituted (Dunn et al., 2000). Pinnipeds
Cases of infection in pinnipeds with atypical mycobacterial species have been reported in the literature for many years. Mycobacterium fortuitum, M. marinum, M. smegmatis, M. chelonei (M. chelonae), and M. chitae have all been implicated. Bernadelli et al. (1990) documented atypical mycobacteriosis (M. chelonae and M. fortuitum) in captive fur seals. Wells et al. (1990) describe a case of cutaneous mycobacteriosis in a captive harbor seal from which both M. chitae and M. fortuitum were isolated. Thorel and Moutou (1994) suggest that captivity in zoos increases the risk of development of mycobacterial infection. They postulate that this occurs because many
0839_frame_C16 Page 320 Tuesday, May 22, 2001 11:12 AM
320
CRC Handbook of Marine Mammal Medicine
species that are not normally found together in nature are held in comparatively small areas. Their review documented mycobacterial infection in otariids, phocids, and sirenians along with a host of nonmarine mammal species. Their caution about the possibility of transmitting mycobacterial infections via the reintroduction of formerly captive endangered species holds equally true for the reintroduction of rehabilitated stranded marine mammals. Needham (1992) discussed the technique of intradermal testing (IDT) for tuberculosis and compared the sensitivity and specificity of IDT and ELISA in diagnosing tuberculosis in pinnipeds. Tuberculosis was diagnosed in pinnipeds for the first time in 1986, in a western Australian marine park. Australian sea lions (Neophoca cinerea) and New Zealand fur seals (Arctocephalus forsteri), the species involved, were diagnosed at necropsy, or by intradermal tuberculin tests using bovine-purified protein derivative (PPD), or by ELISA (Cousins, 1987). About 4 years after this outbreak, the first bacteriologically confirmed diagnoses of tuberculosis in wild pinnipeds of the same species were made (also in western Australia). While these cases initially appeared to be caused by M. bovis, later PCR and restriction endonuclease analysis of samples from the wild seals identified those strains as a “unique cluster in the M. tuberculosis complex” (Cousins et al., 1993). Spread of the disease was thought to be via inhalation, because the lung was the focus of infection in both the wild and captive seal cases. The gregarious nature of many pinniped species would likely predispose to transmission via this route. All three cases diagnosed in wild seals were in older males. This suggests that the male seals’ occasional use of mainland beaches might put them at greater risk to the disease than females, which tend to congregate on offshore islands. Most recently, M. tuberculosis has been diagnosed in wild subantarctic fur seals (Arctocephalus tropicalis). As in the other four species of wild, Southern Hemisphere pinnipeds in which this diagnosis has been made, the principal lesions involve pulmonary granulomata with central areas of caseous necrosis. The authors (Bastida et al., 1999) speculate that the subantarctic fur seal represents the common link in the spread of the disease to other Southern Hemisphere pinnipeds, since it is found in association with all the other pinniped species in which tuberculous mycobacteriosis has been diagnosed.
Leptospirosis Leptospirosis is a serious, geographically widespread disease affecting the liver, kidney, and reproductive systems of many terrestrial and aquatic vertebrates. Most of the earliest reports of leptospirosis in marine mammals cite Leptospira pomona (L. interrogans, serovar pomona), or an organism from which it is serologically indistinguishable, as the etiological agent. More recent case reports have implicated L. interrogans, serovar grippotyphosa. No definitive diagnoses of leptospirosis have yet been made in cetaceans. The disease is most commonly seen in California sea lions and northern fur seals. Northern elephant seals have shown elevated titers, and both wild and rehabilitated harbor seals maintained in close proximity to both California sea lions and northern elephant seals have been diagnosed with the disease (Stamper et al., 1998; Stevens et al., 1999). Pinnipeds
The disease was first recognized in California sea lions along the California and Oregon coasts in 1970 (Schoenwald et al., 1971; Vedros et al., 1971). During this epizootic, in which hundreds of animals were involved, the number of stranded sea lions was approximately four times higher than normally reported. Only adult and subadult males were affected. A later report (Smith et al., 1974b) detected the presence of antibodies to L. pomona in four of ten aborting female California sea lions, and it was speculated that the disease caused reproductive failure in adults
0839_frame_C16 Page 321 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
321
and multiple hemorrhagic syndromes in fetuses and neonates. Additional epizootics continue to occur (Gulland et al., 1996). Serological evidence in the northern fur seal suggests that the exposure to L. pomona occurs at sea; however, the isolation of L. pomona from the placentas of California sea lions on San Miguel Island, California indicates that rookery exposure is also possible (Smith et al., 1974b). Dierauf et al. (1985) suggest the contamination of fresh water by domestic animals and wild rodents as possible sources of the spread of the pathogen into the sea lion population. Clinical signs of the disease include depression, extreme thirst, reluctance to use the rear limbs, fever, polydipsia, anorexia, and, on occasion, icterus, muscle tremors, and vomiting. Some authors report a leukocytosis along with elevated serum creatinine, blood urea nitrogen (BUN), and globulin (Dierauf et al., 1985; Stamper et al., 1998), whereas others have found no significant hematological changes (Gulland et al., 1996). In adults, gross necropsy findings include kidneys that are markedly swollen and hard, with a loss of differentiation between the medullae and cortices, gallbladders containing thick, black bile, and thick, pale yellow pericardial fluid. A third of the affected animals showed swollen, friable livers (Dierauf et al., 1985; Gulland et al., 1996). Histopathological changes have included diffuse interstitial nephritis and glomerulonephritis. Large numbers of the spirochetes have been seen in some silver-stained kidney sections. In fur seal pups at necropsy, free blood was present in the abdominal cavity, and subcapsular hemorrhage of the kidneys and liver were prominent. The livers were friable and may have been the source of blood present in the abdominal cavity. Ocular anterior chamber hemorrhage was a common finding, as was subperiosteal hemorrhage (Smith et al., 1977). Vedros et al. (1971) mentioned the successful treatment of one leptospirosis-affected California sea lion with penicillin, streptomycin, and vitamin B complex, as well as free access to water. Dierauf and colleagues (1985) reported the successful treatment and release of 66 California sea lions; these animals were treated with either tetracycline (22 mg/kg TID PO) or potassium penicillin G (44,000 U/kg BID PO or IM) for 10 to 14 days. Stamper et al. (1998) treated a harbor seal successfully with tetracycline, oral phosphorus binders, and oral electrolyte supplementation. Smith et al. (1977) speculated on the use of a vaccine to protect fur seals, and at Mystic Aquarium one of the authors (J.L.D.) vaccinates fur seals twice each year with a five-serovar commercial livestock vaccine (Lepto-5, Biocor Animal Health, Omaha, NE). There is considerable variability in the titer levels achieved against the different serovars, but the incidence of leptospirosis cases has decreased since initiation of the vaccination program.
Nocardia Nocardia spp. are classified as aerobic actinomycetes in the order Actimomycetales. Pathogenic Nocardia spp. include N. asteroides (and its several serotypes), N. brasiliensis, N. otitidiscaviarum, and N. transvalensis. Nocardia is ubiquitous in the environment, found in soil, vegetable matter, and water (Williams et al., 1983). Infections with nocardioforms result in a wide variety of clinical manifestations depending on the organ system(s) involved. Granulomatous inflammatory reactions, which may be acute or chronic and which often progress to abscess formation, characterize typical nocardial infections (McNeil and Brown, 1994). At present, diagnosis requires identification of the organism via cytological or histological preparations, in conjunction with isolation of the organism from infected tissue. Successful treatment relies on early diagnosis and administration of appropriate antibiotics. Although it provides a definitive diagnosis, culture from blood or infected tissue is often unrewarding. It sometimes requires weeks for a successful isolation, while an infectious
0839_frame_C16 Page 322 Tuesday, May 22, 2001 11:12 AM
322
CRC Handbook of Marine Mammal Medicine
process is continuing its destructive activities. However, the presence of acid-fast (or partially acid-fast), branched filaments in accessible lesions, together with clinical signs, may provide a presumptive diagnosis pending isolation of the pathogen. A serology test, based on both ELISA and Western immunoblot reactions to specific, key nocardial antigens, has been developed. The specificity to the diagnostic antigens, which are found only during infection, is high, with crossreactions only to some mycobacteria. The test is not as yet commercially available, and is only conducted at the University of California at Davis. A positive antigen isolation indicates a clinical or subclinical Nocardia spp. infection. Unfortunately, not all species of Nocardia produce these antigens, so a nonreacting Nocardia spp. infection cannot be ruled out (Beaman, pers. comm.). There are at least six basic forms of clinical Nocardia spp. infections identified in humans. These include pulmonary, systemic, extrapulmonary, cutaneous, subcutaneous, and lymphocutaneous nocardiosis. Pulmonary nocardiosis exhibits no specific clinical signs. The disease can be acute or chronic, and may result in pneumonia, abscess formation, or both. The infections usually involve a suppurative response and may be granulomatous. In humans, the condition is often misdiagnosed as pyogenic, or of mycotic origin (Beaman and Beaman, 1994). Systemic nocardiosis can result either from traumatic lesions or when a localized, often pulmonary, infection erodes vessel walls and allows the pathogen to be broadly disseminated. Unlike pulmonary nocardiosis, disseminated nocardiosis tends to be relentlessly progressive and not readily treated with chemotherapeutics. Extrapulmonary nocardiosis includes infections isolated to a site other than pulmonary or cutaneous. The most common extrapulmonary sites involve the brain or skeleton. Cutaneous, subcutaneous, and lymphocutaneous nocardial infections are afforded their own category because of the frequency of their involvement with these sites. These types of infection often occur following traumatic injury, with the infection later becoming circumscribed and forming an abscess. Usually these infections are self-limiting, but on occasion they progress. All the Nocardia spp. can cause cutaneous lesions, but N. brasiliensis most frequently results in a progressive disease (Beaman and Beaman, 1994). Cetaceans
Nocardiosis was first reported in cetaceans by Pier et al. (1970; 1981). They reported three cases at Sea Life Park, Hawaii, that involved a captive pilot whale (Globiocephala scammoni) and two bottlenose dolphins (Tursiops truncatus gilli). They later confirmed another case in a captive harbor porpoise. Since that time, cases have been observed in bottlenose dolphins (Jasmin et al., 1972; Sweeney et al., 1976), a false killer whale, a spinner dolphin (Stenella longirostris), and a killer whale (Sweeney et al., 1976). No other cases were reported until the 1990s, when six additional cases of nocardiosis were diagnosed in belugas and killer whales (Table 1). Pulmonary or extrapulmonary nocardiosis were the forms of the disease most often noted. Because Nocardia spp. are ubiquitous in soil, organic matter, and water, organisms may easily become airborne on dust particles and inhaled or aspirated. Cetaceans appear to be particularly vulnerable to pneumonia; thus any environmental activity that could increase exposure to soil-borne particulate matter should be considered as possible sources of infection. Three of the recent six cases were hypothesized to have been initiated at sites within close geographical proximity of each other, and shortly after a violent, dust-spreading weather pattern. Four cases occurred in animals less than 21/2 years of age, including two neonates less than 40 days old and one animal 132 days old. All young animals were initially examined because of concerns relating to abnormal nursing patterns. The first animal, a 40-day-old male beluga, presented with inappetence, lethargy, and an increased respiratory rate. At 5 days postpartum, the calf was treated with antibiotics, antifungals, and purified beluga IgG after it exhibited inappetence and inadequate weight gain. The calf ’s dam was found to have mastitis. Once the mastitis was successfully treated, the calf ’s nursing patterns returned to normal. A day 40 blood sample and physical examination showed
Adult (F) Subadult (M) Adult (F)
New Zealand New Zealand Hong Kong
Adult (F)
New York
Adult (?)
Hawaii
D. leucus
Adult (M)
Hawaii
Infant (F) Immature (M) Immature (M) Adult (M)
Mature (F) ? Adult (M)
Hawaii Florida Hawaii
Texas Texas Florida Texas
Infant (F)
Hawaii
Infant (M)
Mature (M)
Hawaii
Texas
Immature (F)
Age (Sex)
U.S.A.
Region
Delphinapterus leucus (beluga) O. orca D. leucus O. orca D. leucus
Phocoena phocoena (harbor porpoise) Tursiops truncatus gilli (bottlenose dolphin) Globicephala scammoni (Scammons pilot whale) T. t. gilli T. truncatus Orcinus orca (killer whale) Pseudorca crassidens (false killer whale) Stenella longirostris (spinner dolphin) T. truncatus Hydrurga leptonyx (leopard seal) T. t. gilli
Species Infected
TABLE 1 Nocardiosis in Marine Mammals
Pulmonary Extrapulmonary Pulmonary Subcutaneous
Pulmonary
?
? ?
?
?
Pulmonary Cutaneous ?
Pulmonary
Pulmonary
Pulmonary
First Site of Infection
3 days >1 year 2 years 1 year
19 days
?
? ?
?
?
7 days ? ?
30 days
?
?
Duration of Clinical Disease
N. asteroides, N. brasiliensis
N. asteroides Nocardia spp.
N. asteroides
Nocardia spp.
Nocardia spp. Nocardia spp.
N. asteroides
N. asteroides
N. caviae N. paraquayensis N. asteroides
N. asteroides
N. brasiliensis
Nocardia asteroides
Isolate
Calle, pers. comm.
Dalton and Robeck, 1995 Robeck et al., 1995 Walsh, pers. comm. Robeck and Dalton, 1995
Hammond (1975) in Sweeney et al., 1976 Robeck et al., 1994
Pier et al., 1970 Jasmin et al., 1972 Conklin et al. (1972) in Sweeney et al., 1976 Conklin et al., (1972) in Sweeney et al., 1976 Conklin et al. (1972) in Sweeney et al., 1976 Allen, 1974) in Sweeney et al., 1976 Allen (1974) in Sweeney et al., 1976
Pier et al., 1970
Pier et al., 1970
Geraci (1969) in Pier et al., 1970
Reference
0839_frame_C16 Page 323 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds 323
0839_frame_C16 Page 324 Tuesday, May 22, 2001 11:12 AM
324
CRC Handbook of Marine Mammal Medicine
that the calf had a severe degenerative left shift, inflammation, regenerative anemia, and pleural effusion. The thoracic effusion was drained and a Nocardia spp. was isolated. Despite treatment based on antibiotic sensitivity of the isolate, the animal died on day 59. Necropsy revealed a primary pneumonic nocardiosis, which led to pleuritis and later spread to other organs (Robeck et al., 1994). The second neonatal case involved a 35-day-old female killer whale calf that also presented with lethargy, inappetence, and an elevated respiratory rate. The animal’s hematology parameters showed a left shift, and an elevated erythrocyte sedimentation rate. The animal was administered purified killer whale IgG, amikacin, and itraconazole. The animal died 3 days later, and was diagnosed with multifocal pyogranulomatous pneumonia. In addition to numerous other bacteria, N. asteroides was cultured from the granulomatous lesions. Similar lesions were found in the lungs, pleura, thyroids, spleen, adrenal, heart, mediastinal lymph nodes (Dalton and Robeck, 1995). A third animal was a 132-day-old male beluga calf that presented with an elevated total white blood count and increased erythrocyte sedimentation rate. The animal was administered oral antibiotics and antifungal agents. The animal initially responded to treatment, but required the intermittent administration of antibiotics over the following 8 months. After 10 months of this treatment, the animal began to exhibit an inability or unwillingness to use its flukes. After another year, most directed motion was accomplished only via pectoral movements, and the animal was euthanized. Necropsy revealed a chronic suppurative vertebral osteomyelitis from T10 to L1, with areas of liquefactive necrosis of the adjacent spinal cord. Nocardia spp. were isolated from the lesions (Robeck et al., 1995). The fourth animal was a 27-month-old female killer whale that presented with an acuteonset depression and a body list to the right. As with four of the other cases, clinical signs were observed soon after an increase in dust particles in the air caused by nearby construction. The animal died after 2 years of treatment with a variety of antibiotics. During this time, the only improvement was observed during short courses of amikacin. Necropsy revealed a 10-cm abscess in the left cerebrum; the left humoral scapula joint was ankylosed and swollen to approximately 2.5 times normal diameter. Cultures from both sites grew N. otitidiscaviarum. The fifth animal diagnosed with nocardia infection was an 18-year-old male beluga that presented with a well-circumscribed mass. Hematologically, the animal had a slightly elevated white blood cell count, with a neutrophilia and monocytosis, and decreasing alkaline phosphatase, serum iron, and mean corpuscular volume (MCV). A wedge biopsy of the mass revealed an abscess that was cultured, drained, and flushed with chlorhexidine (Novasan®, Aveco Co., Inc., Fort Dodge, IA). Cultures of the mass revealed a mixed infection with N. brasiliensis and N. asteroides. The presence of N. brasiliensis is usually associated with progressive disease and suggests a poor prognosis. The lesions recurred several times over the next few months, despite appropriate antibiotic therapy. After 1 year, the animal was placed on a 6-month course of intramuscular amikacin (Amiglyde-V, Fort Dodge Lab, Inc., Fort Dodge, IA; 15 mg/kg SID). The lesions were aspirated, flushed, and injected directly with amikacin. All lesions appeared to resolve after 20 days, and isolation attempts either were negative or exhibited only slight growth. A lesion reappeared 30 days after the end of the amikacin treatment. The conclusion was that the lipophilic nature of the organism compounded the difficulty of treating the abscess in the relatively hematogeneously isolated blubber. The lack of response to chemotherapeutics and the potential for transmission of the organism to other animals, as well as the long-term prognosis, aided in the decision to euthanize the animal (Robeck and Dalton, 1995). Diagnosis of nocardial infections depends on isolation from infected tissue (Beaman and Beaman, 1994). Presumptive diagnosis pending cultures can be made when Gram-positive, filamentous organisms are observed on a cytological specimen. Serological tests are in development, but thus far are not totally rewarding. Treatment for these infections is difficult and
0839_frame_C16 Page 325 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
325
depends on how early the organism is detected and the character of the infection. In the authors’ hands, the best response to antibiotic therapy was obtained with amikacin. Single daily dosing at 15 mg/kg was used for up to 6 months in the above-mentioned belugas, in an Asian short clawed otter (Aonyx cinerea) with suspected nocardiosis (Robeck et al., 1996), and in a killer whale, all without development of renal problems. In humans, current recommendations for antibiotic therapy include imipenen, amikacin, or cefotaxime, or a combination (Beaman et al., 1992). However, as with the previously mentioned belugas, many cases have been reported that describe reoccurrence of Nocardia spp. disease, despite repeated and apparent successful chemotherapeutic treatment (Beaman and Beaman, 1994). When trying to identify the source of infection, one needs to recall that the appearance of clinical signs may be significantly delayed from the time of initial exposure, since Nocardia spp. has a propensity to exhibit a prolonged latency period. The large number of cases reported in marine mammals demonstrates their susceptibility to this organism. Efforts should be made to determine if animal habitats are located in endemic or high-risk areas for human Nocardia infections. Practitioners in these areas should be cognizant that certain weather patterns (high wind, seasonal variations in rainfall), combined or separate from local construction or dust-causing activity, may increase the risk of infection. Pinnipeds
One case of Nocardia spp. infection in a pinniped has been reported. This was in a juvenile male leopard seal (Hydrurga leptonyx) that had been held in captivity for 3 weeks (Sweeney et al., 1976). Despite the paucity of reported cases, there is every reason to believe that infections with these organisms readily occur in pinnipeds, and those involved with their care should always suspect Nocardia spp. in any granulomatous infectious process.
Miscellaneous Bacterial Disease Respiratory Disease The respiratory system continues to rank high on the list of organ systems from which pathogenic bacteria are isolated. In wild and stranded marine mammals, bacterial respiratory disease is frequently a sequel to heavy parasitism. Stranded harbor seals have a high prevalence of bronchopneumonia, often secondary to heavy infestation with Otostrongylus circumlitus. Sweeney and Gilmartin (1974) diagnosed pneumonia/pneumonitis in nearly 80% of the California sea lions they examined, and in Scotland, gray seal pups succumbed to bacterial pneumonia at an alarming rate (Gallacher and Waters, 1964). Gerber et al. (1993) list pneumonias as second in their list of primary disease findings in stranded California sea lions. Captive pinnipeds have a lower prevalence of bacterial pneumonia, possibly because of prompt treatment for parasitism, thereby eliminating one significant predisposing factor. Howard et al. (1983) describe gross and histopathological lesions of bacterial bronchopneumonia in stranded pinnipeds. The principal gross pathological changes include the mottled white to mahogany red areas of wet, heavy lung tissue. Purulent material often exudes from bronchi and bronchioles. Histological changes vary with the stage of the infection. Early on, alveolar exudate is mainly composed of neutrophils and monocytes. Later, with an increase in consolidation, neutrophils decrease and macrophages predominate. Cetaceans, too, have a high prevalence of respiratory disease with a bacterial component. Howard et al. (1983) report pure bacterial pneumonia as the principal cause of death in cetaceans in Hawaiian waters. Parsons and Jefferson (2000) report that nearly a third of the finless porpoises (Neophocoena phocoenoides) they examined showed moderate to heavy lungworm infections, and they were able to isolate 15 species of bacteria from nine animals in their
0839_frame_C16 Page 326 Tuesday, May 22, 2001 11:12 AM
326
CRC Handbook of Marine Mammal Medicine
study. Kinoshita et al. (1994) reported multiple mortalities in bottlenose dolphins associated with Staphylococcus aureus pneumonia. Studies on disease in captive cetaceans also found bacterial pneumonia to be a common cause of death (Medway and Schryver, 1973; Sweeney and Ridgway, 1975; Buck and Spotte, 1986a). A significant proportion of cetaceans diagnosed with respiratory tract problems exhibit pulmonary abscesses. Staphylococcus aureus and Pseudomonas aeruginosa have been prominent pathogenic isolates. Less frequent, but common, isolates tend to be Gram-negative organisms. In pinnipeds, bacterial respiratory disease is often accompanied by coughing, whereas in cetaceans most coughing occurs in animals with upper respiratory disease. An increased respiratory rate is often associated with respiratory disease in pinnipeds, but in dolphins is frequently not readily observed until a significant portion of the lung field is damaged. Unilateral pneumonia or large pulmonary abscesses in cetaceans sometimes result in an animal listing to the side of the damaged lung. This finding is not universal, as one cetacean has been observed with a large-volume unilateral pleural effusion and associated total lung collapse that showed no predisposition to list to the affected side. Other generalized signs are not helpful in localizing the problem. Nasal or blowhole swabs and exhalation plates will aid in identifying pathogens as will cytological analysis of sputum (see Chapter 20, Cytology). Bronchoalveolar lavage during bronchoscopic procedures has also proved useful in cetaceans (Harvell et al., 1996; Hawkins et al., 1996). For treatment of bacterial respiratory disease, antibiotic selection should follow routine culture and sensitivity. Treatment must often begin before results from these tests are available; thus, antibiotics with a broad spectrum and good activity against Gram-negative organisms are used initially. In pinnipeds, oxygen supplementation, bronchodilators, and mucolytic agents have proved useful in some bacterial respiratory diseases.
Dermatological Disease Marine mammals appear to be as susceptible to dermatological problems as terrestrial mammals, although most bacterial skin disease follows a primary viral, parasitic, or traumatic insult. Meyer (1997) suggested that the form of locomotion employed by harbor seals (and hence other phocids) on land might predispose them to chemical and mechanical damage to the skin of ventral surfaces. Meyer et al. (2000) reported the presence of significant amounts of saccharide residues in the sebaceous and tubular apocrine glands of the harbor seal. Upon release by microbial action, these saccharides, when found on the skin, may act to inhibit the adherence of microorganisms to the epidermal surface. Sweeney (1974) described a focal dermatitis in the California sea lion that responds to ampicillin, lincomycin, or erythromycin. He also mentioned dermal abscesses, a condition this author (J.L.D.) has seen in stranded young harbor seals. Rand (1979) described an epizootic in Galapagos sea lions (Z. californianus wollebaeki), which resulted in numerous deaths. Pseudomonas aeruginosa was the predominant organism recovered from the suppurative cutaneous nodules that were the hallmark of the disease. VanPelt and Dieterich (1973) described S. aureus abscesses in a young harbor seal. Howard et al. (1983) reported that β-hemolytic streptococci were the most common pathogens isolated from cutaneous abscesses in their study. Brown et al. (1994) speculated on a possible bacterial etiology of a case of orthokeratotic hyperkeratosis in a harbor seal. The cutaneous form of disease caused by Erysipelothrix spp. and the frequently seen contamination of surface wounds of cetaceans with Vibrio spp. have been previously discussed. Cusick and Bullock (1973) described ulcerative dermatitis associated with Aeromonas hydrophila in a bottlenose dolphin. Some of the histological changes in cetacean skin produced by bacterial infections have also been described by Howard et al. (1983). Mullan (1991) suggested that at least some of the bacterial species cultured from the skin of bowhead whales (Balaena
0839_frame_C16 Page 327 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
327
mysticetus) are pathogenic, and that Rhodococcus equi, Corynebacterium pseudotuberculosis, and Moraxella spp. had the greatest potential to be the etiological agents of bowhead whale skin lesions. Wilson et al. (1999) speculated on the impacts of natural and human-related factors on the epidermal health of wild bottlenose dolphins. Their study did not permit determination of the etiology of the skin lesions they observed affecting >60% of the individuals in their study. Many other reports of isolations from skin lesions exist. Most of these represent contamination of dermal wounds with bacterial organisms common to the animals’ environment. When sampling dermal lesions for bacterial isolation, it is often helpful to sample a nearby area of normal skin at the same time. This will act as a control for the normal flora found in the environment.
Urogenital Disease Other than leptospirosis, there have been few reports of bacterially induced disease of the urogenital system in marine mammals. At Mystic Aquarium, excluding leptospirosis cases, there have been three pinniped deaths associated with urogenital disease of bacterial origin in 25 years. These have included an adult male California sea lion with a mixed bilateral pyelonephritis subsequent to a bite on the penis, an aged northern fur seal with a mixed bacterial pyometra, and an adult Atlantic harbor seal with a postpartum Streptococcus spp. metritis and septicemia. The authors have also diagnosed a mixed bacterial pyometra in a Steller sea lion (Eumetopias jubatus) that responded well to drainage, antibiotics, and prostaglandin therapy. Howard et al. (1983) reported stranded California sea lions with both pyometra and metritis. One animal yielded a pure culture of Edwardsiella. A second animal had a mixed infection. Brown et al. (1960) reported renal abscesses in bottlenose dolphins. Sweeney and Ridgway (1975) reported that postfeeding diuresis tends to inhibit ascending urinary tract infections and that renal infections in cetaceans are rare and often associated with infection elsewhere in the animal.
Gastrointestinal Disease Except for clostridial enterotoxemia, primary gastrointestinal disease of bacterial origin is an uncommon problem in captive marine mammals. However, wild pinnipeds have frequently been diagnosed with bacterial gastroenteritis. Infection with Salmonella enteriditis was considered a significant cause of fur seal pup mortality on the Pribilof Islands, Alaska (Jellison and Milner, 1958), but not in fur seal or California sea lion pups on San Miguel Island, California, from which numerous Salmonella isolates were obtained (Gilmartin et al., 1979). Keyes (1963) diagnosed necrotic enteritis and hemorrhagic colitis in northern fur seals. Koski and Vandenbroek (1986) described gastrointestinal signs including anorexia, vomiting, and liquid stools, in Pacific harbor seals from which Plesiomonas spp. were isolated. Beckmen and Nolan (1994) and Thornton et al. (1998) isolated Salmonella spp. from the lower gastrointestinal tract of nearly 10% of the 865 stranded pinnipeds they sampled. Calle et al. (1995) reported multiple Salmonella spp. isolations from captive walruses. Gastritis and gastric ulcers, well-recognized problems in cetaceans, may, as in humans, have a bacterial component as demonstrated by the recent isolation of Helicobacter spp. from gastric mucosal samples of dolphins (Fox et al., 2000). Diamond et al. (1980) described β-hemolytic Escherichia coli–induced edema disease in a California sea lion that had lesions similar to those seen in swine. Grafton (1968) recommended vaccinating cetaceans for clostridial diseases. Walsh et al. (1994) determined that the clostridial toxin most commonly found in cetacean Clostridium perfringens isolates was the type A variant, and they recommended the use of oral nonsystemically absorbable antibiotics.
0839_frame_C16 Page 328 Tuesday, May 22, 2001 11:12 AM
328
CRC Handbook of Marine Mammal Medicine
Vomiting, diarrhea, flatulence, and abdominal cramping have been observed in cetaceans with bacterially induced gastrointestinal disease. Frothy, floating, off-color stools, although not pathognomonic, are useful indicators of enteric problems.
Conclusion The authors’ collective experience has shown that one must always be prepared to make diagnoses of bacterial diseases previously undiagnosed in marine mammals. New genera of bacteria continue to be reported from marine mammals. Furthermore, a review of the references in this chapter reveals that several bacterial diseases previously unknown in marine mammals have recently been diagnosed. Often, reports of such diagnoses are not published, or are found only in source material not ordinarily accessed. Interchange with attendees at annual meetings of such organizations as the International Association for Aquatic Animal Medicine (IAAAM), the European Association for Aquatic Animal Medicine (EAAAM), and the American Association of Zoo Veterinarians (AAZV) will help expand the thinking on the wide range of bacterial agents that affect marine mammals. Internet “user groups” such as MARMAM, AQUACARE, COMPMED, and PROMED (see Chapter 8, The Electronic Whale) are all valuable resources that provide their users access to a host of scientists and practitioners who possess a wealth of collective clinical experiences that no individual could ever hope to duplicate. In nearly all cases, these individuals will freely relate information that will enable us to provide a better quality of life for the marine mammals in our care.
Acknowledgments This work could not have been completed without the assistance of Donna Zyry of Pfizer Central Research, whose assistance with researching the literature was invaluable. Judy Lawrence and Carol House peer-reviewed this chapter, and David St. Aubin kindly reviewed portions of the manuscript. This is contribution number 121 of the Sea Research Foundation, and technical contribution number 2000-06-T from SeaWorld, San Antonio.
References Baker, A.S., Ruoff, K.L., and Madoff, S., 1998, Isolation of Mycoplasma species from a patient with seal finger, Clin. Infect. Dis., 27: 1168–1170. Bastida, R., Loureiro, J., Quse, V., Bernardelli, A., Rodriguez, D., and Costa, E., 1999, Tuberculosis in a wild subantarctic fur seal from Argentina, J. Wildl. Dis., 35: 796–798. Beaman, B.L., and Beaman, L., 1994, Nocardia species: Host–parasite relationships, Clin. Microbiol. Rev., 7: 213–264. Beaman, B.L., Boiron, P., Beaman, L., Brownell, G.H., Schaal, K., and Gombert, M.E., 1992, Nocardia and nocardiosis, J. Med. Vet. Mycol., 30: 317–331. Beckmen, K.B., and Nolan, S., 1994, Salmonella serotypes in pinnipeds at a rehabilitation center, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 162. Benkovsky, L.M., and Golovina, T.I., 1971, Infestation of B-perfringens pyocyanem [sic] schroeter and Erysipelothrix rhusiopathiae in fur seals, Tr. Vses. Nauchno-Issled. Inst. Morsk. Rybn. Khoz. Oceanogr., 82: 124 [in Russian; English summary]. Bernadelli, A., Nader, A.J., Loureiro, J., Michelis, H., and Debenedetti, R., 1990, Micobacteriosis en mamiferos y aves marinas, Rev. Sci. Tech. Of. Int. Epizootie, 9: 1121–1129. Blair, W.R., 1913, Report of the veterinarian, in 17th Annual Report of New York Zoological Society, New York, 73 pp.
0839_frame_C16 Page 329 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
329
Boever, W.J., Thoen, C.O., and Wallach, J.D., 1976, Mycobacterium chelonei infection in a Natterer manatee, J. Am. Vet. Med. Assoc., 169: 927–929. Bossart, G.D., and Eimstad, E., 1988, Erysipelothrix vesicular glossitis in a killer whale (Orcinus orca), J. Zoo. Anim. Med., 19: 42–47. Brew, S.D., Perrett, L.L., Stack, J.A., and MacMilllan, A.P., 1999, Human exposure to Brucella recovered from a sea mammal, Vet. Rec., 144: 483. Bricker, B.J., Ewalt, D.R., MacMillan, A.P., Foster, G., and Brew, S.D., 2000, Molecular characterization of Brucella strains isolated from marine mammals, J. Clin. Microbiol., 38: 1258–1262. Brown, D.H., McIntyre, R.W., DelliQuadri, C.A., and Schroeder, R.J., 1960, Health problems of captive dolphins and seals, J. Am. Vet. Med. Assoc., 137: 534. Brown, S.R., Heidel, J.R., and Glenn, M., 1994, Case report: Hyperkeratosis in a harbor seal, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 137–140. Buck, J.D., 1982, Nonstaining (KOH) method for determination of Gram reactions of marine bacteria, Appl. Environ. Microbiol., 44: 992. Buck, J.D., and Spotte, S., 1986a, Microbiology of captive white-beaked dolphins, Lagenorhynchus albirostris, with comments on epizootics, Zoo Biol., 5: 321. Buck, J.D., and Spotte, S., 1986b, The occurrence of potentially pathogenic vibrios in marine mammals, Mar. Mammal Sci., 2: 319. Buck, J.D., Spotte, S., and Gadbaw, J.J., 1984, Bacteriology of the teeth from a great white shark: Potential medical implications for shark bite victims, J. Clin. Microbiol., 20: 849. Buck, J.D., Shepard, L.L., and Spotte, S., 1987, Clostridium perfringens as a cause of death of a captive Atlantic bottlenose dolphin (Tursiops truncatus), J. Wildl. Dis., 23: 488. Buck, J.D., Bubucis, P.M., and Spotte, S., 1988, Microbiological characterization of three Atlantic whiteside dolphins (Lagenorhynchus acutus) from stranding through captivity with subsequent rehabilitation and release of one animal, Zoo Biol., 7: 133–138. Buck, J.D., Overstrom, N.A., Patton, G.W., Anderson, H.F., and Gorzelany, J.F., 1991, Bacteria associated with stranded cetaceans from the northeast USA and southwest Florida coast, Dis. Aquat. Organisms, 10: 147–152. Calle, P.P., Kenny, D.E., and Cook, R.A., 1993, Successful treatment of suspected erisypelas septicemia in a beluga whale (Delphinapterus leucas), Zoo Biol., 12: 483–490. Calle, P.P., Stetter, M.D., Cook, R.A., McClave, C.A., and Massucci, S., 1995, Enteric salmonellosis of captive Pacific walrus (Odobenus rosmarus divergens), in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 92–93. Castro Ramos, M., Ayla, M., Errico, F., and Silvera, F.V., 1998, Aislamiento de Mycobacterium bovis en Pinnipedos Otaria byronia (Lobo marino comun) en Uraguay, Rev. Med. Vet., 79: 197–200. Clavareau, C., Wellemans, V., Walravens, K., Tryland, M., Verger, J.M., Grayon, M., Cloeckaert, A., Letesson, J.J., and Godfroid, J., 1998, Phenotypic and molecular characterization of a Brucella strain isolated from a minke whale (Balaenoptera acutorostrata), Microbiology, 144: 3267–3273. Cohen, M.A., Mitchell, G., and Sullivan, N., 1993, Melioidosis in two Australian pinnipeds, in Proceedings of the 29th Annual Workshop of the International Association for Aquatic Animal Medicine, 72–74. Coles, B.M., Stroud, R.K., and Sheggeby, S., 1978, Isolation of Edwardsiella tarda from three Oregon sea mammals, J. Wildl. Dis., 14: 339. Cousins, D.V., 1987, ELISA for the detection of tuberculosis in seals, Vet. Rec., 121: 305. Cousins, D.V., Francis, B.R., Gow, B.L., Collins, D.M., McGlashan, C.H., Gregory, A., and Mackenzie, R.M., 1990, Tuberculosis in captive seals: Bacteriological studies on an isolate belonging to the Mycobacterium tuberculosis complex, Res. Vet. Sci., 48: 196–200. Cousins, D.V., Williams, S.N., Reuter, R., Forshaw, D., Chadwick, B., Coughran, D., Collins, P., and Gales, N., 1993, Tuberculosis in wild seals and characterisation of the seal bacillus, Aust. Vet. J., 70: 92–97.
0839_frame_C16 Page 330 Tuesday, May 22, 2001 11:12 AM
330
CRC Handbook of Marine Mammal Medicine
Cowan, D.F., Turnbull, B.S., and Haubold, E.M., 1998, Organisms cultured from stranded cetaceans: Implications for rehabilitation and for safety of handlers, in Proceedings of the 29th Annual Workshop of the International Association for Aquatic Animal Medicine, 148–149. Cusick, P.K., and Bullock, B.C., 1973, Ulcerative dermatitis and pneumonia associated with Aeromonas hydrophila infection in the bottle-nosed dolphin, J. Am. Vet. Med. Assoc., 163: 578. Dailey, M. D., 1985, Diseases of Mammalia: Cetacea, in Diseases of Marine Animals, Kinne, O. (Ed.), Biologische Anstalt Helgoland, Hamburg, 805. Dalton, L.M., and Robeck, T.R., 1995, Multifocal pyogranulomatous pneumonia in a killer whale calf, in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 26: 84–85. De Guise, S., Martineau, D., Beland, P., and Fournier, M., 1995a, Possible mechanisms of action of environmental contaminants on St. Lawrence beluga whales, Environ. Health Perspect., 103(Suppl. 4): 73–77. De Guise, S., Lagace, A., Beland, P., and Higgins, R., 1995b, Non-neoplastic lesions in beluga whales (Delphinapterus leucas) and other marine mammals from the St. Lawrence estuary, J. Comp. Pathol., 112: 257–271. Diamond, S.S., Raflo, C.P., Beleau, M.H., and Caldwell, G.A., 1980, Edema disease in a California sea lion, J. Am. Vet. Med. Assoc., 177: 808. Dierauf, L.A., Vandenbroek, D.J., Roletto, J., Koski, M., Amaya, L., and Gage, L.J., 1985, An epizootic of leptospirosis in California sea lions, J. Am. Vet. Med. Assoc., 187: 1145. Dunn, J.L., Bowenkamp, K., Frasca, S., Montali, R.J., Goertz, C.E.C., Draghi, A.J., Tsongalis, G.J., and St. Aubin, D.J., 2000, Fusariomycosis, atypical mycobacterial pyogranulomatous panniculitis, proliferative pleuritis and fatal aortic rupture in a white whale, Delphinapterus leucas, in Proceedings of the Joint Meeting of American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, New Orleans, LA, 295–296. Ehlers, K., 1965, Records of the hooded seal (Cystophora crestata) [sic] and the other animals at the Bremerhaven Zoo, Int. Zoo Yearb., 5: 149–152. Ewalt, D.R., Payeur, J.B., Martin, B.M., Cummins, D.R., and Miller, W.G., 1994, Characterization of a Brucella species from a bottlenose dolphin (Tursiops truncatus), J. Vet. Diagn. Invest., 6: 448–452. Forbes, L.B., Nielsen, O., Measures, L., and Ewalt, D.R., 2000, Brucellosis in ringed seals and harp seals from Canada, J. Wildl. Dis., 36: 595–598. Forshaw, D., and Phelps, G.R., 1991, Tuberculosis in a captive colony of pinnipeds, J. Wildl. Dis., 27: 288–295. Foster, G., Jahans, K.L., Reid, R.J., and Ross, R.M., 1996, Isolation of Brucella species from cetaceans, seals and an otter, Vet. Rec., 138: 583–586. Fox, J.G., Harper, C.M.G., Dangler, C.A., Xu, S., Stamper, A., and Dewhirst, F.E., 2000, Isolation and characterization of Helicobacter sp. from the gastric mucosa of dolphins, in Proceedings of the Joint Meeting of American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, New Orleans, LA, 339–340. Fujioka, R.S., Greco, S.B., Cates, M.B., and Schroeder, J.P., 1988, Vibrio damsela from wounds in bottlenose dolphins, Tursiops truncatus, Dis. Aquat. Organisms, 4: 1. Gales, N., 1993, Tuberculosis in wild seals and characterisation of the seal bacillus, Aust. Vet. J., 70: 92–97. Gall, D., Nielsen, K., Forbes, L., Davis, D., Elzer, P., Olsen, S., Balsevicius, S., Kelly, L., Smith, P., Tan, S., and Joly, D., 2000, Validation of the fluorescence polarization assay and comparison to other serological assays for the detection of serum antibodies to Brucella abortus in bison, J. Wildl. Dis., 36: 469–476. Gallacher, J.B., and Waters, W.E., 1964, Pneumonia in grey seal pups at St. Kilda, Proc. Zool. Soc. London, 142: 177. 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. Geraci, J.R., Sauer, R.M., and Medway, W., 1966, Erysipelas in dolphins, Am. J. Vet. Res., 27: 597.
0839_frame_C16 Page 331 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
331
Geraci, J.R., St. Aubin, D.J., Barker, I.K., Webster, R.G., Hinshaw, V.S., Bean, W.J., Rukake, H.J., Prescott, J.M., Early, G., Baker, A.S., Madoff, S., and Schooley, R.T., 1982, Mass mortality of harbor seals: Pneumonia associated with influenza A virus, Science, 215: 1129. Gerber, J.A., Roletto, J., Morgan, L.E., Smith, D.M., and Gage, L.J., 1993, Findings in pinnipeds stranded along the central and northern California coast, 1984–1990, J. Wildl. Dis., 29: 423–433. Gilmartin, W.G., Allen, J.F., and Ridgway, S.H., 1971, Vaccination of porpoises (Tursiops truncatus) against Erysipelothrix rhusiopathiae infection, J. Wildl. Dis., 7: 292. Gilmartin, W.G., Vainik, P.M., and Neill, V.M., 1979, Salmonellae in feral pinnipeds of the southern California coast, J. Wildl. Dis., 15: 511. Grafton, T.S., 1968, Necropsy of a sacred dolphin of the Amazon (Sotalia fluviatilis), in Proceedings of the 2nd Symposium on the Diseases and Husbandry of Aquatic Mammals. Greco, S.B., Fujioka, R.S., Cates, M.B., and Schroeder, J.P., 1985, Assessing the concentrations and significance of Vibrio bacteria in pens used to house marine mammals, in Proceedings of the 16h Annual Workshop of the International Association for Aquatic Animal Medicine, 139. Greco, S., Fujioka, R.S., and Schroeder, J.P., 1986, Patterns of antiobiotic resistance in Vibrio spp. isolated from cold-blooded animals, marine mammals and environmental sources, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 132. Gulland, F.M.D., Koski, M., Lowenstine, L.J., Colagross, 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. Gutter, A.E., Wells, S.K., and Spraker, T.R., 1987, Generalized mycobacteriosis in a California sea lion (Zalophus californianus), J. Zoo Anim. Med., 18: 118–120. Hall, J.D., Gilmartin, W.D., and Mattson, J.L., 1971, Investigation of a Pacific pilot whale stranding on San Clemente Island, J. Wildl. Dis., 7: 324. Harvell, C.D., Kim, K., Burkholder, J.M., Colwell, R.R., Epstein, P.R., Grimes, D.J., Hoffman, 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: 1505–1509. Hawkins, E.C., Townsend, F.I., Lewbart, G.A., Stamper, M.A., Thayer, V.G., and Rhinehart, H.L., 1996, Bronchoalveolar lavage in a stranded bottlenose dolphin, in Proceedings of the 27th Annual Workshop of the International Association for Aquatic Animal Medicine, 124. Hermans, J., Cox, E., Goddeeris, B., and Lacave, G., 1997, Protection of mice against dolphin erysipelas isolates following vaccination with a commercial erysipelas vaccine, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 82. 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. Higgins, R., 2000, Bacteria and fungi of marine mammals, Can. Vet. J., 41: 105–116. Howard, E.B., Britt, J.O., Matsumoto, G.K., Itahara, R., and Nagano, C.N., 1983, Bacterial diseases, in Pathobiology of Marine Mammal Diseases, Vol. 1, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 69–118. Hunter, J.E.B., Duignan, P.J., Dupont, C., Fray, L., Fenwick, S.G., and Murray, A., 1998, First report of potentially zoonotic tuberculosis in fur seals in New Zealand, N. Z. Med. J., April 10: 130–131. Jahans, K.L., Foster, G., and Broughton, E.S., 1997, The characterization of Brucella strains isolated from marine mammals, Vet. Microbiol., 57: 373–382. Jasmin, C., Powell, C.P., and Baucom, J.N., 1972, Actinomycotic mycetoma in the bottlenose dolphin (Tursiops truncatus) due to Nocardia paraguayenesis, Vet. Med. Small Anim. Clin., 67: 542–543. Jellison, W.L., and Milner, K.C., 1958, Salmonellosis (bacillary dysentery) of fur seals, J. Wildl. Manage., 22: 199. Jensen, A.E., Cheville, N.F., Thoen, C.O., MacMillan, A.P., and Miller, G.W., 1999, Genomic fingerprinting and development of a dendrogram for Brucella spp. isolated from seals, porpoises, and dolphins, J. Vet. Diagn. Invest., 11: 152–157.
0839_frame_C16 Page 332 Tuesday, May 22, 2001 11:12 AM
332
CRC Handbook of Marine Mammal Medicine
Jepson, P.D., Brew, S., MacMillan, A.P., Baker, J.R., Barnett, J., Kirkwood, J.K., Kuiken, T., Robinson, I.R., and Simpson, V.R., 1997, Antibodies to Brucella in marine mammals around the coast of England and Wales, Vet. Rec., 141: 513–515. 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. Jones, J.C., Patterson, R.A., and Middlebrooks, B.L., 1995, A biotin-avidin amplified ELISA for the detection of antibodies against Erysipelothrix rhusiopathiae in captive dolphins, in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 54. Jones, J.C., Patterson, R.A., and Middlebrooks, B.L., 1996, Comparison of ELISA and agglutination titers against Erysipelothrix rhusiopathiae in cetaceans, in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 52. Kennedy-Stoskopf, S., Stoskopf, M.K., Eckhaus, M.A., and Strandberg, J.D., 1986, Isolation of a retrovirus and a herpesvirus from a captive California sea lion, J. Wildl. Dis., 22: 156. Keyes, M.C., 1963, Necrotic hemorrhagic enteritis in a seal, Small Anim. Clin., 3: 627. Kilgen, M.B., 2000, An overview of ionizing irradiation technology for pathogen control in fishery feed products for marine mammals, in Proceedings of the Joint Meeting of American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, New Orleans, LA, 200–202. Kinoshita, R., Brook, F., Vedros, N., Wad, H.S., Lung, R., Ng, T., Tao, L.Y., and Yuen, C.S., 1994, Staphylococcal isolations and clinical cases of Staphylococcus aureus in bottlenose dolphins at Ocean Park, Hong Kong, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 159. Kinsel, M.J., Boehm, J.R., Harris, B., and Murnane, R.D., 1997, Fatal Erysipelothrix rhusiopathiae septicemia in a captive Pacific white-sided dolphin (Lagenorhynchus obliquidens), J. Zoo Wildl. Med., 28: 494–497. Koski, M., and Vandenbroek, D.J., 1986, Plesiomonas shigelloides-associated gastroenteritis in harbor seals (Phoca vitulina richardsi), in Proceedings of the 17th Annual Workshop of the International Association for Aquatic Animal Medicine, 125–131. Lauckner, G., 1985, Diseases of Mammalia: Pinnipedia, in Diseases of Aquatic Animals, Kinne, O. (Ed.), Biologische Anstalt Helgoland, Hamburg, 654 pp. Lipscomb, T.P., Kennedy, S., Moffett, D., and Ford, B.K., 1994, Morbilliviral disease in an Atlantic bottlenose dolphin (Tursiops truncatus) from the Gulf of Mexico, J. Wildl. Dis., 30(4): 572–576. Makino, S., Okada, Y., Maruyama, T., Ishikawa, K., Takahashi, T., Nakamura, M., Ezaki, T., and Morita, H., 1994, Direct and rapid detection of Erysipelothrix rhusiopathiae DNA in animals by PCR, J. Clin. Microbiol., 32: 1526–1531. Martineau, D., Lagacé, 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, Can. J. Comp. Pathol., 98: 287–311. McNeil, M.M., and Brown, J.M., 1994, The medically important aerobic actinomycetes: Epidemiology and microbiology, Clin. Microbiol. Rev., 7: 357–417. Measures, L., 1998, Oral mycoplasmal infections in Canadian pinnipeds, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 50. Medway, W., and Schryver, H.F., 1973, Respiratory problems in captive small cetaceans, J. Am. Vet. Med. Assoc., 163: 571. Meyer, W., 1997, Immunohistochemical demonstration of transferrin and transferrin receptor in mammalian integument, Biotech. Histochem., 72: 223–228. Meyer, W., Bollhorn, M., and Stede, M., 2000, Aspects of general antimicrobial properties of skin secretions in the common seal Phoca vitulina, Dis. Aquat. Organisms, 41: 77–79. Miller, W.G., Adams, L.G., Ficht, T.A., Cheuible, N.F., Payeur, J.P., Harley, D.R., House, C., and Ridgway, S.H., 1999, Brucella-induced abortions and infection in bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med., 30: 100–110.
0839_frame_C16 Page 333 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
333
Morales, P., Madin, S.H., and Hunter, A., 1985, Systemic Mycobacterium marinum infection in an Amazon manatee, J. Am. Vet. Med. Assoc., 187: 1230–1231. Mullan, D.L., 1991, A Classification of Epidermal Lesions Observed on the Bowhead Whale, Balaena mysticetus, and the Initial Steps in the Development of a Probe for the Identification of Lesion Associated Bacteria, Master’s thesis, Louisiana State University and Agricultural and Mechanical College, Baton Rouge, LA, 84 pp. Munford, R.S., 1994, Sepsis and septic shock, in Harrison’s Principles of Internal Medicine, Isselbacher, K.J., Braunwald, E., Wilson, J.O., Martin, J.B., Fauci, A.S., and Kasper, D.L. (Eds.), McGraw-Hill, New York, 511–515. Needham, D.J., 1992, Tuberculosis/mycobacteriosis in pinnipeds, in Proceedings of the 23rd Annual Workshop of the International Association for Aquatic Animal Medicine, 92–96. Nielsen, O., Nielsen, K., and Stewart, R.E., 1996, Serologic evidence of Brucetta spp. exposure in Atlantic walruses (Odobenus rosmarus rosmarus) and ringed seals (Phoca hispida), Arctic, 49: 383–386. Parsons, E.C.M., and Jefferson, T.A., 2000, Post-mortem investigations of stranded dolphins and porpoises from Hong Kong waters, J. Wildl. Dis., 36: 342–356. Patterson, R.A., 2000, What’s new in Erysipelothrix rhusiopathiae research in marine mammals, in Proceedings of the Joint Meeting of American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, New Orleans, LA, 210. Patterson, R.A., Jones, J.C., and Middlebrooks, B.L., 1997, Determination of serum antibody titers against Erysipelothrix rhusiopathiae in Delphinapterus leucas and Lagenorhynchus obliquidens, in Proceedings of the 28th Annual Workshop of the International Association for Aquatic Animal Medicine, 79. Pier, A.C., and Fitchner, R.E., 1981, Distribution of serotypes of Nocardia asteroides from animal, human and environmental sources, J. Clin. Microbiol., 13: 548–553. Pier, A.C., Takayama, A.K., and Miyahara, A.Y., 1970, Cetacean nocardiosis, J. Wildl. Dis., 6: 112–118. Rand, C.S., 1979, Nodular suppurative cutaneous cellulitis in a Galapagos sea lion, J. Wildl. Dis., 11: 325. Robeck, T.R., and Dalton, L.M., 1995, Subcutaneous nocardiosis in a beluga whale, in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 26. Robeck, T., Dalton, L.M., and Young, W.G., 1994, Systemic nocardiosis in a 6 week old beluga calf, in Proceedings of the 25th Annual Workshop of the International Association for Aquatic Animal Medicine, 109. Robeck, T.R., Dalton, L.M., and Young, W.G., 1995, Nocardia spp. induced chronic suppurative osteomylitis in a beluga whale, in Proceedings of the 26th Annual Workshop of the International Association for Aquatic Animal Medicine, 28. Robeck, T.R., Dalton, L.M., and Young, W.G., 1996, Single daily dosing of amikacin in two beluga whales and an Asian small clawed otter, in Proceedings of the 27th Annual Workshop of the International Association for Aquatic Animal Medicine, 19–21. Ross, H.M., Jahans, K.L., MacMillan, A.P., Reid, R.J., Thompson, P.M., and Foster, G., 1996, Brucella species infection in North Sea seal and cetacean populations, Vet. Rec., 138: 647–648. Schoenwald, J., Migaki, G., and Hubbard, R.C., 1971, Leptospirosis epizootic among California sea lions, Science, 172: 1250. Siebold, H.R., and Neal, J.C., 1956, Erysipeolthrix septicemia in the porpoise, J. Am. Vet. Med. Assoc., 128: 537. Smith, A.W., Prato, C.M., Gilmartin, W.G., Brown, R.J., and Keyes, M.C., 1974a, Preliminary report on potentially pathogenic microbial agents recently isolated from pinnipeds, J. Wildl. Dis., 10: 54. Smith, A.W., Brown, R.J., Skilling, D.E., and DeLong, R.L., 1974b, Leptospira pomona and reproductive failure in California sea lions, J. Am. Vet. Med. Assoc., 165: 996. Smith, A.W., Brown, R.J., Skilling, D.E., Bray, H.L., and Keyes, M.C., 1977, Naturally occurring leptospirosis in northern fur seals (Callorhinus ursinus), J. Wildl. Dis., 13: 144. Stamper, M.A., Gulland, F.M.D., and Spraker, T., 1998, Leptospirosis in rehabilitated Pacific harbor seals in California, J. Wildl. Dis., 34: 407–410.
0839_frame_C16 Page 334 Tuesday, May 22, 2001 11:12 AM
334
CRC Handbook of Marine Mammal Medicine
Stevens, E., Lipscomb, T.P., and Gulland, F.M.D., 1999, An additional case of leptospirosis in a harbor seal, J. Wildl. Dis., 35: 150. Stoskopf, M.K., Moench, K.T., Thoen, T., and Charache, P., 1987, Tuberculosis in pinnipeds, in Proceedings of the Annual Meeting of the American Association of Zoo Veterinarians, 393. Stroud, R.K., and Jaffe, T.J., 1979, Causes of death in marine mammals stranded along the Oregon coast, J. Wildl. Dis., 15: 91. Suer, L.D. and Vedros, N.A., 1988, Erysipelothrix rhusiopathiae. I. Isolation and characterization from pinnipeds and bite/abrasion wounds in humans, Dis. Aquat. Organisms, 5: 1–5. Suer, L.D., Vedros, N.A., Schroeder, J.P., and Dunn, J.L., 1988, Erysipelothrix rhusiopathiae. II. Enzyme immunoassay of sera from wild and captive marine mammals, Dis. Aquat. Organisms, 5: 7–13. Sugita, H., Kumazawa, J., and Daguchi, Y., 1996, Production of chitinase and beta-N-acetylglucosaminidase by intestinal bacteria of pinnipedian animals, Lett. Appl. Microbiol., 23: 275–278. Sweeney, C., 1974, Common diseases of pinnipeds, J. Am. Vet. Med. Assoc., 165: 805. Sweeney, J.C., 1978, Infectious diseases, in Zoo and Wild Animal Medicine, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 777. Sweeney, J.C., and Gilmartin, W.G., 1974, Survey of diseases in free-living California sea lions, J. Wildl. Dis., 10: 370. Sweeney, J.C., and Ridgway, S.H., 1975, Common diseases of small cetaceans, J. Am. Vet. Med. Assoc., 167: 533. Sweeney, J.C., Miyaki, G., Vainik, P.M., and Conklin, R.H., 1976, Systemic mycoses in marine mammals, J. Am. Vet. Med. Assoc., 169: 946–948. Tangredi, R.C., and Medway, W., 1980, Post-mortem isolation of Vibrio alginolyticus from an Atlantic white-sided dolphin (Lagenorhynchus acutus), J. Wildl. Dis., 16: 329. 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. Thorel, M.F., and Moutou, F., 1994, Tuberculose et animaux sauvages, Point Vet., 26: 27–43. Thorel, M.F., Karoui, C., Varnerot, A., Fleury, C., and Vincent, V., 1998, Isolation of Mycobacterium bovis from baboons, leopards and a sea-lion, Vet. Res., 29: 207–212. Thornton, S.M., Nolan, S., and Gulland, F.M.D., 1998, Bacterial isolates from California sea lions (Zalophus californianus), harbor seals (Phoca vitulina), and northern elephant seals (Mirounga angustirostris) admitted to a rehabilitation center along the central California coast, J. Zoo Wildl. Med., 29: 171–176. VanPelt, R.W., and Dieterich, R.A., 1973, Staphylococcal infection and toxoplasmosis in a young harbor seal, J. Wildl. Dis., 9: 258. Vedros, N.A., 1982, A potential vaccine for Pasteurella multocida in marine mammals, in Proceedings of the 13th Annual Workshop of the International Association for Aquatic Animal Medicine, 51. Vedros, N.A., Smith, A.W., Schoenwald, J., Migaki, G., and Hubbard, R.C., 1971, Leptospirosis epizootic among California sea lions, Science, 172: 1250. Visser, I.J., Vellema, P., van Dokkum, H., and Shimada, T., 1999, Isolation of Vibrio cholerae from diseased farm animals and surface water in the Netherlands, Vet. Rec., 144: 451–452. Walsh, M.T., Thomas, L.A., Songer, J.G., Campbell, T.W., and Tucker, L.S., 1994, Clostridium perfringens isolates from cetaceans, in Proceedings of the 28th Annual Workshop of the International Association for Aquatic Animal Medicine, 95. Wells, S.K., Gutter, A., and Van Meter, K., 1990, Cutaneous mycobacteriosis in a harbor seal: Attempted treatment with hyperbaric oxygen, J. Zoo Wildl. Med., 21(1): 73–78. West, P.A., and Colwell, R.R., 1984, Identification and classification of Vibrionaceae: An overview, in Vibrios in the Environment, Colwell, R.R. (Ed.), John Wiley & Sons, New York, 285 pp. Weynants, V., Godfroid, J., Limbourg, B., Saegerman, C., and Letesson, J.J., 1995, Specific bovine brucellosis diagnosis based on in vitro antigen-specific gamma interferon production, J. Clin. Microbiol., 33: 706–712.
0839_frame_C16 Page 335 Tuesday, May 22, 2001 11:12 AM
Bacterial Diseases of Cetaceans and Pinnipeds
335
Williams, S.T., Lanning, S., and Wellington, E.M.H., 1983, Ecology of actimomycetes, in The Biology of Actinomycetes, Goodfellow, M., Mordarski, M., and Williams, S.T. (Eds.), Academic Press, London, 481–528. Wilson, B., Arnold, A., Bearzi, G., Fortuna, C.M., Gaspar, R., Ingram, S., Liret, C., Pribanic, S., Read, A.J., Ridoux, V., Schneider, K., Urian, K.W., Wells, R.S., Wood, C., Thompson, P.M., and Hammond, P.S., 1999, Epidermal disease in bottlenose dolphins: Impacts of natural and anthropogenic factors, Proc. R. Soc. London B, 266: 1077–1083. Young, J.F., Huff, D.G., Ford, J.K.B., Anthony, J.M.G., Ellis, G., and Lewis, R.L., 1997, First case report—Mortality of wild resident killer whale (Orcinus orca) from Erysipilothrix rhusopathiae, in Proceedings of the 28th Annual Workshop of the International Association for Aquatic Animal Medicine, 97. Ziesenis, A., Bernard, T., Petermann, M., Franz, B., and Leibold, W., 1992, Monoclonal antibodies preventing the development of polyarthritis in rats induced by experimental infection with erysipelas bacteria, Scand. J. Rheumatol., 21: 60–67.
0839_frame_C16 Page 336 Tuesday, May 22, 2001 11:12 AM
0839_frame_C17.fm Page 337 Tuesday, May 22, 2001 11:12 AM
17 Mycotic Diseases Thomas H. Reidarson, James F. McBain, Leslie M. Dalton, and Michael G. Rinaldi
Introduction Clinicians face an immense challenge diagnosing mycotic infections in sick animals. Even more challenging is determining the significance of finding a yeast or mold in a specimen from a clinically normal individual. Should treatments commence after finding a potentially pathogenic fungus such as Aspergillus fumigatus in the blowhole of a cetacean or Candida spp. in a skin or gastric lesion? Unfortunately, relatively little is known concerning mycotic infections in marine mammals. Interestingly, since the early work on mycoses (Sweeney et al., 1976), the development of new therapeutic agents has advanced, while the ability to make timely ante-mortem diagnoses has lagged. This chapter discusses the diagnostic challenges faced by clinicians and current therapeutic modalities for mycotic diseases in marine mammals.
Mycotic Diseases The most common pathogenic fungi belong to an artificial taxon called the Fungi Imperfecti (asexual fungi), which includes A. fumigatus, Blastomyces dermatitidis, Candida spp., Cryptococcus neoformans, Coccidioides immitis, and Histoplasma capsulatum, and the Class Zygomycetes (which includes Apophysomyces elegans, Rhizomucor pusillus, Saksenaea vasiformis, and others), as well as an unclassified fungus called Lacazia loboi (formerly Loboa loboi), perhaps related to B. dermatitidis. Aspergillus fumigatus, Candida spp., Cryptococcus neoformans, Fusarium spp., and the zygomycetes are true opportunistic pathogens, whereas the other endemic pathogens (B. dermatitidis, Coccidioides immitis, and H. capsulatum) are capable of infecting healthy hosts (Rippon, 1988). In humans and animals, mycoses represent only a small, but often critically significant, fraction of infectious diseases (Nicholls et al., 1993). A recent survey of mycotic infections in captive and wild marine mammals contributed by 92 clinicians and pathologists reported 168 cases, including 22 species of fungi in 27 species of marine mammals (Reidarson et al., 1999). Of these cases, 44% were stranded animals, of which 60% had another underlying illness, and the remaining 56% were residents of various oceanaria. Of the captive animals, 40% had some type of preexisting disease, 54% were apparently healthy, and 6% percent were neonates. The most commonly reported mycotic infection in any marine mammal is pulmonary aspergillosis, which has been isolated from ten species of marine mammals. The greatest number
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
337
0839_frame_C17.fm Page 338 Tuesday, May 22, 2001 11:12 AM
338
CRC Handbook of Marine Mammal Medicine
of cases has occurred in stranded bottlenose dolphins (Tursiops truncatus) infected with dolphin morbillivirus, an immunosuppressive virus closely related to, but distinct from, human measles and canine distemper (see Chapter 15, Viral Diseases). Other opportunistic fungi include systemic and local Candida albicans infections in ten species of marine mammals, Cryptococcus neoformans infections in cetaceans and sea otters (Enhydra spp.), systemic zygomycetes infections in nine species of cetaceans and a harp seal (Pagophilus groenlandicus), and dermatophyte infections including Fusarium spp., Malassezia pachydermatis, Microsporum canis, Sporothrix schenckii, Trichophyton spp., and Trichosporon pullulans in cetaceans and pinnipeds. Infections involving endemic fungi include systemic Blastomyces dermatitidis in a bottlenose dolphin, a gray seal (Halichoerus grypus), and a California sea lion (Zalophus californianus); systemic Coccidioides immitis infections in California sea otters (Enhdra lutris nereis), California sea lions, and a bottlenose dolphin; disseminated Histoplasma capsulatum infections in bottlenose dolphins, a false killer whale (Pseudorca crassidens), a harp seal, and a Pacific white-sided dolphin (Lagenorhynchus obliquidens); and Lacazia loboi (keloidal blastomycosis) in a freshwater dolphin (Sotalia fluviatilis guianensis) and bottlenose dolphins.
Epidemiology of Fungi Modes of Transmission Each of the fungi, except Candida spp. and L. loboi, is ubiquitous in certain environments. They grow as saprobes, producing mycelia and/or conidia, or exist as yeasts comprising a part of the normal microbiota or in stool (in the case of Cryptococcus neoformans) as pathogenic forms capable of producing infection. These fungi gain entry into hosts by inhalation, trauma, or ingestion, and then settle into the lungs, skin, or alimentary tract (Muller, 1994). Candida albicans is a normal fungal resident of mucous membranes, where it resides as a commensal. Buck (pers. comm., 1996) found C. albicans and other Candida species in 4 to 54% of wild bottlenose dolphins examined in Sarasota Bay, Florida from 1990 to 1992. This is comparable with what is observed in many cetaceans living in modern oceanaria. Because fungi are poorly communicable between animals, mycoses are often endemic and rarely epidemic (Rippon, 1988). Except for two reported zoonotic cases involving Blastomyces dermatitidis in bottlenose dolphins (Cates et al., 1986), and L. loboi in an Amazon dolphin (de Vries and Laarman, 1973), infection of a host by a fungus is generally dead-end, and is not contagious to other hosts or capable of becoming widespread in the species (Geraci and Ridgway, 1991; Nicholls et al., 1993).
Mechanisms of Pathogenesis After invading the alveoli of the host, the conidia of the endemic dimorphic fungi (B. dermatitidis, Coccidioides immitis, and Histoplasma capsulatum) transform into parasitic forms and stimulate local humoral and cellular immune responses that lead to granuloma formation, caseation, fibrosis, and ultimately calcification (Walsh and Mitchell, 1991). Most infections are self-limiting and produce minimal clinical signs. However, some become blood-borne and disseminate (Christin and Sugar, 1996; Wheat, 1996). For a normal host, the skin and mucosa are effective barriers against invasive candidiasis. However, when these barriers are broken and/or the host’s defense mechanisms are impaired, local invasion and fungemia may occur. Candida spp. may infect any organ; the kidneys, central nervous system, and heart valves are the most common sites of systemic invasion (Nicholls et al., 1993).
0839_frame_C17.fm Page 339 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases
339
Because many animals have resident Candida spp. microflora, the discovery of fungus in blowhole, stomach, or stool samples may cause some confusion. Unless a thorough cytological examination reveals evidence of local invasion, such as the presence of budding yeast cells, pseudohyphae, and/or true hyphal elements, and the presence of inflammatory cells, the fungus should be considered a resident or contaminant, having entered by way of an aerosol or as an accidental introduction at the diagnostic laboratory (Jones, 1990) (see Chapter 20, Cytology, and Clinical Diagnostic Features of Fungi, below). Zygomycetes and Aspergillus fumigatus infections are the most devastating of the filamentous opportunistic fungi. Zygomycete infections begin with entry of spores by inhalation or through wounds. Cutaneous disease arising from disseminated disease, in contrast to wound zygomycosis, usually presents as nodular subcutaneous lesions, which may ulcerate. Disseminated zygomycosis may originate from any of the primary sites of infection, particularly the lung and sinus, or the alimentary tract. Once spores enter the site of infection, they rapidly proliferate and aggressively invade skin or bronchial tissue causing infarction. Dissemination may occur to nearly all internal organs, usually followed by rapid death (Ribes et al., 2000). In contrast, A. fumigatus spores primarily enter through inhalation and produce several forms of disease, namely, allergic aspergillosis, chronic necrotizing aspergillosis, aspergillar fungal balls (aspergillomas), and invasive aspergillosis (Haque, 1992). The first three are chronic debilitating forms, whereas the latter acts similarly to invasive zygomycosis. Finding A. fumigatus or a zygomycete in a culture from a respiratory mucous membrane may be cause for concern. However, unless there is a high clinical suspicion of invasion, the fungus should be considered saprobic, as both are easily airborne and may be demonstrated during sampling of both outdoor and indoor air. Demonstration of invasive disease by these organisms generally requires the identification of fungal elements directly in the clinical specimen, or organism growth from more than one specimen obtained from a normally sterile site (Ribes et al., 2000). Other less common mycotic diseases are caused by Cryptococcus neoformans, Fusarium spp., and L. loboi. Cryptococcal organisms settle in the lungs, producing localized fungal balls (cryptococcomas) and occasionally extrapulmonary dissemination to the brain and meninges. Fusarium species are soil and plant saprobes that are capable of infecting damaged or devitalized skin, primarily causing skin disease. However, when disseminated infection occurs, mortality rates are particularly high (Kwon-Chung and Bennett, 1992). Lobomycosis is a chronic, localized disease of the skin caused by infection with the yeastlike organism called L. loboi (Taborda et al., 1999). The organism has never been successfully cultured, and systemic spread has never been demonstrated.
Clinical Manifestations Clinical presentations of mycotic diseases are frustratingly nonspecific, ranging from chronic to fulminating, just as with bacterial or viral diseases. Fungi can affect any tissue, so unless the fungus can be identified, the origin of the disease may remain unknown. A thorough history focusing on types and extent of previous illnesses and response to past and present treatments may give clues to the identity of the fungus. A disease that initially appears responsive to antibiotics and then apparently becomes unresponsive could be evidence of a change to a mycotic etiology. The only unique presentations are those of lobomycosis, which produces multifocal white crusts involving large areas of skin, and central nervous system mycoses, in which animals have rapidly escalating abnormal neurological clinical signs.
0839_frame_C17.fm Page 340 Tuesday, May 22, 2001 11:12 AM
340
CRC Handbook of Marine Mammal Medicine
Clinical Diagnostic Features of the Fungi Laboratory findings from individual marine mammals with opportunistic mycotic infections often reveal hematological and biochemical changes indistinguishable from bacterial or viral infections (Joseph et al., 1986; Magnussen, 1992; Coleman et al., 1995). An example is an individual with initial stages of zygomycosis in which laboratory findings are unremarkable until the later stages of disease, at which time all muscle and liver specific enzymes escalate rapidly because of tissue infarction. On the other hand, individuals with endemic infections generally have blood work indicative of chronic granulomatous diseases showing persistent leukocytosis, monocytosis, eosinophilia, and hyperproteinemia with hypergammaglobulinemia; however, these features may be inconsistent (Harley and Blaser, 1994; see Chapter 19, Clinical Pathology). Radiography, ultrasonography, and endoscopy aid in localizing lesions (Reef, 1991; McAdams et al., 1995; Hawkins et al., 1996; Harrell et al., 1996). The practice of culturing nasal, gastric, and stool samples has limited use. In cetaceans, there appears to be very low correlation between pneumonia-causing organisms and those organisms isolated from the blowholes of the same animals. In humans, there appears to be a high correlation between bronchoalveolar lavage isolates and pneumonia-causing organisms (J. Harrell, pers. comm.). Preliminary evidence suggests the same correlation exists in dolphins. In fact, for most pulmonary mycoses listed in Table 1, the pathogen discovered at necropsy was not identified in ante-mortem blowhole samples. On the other hand, one extremely important value of sampling nasal exudates is to compare ratios of inflammatory cells to epithelial cells as a means of predicting early infectious respiratory tract disease (Jeraj and Sweeney, 1996; see Chapter 20, Cytology). Biopsy, aspiration, scrapings, or bronchoalveolar lavage are definitive diagnostic procedures. For biopsies, it is advisable to fix one piece of tissue for histopathological study, and to place another directly on selective culture media (e.g., Sabouraud’s dextrose agar). This is because certain fungi, especially the zygomycetes, may be difficult to grow from a culturette swab only, and may be rendered nonviable if the tissue is ground before plating out on microbiologic media (Rinaldi, 1989). Many produce erect aerial mycelia, described as fibrous or “cotton candylike,” with vigorous growth characteristics responsible for the group designation as “lid lifters” (Ribes et al., 2000). For blood culture, a method for concentrating small numbers of fungi, called saponin lysis and centrifugation, is the quickest procedure available with the greatest yield. A negative culture does not preclude infection, as only 50 to 80% of blood cultures from proven cases of invasive candidiasis are positive (Jones, 1990). Unfortunately, molds are rarely recovered from blood, even in rapidly progressive, fulminating, terminal diseases. Each of the fungi may be differentiated from other mycotic agents on examination of cytological specimens or tissue sections. A positive culture linked to hyphal identification in cytological specimens or tissue sections is considered diagnostic. Zygomycetes in respiratory specimens can be differentiated from the dimorphic mycotic pathogens (i.e., Histoplasma capsulatum, Blastomyces dermatitidis, and Coccidiodes immitis), as they do not produce a yeast phase at this site. The major differentiation must be made between the zygomycetes, Candida spp., and other filamentous fungi. The morphology of the hyphae is most important, with zygomycetes species producing wide-angle branching ribbonlike hyphae (of which each species is differentiated based on the morphology of the individual sporangia), with Aspergillus fumigatus producing septate hyphae, and with Candida spp. producing pseudohyphae and blastoconidia in clinical specimens (Kwon-Chung and Bennett, 1992; Ribes et al., 2000). Although serodiagnostic tests for systemic mycotic diseases in humans and animals have improved over the years, most only help to corroborate physical and other diagnostic assessments. The tests with the greatest diagnostic potential are immunodiffusion, complement fixation, antigen titers, and detection of metabolic by-products.
Stenella coeruleoalba (striped dolphin) Tursiops truncatus (bottlenose dolphin)
Arctocephalus gazella (Antarctic fur seal) Callorhinus ursinus (northern fur seal) Cephalorhynchus commersonii Commerson’s dolphin Delphinapterus leucas (beluga) Lissodelphis borealis (northern right whale dolphin) Mesoplodon carlhubbsi (Hubbs’ beaked whale) Phoca vitulina (harbor seal) Orcinus orca (killer whale)
Host Species
Histopathology and culture Histopathology and culture Histopathology and culture Histopathology and culture Histopathology and culture Histopathology and a culture (2) Histopathology and culture Biopsy with histopathology and culture Histopathology and culture (3) Bronchoscopy, culture, and immunodiffusion serology (3)
Lungs
Periauricular and cerebrum Lungs
Lungs
Lung and endobronchial lesions
Lungs and brain
Vertebral bone marrow
Tongue and lungs
Brain
2.5 mg/kg itraconazole BID (approx. 1 year)
Necropsy (morbillivirus positive)
Mycotic Diseases
(Continued)
Guillot et al., 1998; Calle, pers. comm.; Lacave, pers. comm.; Jensen et al., 1998a; Muller, 1994; Bossart and Ewing, pers. comm.
Domingo et al., 1992
Sweeney, pers. comm.
Bossart and Ewing, pers. comm.; McBain and Reidarson, pers. obs. McBain and Reidarson, pers. obs.
Necropsy 1.25 mg/kg itraconazole BID (2 wk) (unsuccessful) Not treated
Lipscomb, pers. comm.
Gage and Lowenstine, pers. comm.
Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm. Joseph et al., 1986
Necropsy
Necropsy
Necropsy
Necropsy
Joseph et al., 1986
Gulland and Haulena, pers. comm.
Necropsy Necropsy
Eyras, pers. comm.
Reference
Necropsy
Aspergillus fumigatus Histopathology and culture Culture
Lungs
Treatment(s)/Comment(s)
OPPORTUNISTIC FUNGI
Diagnostic Method(s)
Lungs, pericardium, lymph nodes Flipper lesions
Lesion(s)
TABLE 1 Mycotic Infections in Marine Mammals
0839_frame_C17.fm Page 341 Tuesday, May 22, 2001 11:12 AM
341
Skin scraping and culture Histopathology and culture Endoscopy, cytology, and immunodiffusion (3) Culture and cytology (4)
Culture and cytology
Pectoral flipper lesion Disseminated Erosive blowhole lesion Tongue, blowhole
Mouth, bladder, kidneys Lung, endobronchial
Disseminated
Histopathology and culture Histopathology and culture (3)
Histopathology and culture
Bronchoscopy and culture
Necropsy
Necropsy
Necropsy
Treatment(s)/Comment(s)
Necropsy (2) 1 mg/kg itraconazole BID (2 mo) (unsuccessful)
10 mg/kg flucytosine TID (2 wk) 10 mg/kg fluconazole TID (2 wk) 5 mg/kg/day ketoconazole 2.5 mg/kg itraconazole BID and 20 mg/kg flucytosine TID (1 yr) 2.5 mg/kg itraconazole BID (8 mo)
Greenwood and Taylor, pers. comm. Dover, pers. comm.
McBain and Reidarson, pers. obs.
Bigg, pers. comm.; Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm. Lacave, pers. comm. Katsumata, pers. comm. Katsumata, pers. comm. Bossart and Ewing, pers. comm. Reidarson and McBain, pers.obs.
Chen, pers. comm.
Necropsy 1.1 mg/kg fluconazole (21 d) (2) Desensitization (1)
Eyras, pers. comm.
Dunn, pers. comm.
Povidone iodine, miconazole, clotrimazole, chlorhexidine
20 mg/kg ketoconazole BID
Griner, 1992
Lipscomb, pers. comm.; Townsend, pers. comm. Greenwood and Taylor, pers. comm.; Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm.; Lipscomb, pers. comm.
Reference
Necropsy
Candida albicans
Histopathology and culture (3) Histopathology and culture (3) Histopathology and culture (29)
Diagnostic Method(s)
Skin, kidneys, lungs, and myocardium Nail bed and external
Brain, lung, and alimentary Lungs
Lesion(s)
342
Zalophus californianus (California sea lion)
Orcinus orca (killer whale) Phoca vitulina (harbor seal) and Callorhinus ursinus (northern fur seal) Pontoporia blainvillei (franciscana) Pseudorca crassidens (false killer whale) Tursiops truncatus (bottlenose dolphin)
Tursiops truncatus (cont’d) (bottlenose dolphin)
Host Species
TABLE 1 Mycotic Infections in Marine Mammals (continued)
0839_frame_C17.fm Page 342 Tuesday, May 22, 2001 11:12 AM
CRC Handbook of Marine Mammal Medicine
Esophageal erosion
Brain
Lungs
Zalophus californianus (California sea lion)
Phocoena phocoena (harbor porpoise)
Enhydra lutris (sea otter) Lagenorhynchus obliquidens (Pacific white-sided dolphin) Phocoenoides dalli (Dall’s porpoise) Lungs, periuterine lymph nodes
Lungs
Lungs, nodes, spleen
Lungs, kidneys, bladder
Lungs
Blowhole lesion
Disseminated
Brain/cranial sinuses
Blowhole lesion
Tursiops truncatus gilli (bottlenose dolphin)
Kogia breviceps (pygmy sperm whale) Tursiops truncatus (bottlenose dolphin)
Necropsy
Histopathology and culture Histopathology and culture
Histopathology and culture Histopathology and culture
Necropsy
Necropsy
Necropsy
Necropsy
Cryptococcus neoformans
Histopathology and culture
Cladophialophora bantiana
Necropsy
Necropsy
Necropsy
1 mg/kg fluconazole SID (1 mo)
2.5 mg/kg itraconazole (11 d) 5.0 mg/kg ketoconazole (12 d) (unsuccessful) Necropsy
1 mg/kg fluconazole SID (1 mo)
Other Candida Culture and cytology (C. parapsilosis) Histopathology and culture (C. rugosa, C. glabrata) Histopathology and culture (C. glabrata) Culture and cytology (C. tropicalis) Histopathology and culture Histopathology and culture (C. tropicalis) Histopathology and culture
Raverty, pers. comm.
Raverty, pers. comm.
Joseph, pers. comm.
Joseph, pers. comm.
(Continued)
Gage and Lowenstine, pers. comm.
Chen, pers. comm.
Chen, pers. comm.
Joseph, pers. comm.
Stamper and Whitaker, pers. comm.
Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm.; Lipscomb, pers. comm.
Dalton and Robeck, pers. comm.
Stamper and Whitaker, pers. comm.
0839_frame_C17.fm Page 343 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases 343
Skin
Skin
Zalophus californianus (California sea lion)
Zalophus californianus (California sea lion)
Histopathology and culture
Unknown
Guillot et al., 1998
Migaki and Jones, 1983
Abt, pers. comm. Frasca et al., 1996
1 mg/kg/day itraconazole (4 mo) Necropsy Unknown
Gulland and Haulena, pers. comm.
Frasca et al., 1996
Frasca et al., 1996
Migaki and Jones, 1983
Haubold et al., 1997
Murnane and Kinsen, 1998; Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm. Migaki et al., 1978b
Reference
0.5 mg/kg fluconazole BID (3 wk)
Malassezia pachydermatis
Skin scraping and culture Culture Histopathology and culture (2) Culture
All mucocutaneous junctions Nail bed Diffuse cutaneous
5 mg/kg ketoconazole SID (10 d)
Necropsy
Unknown
Fusarium spp.
Necropsy
Deuteromyces
Necropsy
Necropsy
Treatment(s)/Comment(s)
344
Skin
Skin
Histopathology and culture Histopathology and culture Histopathology and culture
Skin
Halichoerus grypus (gray seal) Kogia breviceps (pygmy sperm whale) Lagenorhynchus acutus (Atlantic white-sided dolphin) Mirounga angustirostris (northern elephant seal) Phoca vitulina (harbor seal)
Histopathology
Histopathology and culture
Lung
Brain, trachea, cervical lymph nodes
Histopathology and culture (2)
Diagnostic Method(s)
Disseminated
Lesion(s)
Tursiops truncatus (bottlenose dolphin)
Tursiops truncatus (bottlenose dolphin)
Host Species
TABLE 1 Mycotic Infections in Marine Mammals (continued)
0839_frame_C17.fm Page 344 Tuesday, May 22, 2001 11:12 AM
CRC Handbook of Marine Mammal Medicine
Skin
Skin
Lungs
Disseminated
Lagenorhynchus obliquidens (Pacific white-sided dolphin)
Callorhinus ursinus (northern fur seal)
Tursiops truncatus (bottlenose dolphin)
Tursiops truncatus (bottlenose dolphin)
Tursiops truncatus (bottlenose dolphin)
Disseminated
Mirounga angustirostris (northern elephant seal)
Subcutaneous fat, muscle, and lung
Blubber, muscle, lungs Skin and brain
Skin
Phoca vitulina (harbor seal)
None
Trichophyton sp.
Necropsy
Bronchoscopy and culture (2) Histopathology and culture Histopathology and culture Histopathology and culture
Robeck et al., in prep.
Townsend, pers. comm.
Skin biopsy, culture, and necropsy Necropsy
Robeck and Dalton, in prep.
Townsend et al., 1996
(Continued)
McBain and Reidarson, pers. comm.
Vedros et al., 1982
Migaki et al., 1978a
Haulena et al., 2000
Fansworth et al., 1975
15–18 mg/kg cum. dose Amphocil IV (12–14 d) (unsuccessful) 2.4–4.2 mg/kg Nyotran IV SID
ZYGOMYCETES Apophysomyces elegans
2.5 mg/kg itraconazole and 20 mg/kg flucytosine TID
Trichosporon pullulans Bronchoscopy and culture
Culture
Histopathology and culture
Sporothrix schenckii
Necropsy
Scedosporium apiospermum
Unknown
Microsporum canis
Histopathology and culture
Culture
0839_frame_C17.fm Page 345 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases 345
Disseminated
Cystophora cristata (hooded seal) Eubalaena australis (southern right whale) Kogia breviceps (pygmy sperm whale) Lagenorhynchus obliquidens (Pacific white-sided dolphin) Orcinus orca (killer whale) Lung, gastrointestinal, heart, and skeletal muscle
Brain
Pyloric stomach and local lymph nodes Disseminated
Necropsy
1 mg/kg itraconazole BID (3 d) (unsuccessful)
Histopathology and culture (2) Histopathology and culture
Histopathology and culture Histopathology and culture Histopathology and culture Histopathology and culture
Necropsy
Robeck et al., in prep.
Bossart and Ewing, pers. comm.
Robeck et al., in prep.
2.5 mg/kg itraconazole (23 d) (unsuccessful) Necropsy
Bossart and Ewing, pers. comm.
Best and McCully, 1979
Bossart, and Ewing, pers. comm.
Robeck et al., in prep.
Wünschmann et al., 1999
Necropsy
Necropsy
Necropsy
Other Zygomycetes
Histopathology and culture
Saksenaea vasiformis
Histopathology and culture
Robeck et al., in prep.
Lipscomb, pers. comm.
Reference
346
Skeletal muscle
Lung, uterus, and brain
Orcinus orca (killer whale)
2.5 mg/kg itraconazole (7 d) (unsuccessful)
Necropsy
Treatment(s)/Comment(s)
Rhizomucor pusillus
Histopathology and culture
Disseminated
Brain, lung, kidneys, testis, lymph nodes
Histopathology and culture
Diagnostic Method(s)
Brain
Lesion(s)
Phocoena phocoena (harbor porpoise)
Lagenorhynchus acutus (Atlantic white-sided dolphin) Lagenorhynchus obliquidens (Pacific white-sided dolphin)
Host Species
TABLE 1 Mycotic Infections in Marine Mammals (continued)
0839_frame_C17.fm Page 346 Tuesday, May 22, 2001 11:12 AM
CRC Handbook of Marine Mammal Medicine
Enhydra lutris (sea otter) Tursiops truncatus gilli (bottlenose dolphin) Zalophus californianus (California sea lion)
Eumetopias jubatus (Steller sea lion) Tursiops truncatus (bottlenose dolphin) Zalophus californianus (California sea lion)
Tursiops truncatus (bottlenose dolphin)
Pagophilus groenlandicus (harp seal) Phocoenoides dalli (Dall’s porpoise)
Lung, liver, pulmonary lymph nodes
Disseminated
Disseminated
Disseminated
Disseminated
Disseminated
Skin and brain
Lungs, mandible
Hilar lymph node
Disseminated
Disseminated
Disseminated
Lung
2 mg/kg Amphotec IV for two separate treatments (unsuccessful) 2.5 mg/kg itraconazole (9 mo) (unsuccessful)
Necropsy
Necropsy
Necropsy
Necropsy
Necropsy
Necropsy
Necropsy
Necropsy
Histopathology, culture, and serology (7) histopathology, culture, and serology Histopathology and culture (9)
Necropsy
Necropsy
Necropsy
Coccidioides immitis
Histopathology and culture Histopathology and culture histopathology and culture
ENDEMIC FUNGI Blastomyces dermatitidis
Histopathology and culture (Paecilomyces lilacinus)
Histopathology and culture Histopathology and culture Histopathology and culture Histopathology and culture (7) Histopathology and culture Biopsy and culture
(Continued)
Fauquier et al., 1996; Reed et al., 1972; Williams, pers. comm.
Thomas et al., 1994; Williams, pers. comm.; Joseph, pers. comm. Reidarson et al., 1998
Kapustin, pers. comm.
Cates et al., 1986
Williamson et al., 1959
Sweeney et al., 1976; Mathey, pers. comm.
Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm.; Townsend, pers. comm. Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm. Walsh, pers. comm.
Young, pers. comm.
Sweeney et al., 1976
Kaplan et al., 1960
0839_frame_C17.fm Page 347 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases 347
Number of cases in parentheses.
a
Histopathology and culture Histopathology and culture (3)
Histopathology and culture Histopathology and culture Histopathology Histopathology and culture
Disseminated
Pulmonary, meningeal Disseminated Necropsy Necropsy
Necropsy
Necropsy
Necropsy
Necropsy
Histopathology Histopathology and electron microscopy (11)
Cutaneous (melon, pectoral flipper, or disseminated)
10–16 mg/kg/day ketoconazole 0.5 mg/kg fluconazole BID 1.0 mg/kg miconazole 2.5 mg/kg itraconazole BID
Necropsy
Lacazia loboi Cutaneous
Lung and skin Lung
Treatment(s)/Comment(s)
Histoplasma capsulatum
Diagnostic Method(s)
Gastrointestinal
Lesion(s)
Haubold et al., 1997; Bossart, 1984; Caldwell and Caldwell, 1975; Dudok, 1977; Migaki and Valerio, 1971; Poelma et al., 1974; Symmers, 1983; Bossart and Ewing, pers. comm.; Cowan, pers. comm.; Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm.; Townsend, pers. comm.; Schroeder, pers. comm.
De Vries and Laarman, 1973
Lipscomb, pers. comm. Greenwood and Taylor, pers. comm.
Wilson et al., 1974
Jensen et al., 1998b; Bossart and Ewing, pers. comm.; Jensen, Linnehan, Miller, Ridgway, and Van Bonn, pers. comm.; Townsend, pers. comm. Dalton and Robeck, pers. comm.
Dalton and Robeck, pers. comm.
Reference
348
Sotalia fluviatilis guianensis (tucuxi) Tursiops truncatus (bottlenose dolphin)
Pseudorca crassidens (false killer whale)
Pagophilus groenlandicus (harp seal)
Lagenorhychus obliquidens (Pacific white-sided dolphin) Tursiops truncatus (bottlenose dolphin)
Host species
TABLE 1 Mycotic Infections in Marine Mammals (continued)
0839_frame_C17.fm Page 348 Tuesday, May 22, 2001 11:12 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C17.fm Page 349 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases
349
Immunodiffusion (ID) assays detect circulating antibodies, which are helpful in assessing exposure to a fungus. Qualitative ID measures the presence of different antibody classes (either IgM or IgG), and is reported by the number of antibody–antigen bands in a gel (or bands of identity). Quantitative tests determine the antibody titer for a specific fungal antigen. Qualitative tests may be useful with candidiasis where a change in bands of identity, from zero to one or more, is indicative of active infection, although not all investigators feel such tests are useful. Quantitative ID is most useful in coccidioidomycosis in which a fourfold rise in titer over 30 days indicates an active infection, and a titer of greater than 1:128 is associated with extrapulmonary dissemination (Christin and Sugar, 1996). There has been considerable progress in the development of antigen-specific tests for mycotic infections. The tests with the greatest potential to detect systemic candidiasis are ones designed to detect metabolic by-products or secreted cell wall constituents. These materials are secreted unpredictably and in minute concentrations, however, so the tests are very unreliable (Jones, 1990; Na and Song, 1999). In contrast, tests for wall constituents have been developed for A. fumigatus, B. dermatitidis, Cryptococcus neoformans, and H. capsulatum (Williamson et al., 1959; Walsh and Mitchell, 1991; Kwon-Chung and Bennett, 1992; Soufleris et al., 1994; Christin and Sugar, 1996; Jensen et al., 1998a,b; Maesaki et al., 1999; Chumpitazi et al., 2000; Hurst et al., 2000). Although serological diagnostic tests for detecting zygomycosis are rarely clinically useful, enzyme-linked immunosorbent assays (ELISA) seem to have the best sensitivity and specificity for detecting antibodies produced during invasive zygomycosis (Kwon-Chung and Bennett, 1992; Ribes et al., 2000). With the exception of the A. fumigatus competitive binding inhibition assay, cryptococcal latex agglutination, the ELISA antigen method for histoplasmosis, and the diagnostically and prognostically useful serological tests for coccidioidomycosis, the utility of serology for the diagnosis of mycoses is generally dismal.
Therapeutics Various antifungal drugs have been used to treat mycotic infections in marine animals (see Table 2). The azoles fluconazole (Diflucan®, Pfizer, New York, NY), itraconazole (Sporanox®, Janssen Pharmaceuticals Co., Princeton, NJ), and ketoconazole (Nizoral®, Janssen Pharmaceuticals Co., Princeton, NJ), a microencapsulated colloidal dispersion of amphotericin B called Amphotec® (Sequus Pharmaceutical, Inc., Menlo Park, CA), an experimental liposomal form of nystatin called Nyotran® (Aronex Pharmaceuticals, Inc., The Woodlands, TX), flucytosine (Ancobon®, ICN Pharmaceuticals, Inc., Costa Mesa, CA) and the combination of itraconazole and flucytosine have been used with variable success (Allendoerfer et al., 1991; Presterl and Graninger, 1994; Reidarson and McBain, 1995). Because fluconazole is water soluble and excreted in urine, it may be used in cases of renal or urinary candidiasis, as well as systemic disease. Although itraconazole is effective for candidiasis, it has been used mainly to treat A. fumigatus infections (Stevens and Lee, 1997). To date, only the microencapsulated nystatin and the microencapsulated amphotericin B products have shown limited, but in some instances encouraging, efficacy against the zygomycetes in humans and marine mammals (Herbrecht, 1996; Townsend et al., 1996). Experience with intravenously administered antifungal drugs (polyenes and their lipid formulations) is limited because of their adverse renal effects, and technical difficulties associated with intravenous (IV) administration in marine mammals (Townsend et al., 1996; Robeck and Dalton, in prep.). In one instance, however, short-term remission was achieved with IV administration of Nyotran for systemic Aphophysomyces elegans infection in a bottlenose dolphin (Robeck and Dalton, in prep.). The key to successful treatment of zygomycoses rests in early diagnosis, radical resection of affected tissues, and long-term IV therapy.
b
Dosages are in mg/kg; N.D. = not done. Used with low-dose prednisolone (0.01 mg/kg SID).
N.D. N.D. 1.25 BID N.D. N.D.
N.D.
Orcinus orca (killer whale) 1.0–2.0 IV SID (2.5 g cum. dose) 20 TID 2.0 BID 2.5–5.0 BID 5.0 BID 2.4–4.2 IV SID
Tursiops truncatus (bottlenose dolphin)
N.D. N.D. 5.0 BID N.D. N.D.
N.D.
Cephalorhynchus commersonii (Commerson’s dolphin)
a
N.D. N.D. 1.25 BID N.D. N.D.
N.D.
Globicephala macrorhynchus (pilot whale)
N.D. N.D. 2.5 BID 1.9 BID N.D.
N.D.
Delphinapterus leucas (beluga)
350
a
Lipid formulation Amphotericin B Flucytosine Fluconazole Itraconazole b Ketoconazole Nystatin
Drug
TABLE 2 Formulary of Antimycotic Drugs Used in Certain Marine Mammal Species
0839_frame_C17.fm Page 350 Tuesday, May 22, 2001 11:12 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C17.fm Page 351 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases
351
Because itraconazole is lipophilic, tissue levels are generally higher than blood levels. Because of the slow release of itraconazole, significant blood levels in marine mammals can persist for weeks after discontinuing therapy. The levels and length of release appear to be dependent on the amount of blubber; however, there is some individual variation in pharmacokinetics (Dalton and McBain, unpubl. data). Inappetence occasionally occurs with the administration of either ketoconazole or itraconazole. Since ketoconazole appears to inhibit glucocorticoid production, supplementation with 0.01 mg/kg prednisolone is recommended. For itraconazole, reducing the dose generally reverses the inappetence. Each of the azoles (except generally fluconazole) is capable of producing 2- to 25-fold elevations of the liver-associated enzymes, asparate and alanine transaminases and lactate dehydrogenase (LDH) (Reidarson and McBain, 1994). The patterns of LDH isoenzyme production help distinguish between enzyme induction due to azole administration and liver pathology. There appears to be no liver pathology associated with itraconazole and fluconazole, in contrast to ketoconazole (Reidarson and McBain, 1994). Furthermore, as long as drug levels persist, enzyme levels also remain elevated. The authors’ experience in the treatment of invasive mycoses in marine mammals has shown that mycotic infections often persist if only the treatment regimes outlined in the medical literature are followed. Treatment must be extended well beyond the apparent elimination of the mycotic infection, which can only be determined by clinical reassessment of each animal. Clinical signs, cultures, and follow-up diagnostic tests assist in making the most informed decision concerning the discontinuation of therapy. Treatment of invasive mycotic infections must continue past an apparent cure as indicated by a clinical, radiological, or endoscopic disease-free state. As fungi have developed resistance to currently accepted pharmaceuticals, newer antifungals have emerged. These include the azoles voriconazole (Pfizer Pharmaceuticals, New York, NY) and posaconazole (Schering Pharmaceuticals, Kenilworth, NJ), and the new squalene epoxidase inhibitor terbinafin (Lamisil®, Novartes Pharmaceuticals Corp., East Hanover, NJ), each of which offers an expanded spectrum of activity and is fungicidal rather than fungistatic at concentrations used in humans (Kappe et al., 1998; De Pauw et al., 1999; Hay, 1999; Patterson, 1999; Xu et al., 2000; Luna et al., 2000;). Recent studies have shown the efficacy of double, and even triple, antifungal therapy in humans with systemic cryptococcosis, candidiasis, and coccidioidomycosis (Kappe et al., 1998). Several mycotic clinical laboratories offer in vitro combination tests that have demonstrated synergy in many instances in which combinations of fungistatic drugs are applied. For individuals on combination therapy containing flucytosine, both medications should be withdrawn simultaneously to prevent the development of resistance. This phenomenon is best illustrated with azoles and flucytosine, when withdrawal of the azole may lead to resistance to flucytosine (Nguyen et al., 1996). Flucytosine is never used as monotherapy because of the rapid generation of resistance.
Conclusion Most mycotic infections share clinical, hematological, and serum biochemical aspects of bacterial and viral infections. Clinical presentations are frustratingly nonspecific, ranging from chronic to fulminating, while laboratory findings simply demonstrate acute or chronic inflammation, just as with bacterial or viral diseases. Although serodiagnostic tests for systemic fungi have improved, most only help to corroborate physical and other diagnostic assessments. It is for this reason that biopsy and culture are the most definitive methods for diagnosing mycotic infections, except for those by Lacazia loboi which has never been successfully cultured.
0839_frame_C17.fm Page 352 Tuesday, May 22, 2001 11:12 AM
352
CRC Handbook of Marine Mammal Medicine
A number of new antimycotic drugs have been introduced. These include itraconazole, fluconazole, and terbinafin (with newer azoles, voriconazole, and posaconazole to be available soon), which appear to be somewhat effective against Aspergillus fumigatus, Candida spp., and the endemic fungi, and the microencapsulated lipid-bound nystatin and amphotericin B products that have limited efficacy against zygomycosis. For the emerging drug-resistant fungi, combinations of various pharmaceuticals may be necessary, and for all mycotic infections, longterm therapy is vital for a successful outcome. Diagnostic modalities have greatly lagged behind therapeutics. However, with improvements in serodiagnostic tests for metabolic by-products and cell wall constituents, as well as emerging methods involving molecular biologic techniques, the ability to make timely ante-mortem diagnoses is expected to improve greatly.
Acknowledgments The authors thank Elsa Haubold for reviewing this chapter, and Rebecca Duerr for editorial assistance. This is SeaWorld of California technical contribution 2000-05-C.
References Allendoerfer, R., Marquis, A.J., Rinaldi, M.G., and Graybill, J.R., 1991, Combined therapy with fluconazole and flucytosine in murine cryptococcal meningitis, Antimicrob. Agents Chemother., 35: 726–729. Best, P.B., and McCully, R.M., 1979, Zygomycosis (phycomycosis) in a right whale (Eubalaena australis), J. Comp. Pathol., 89: 341–348. Bossart, G.D., 1984, Suspected acquired immunodeficiency in an Atlantic bottlenose dolphin with chronic-active hepatitis and lobomycosis, J. Am. Med. Assoc., 185: 1413–1414. Caldwell, D.K., and Caldwell, M.C., 1975, Lobomycosis as a disease of the Atlantic bottlenosed dolphin, Tursiops truncatus, Montague (1821), Am. J. Trop. Med. Hyg., 24: 105–114. Cates, M.B., Kaufman, J.H., Pletcher, J., and Schroeder, J.P., 1986, Blastomycosis in an Atlantic bottlenose dolphin, J. Am.Vet. Med. Assoc., 189: 1148–1150. Christin, L., and Sugar, A.M., 1996, Endemic fungal infections in patients with cancer, Infect. Med., 13: 673–679. Chumpitazi, B.F., Pinel, C., Lebeau, B., Ambroise-Thomas, P., and Grillot, R., 2000, Aspergillus fumigatus antigen detection in sera from patients at risk for invasive aspergillosis, J. Clin. Microbiol., 38: 438–443. Coleman, J.M., Hogg, G.G., Rosenfeld, J.V., and Waters, K.D., 1995, Invasive central nervous system aspergillosis: Cure with liposomal amphotericin B, itraconazole, and radical surgery—Case report and review of the literature, Neurosurgery, 36: 858–860. De Pauw, B.E., Donnelly, J.P., and Kullberg, B.J., 1999, Treatment of fungal infections in surgical patients using conventional antifungals, J. Chemother., 11: 494–503. de Vries, G.A., and Laarman, J.J., 1973, A case of Lobo’s disease in the dolphin Sotalia guianensis, Aquat. Mammals, 1: 26–33. Domingo, M., Visa, J., and Pumarola, M., 1992, Pathologic and immunocytochemical studies of morbillivirus infection in striped dolphins (Stenella coeruleoalba), Vet. Pathol., 29: 1–10. Dudok, W.H., 1977, Successful treatment in a case of lobomycosis (Lobo’s disease) in Tursiops truncatus (Mont.) at the Dolfinarium, Harderwijk, Aquat. Mammals, 5: 8–15. Fansworth, R.J., McKeever, P.J., and Flether, J.A., 1975, Dermatopmycosis in a harbor seal caused by Microsporum canis, J. Zoo. Anim. Med., 6: 26–28.
0839_frame_C17.fm Page 353 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases
353
Fauquier, D.A., Gulland, F.M., Trupkiewicz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996, Coccidioidomycosis in free-living California sea lions (Zalophus californianus) in central California, J. Wildl. Dis., 32: 707–710. Frasca, J., Dunn, J.L., Cooke, J.C., and Buck, J.D., 1996, Mycotic dermatitis in an Atlantic white-sided dolphin, a pygmy sperm whale, and two harbor seals, J. Am. Vet. Med. Assoc., 208: 727–729. Geraci, J.R., and Ridgway, S.H., 1991, On disease transmission between cetaceans and humans, Mar. Mammal Sci., 7: 191–194. Griner, L.A., 1992, Cardiac candidiasis in a captive killer whale, in Erkrankungen der Zootiere, 34, Internationalen Symposiums über die Erkrankungen der Zoo- und Wildtiere, Santander, Spain, 159–162. Guillot, J., Petit, T., Degoree-Rubiales, F., Gueho, E., and Charmette, R., 1998, Dermatitis caused by Malassezia pachydermatis in a California sea lion (Zalophus californianus), Vet. Rec., 142: 311–312. Haque, A.K., 1992, Pathology of common pulmonary fungal infections, J. Thorac. Imaging, 7: 1–11. Harley, W.B., and Blaser, M.J., 1994, Disseminated coccidioidomycosis associated with extreme eosinophilia, Clin. Infect. Dis., 18: 627–629. Harrell, J.H., Reidarson, T.H., McBain, J., and Sheetz, H., 1996, Bronchoscopy of the bottlenose dolphin, Abstr., 27th Annual International Association for Aquatic Animal Medicine Conference, Chattanooga, TN, 33. Haubold, E.M., Cowan, D.F., Cooper, C.R., and McGinnis, M.R., 1997, Unusual mycotic infections in stranded western Gulf of Mexico dolphins: Phaeohyphomycosis and lobomycosis, Abstr., 28th Annual International Association for Aquatic Animal Medicine Conference, Harderwijk, the Netherlands, 26–27. Hawkins, E.C., Townsend, F.I., Lewbart, G.A., Stamper, M.A., Thayer, V.G., and Rhinehart, H.L., 1996, Bronchoalveolar lavage in a stranded bottlenose dolphin, Abstr., 27th Annual International Association for Aquatic Animal Medicine Conference, Chattanooga, TN, 124. Hay, R.J., 1999, Therapeutic potential of terbinafin in subcutaneous and systemic mycoses, Br. J. Dermatol., 141(Suppl. 56): 36–40. Herbrecht, R., 1996, The changing epidemiology of fungal infections: Are the lipid forms of amphotericin B an advance? Eur. J. Haematol., 57: 12–17. Hurst, S.F., Reyes, G.H., McLaughlin, D.W., Reiss, E., and Morrison, C.J., 2000, Comparison of commercial latex agglutination and sandwich enzyme immunoassays with a competitive binding inhibition enzyme immunoassay for detection of antigenemia and antigenuria in a rabbit model of invasive aspergillosis, Clin. Diagn. Lab. Immunol., 7: 477–485. Jensen, E.D., Lipscomb, T., Van Bonn, B., Miller, G., Fradkin, J.M., and Ridgway, S.H., 1998a, Disseminated histoplasmosis in an Atlantic bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 456–460. Jensen, E.D., Van Bonn, W., Lipscomb, T., and Ridgway, S.H., 1998b, Disseminated histoplasmosis in Atlantic bottlenose dolphins, Abstr., 29th Annual International Association for Aquatic Animal Medicine Conference, San Diego, CA, 64. Jeraj, K.P., and Sweeney, J.C., 1996, Blowhole cytology to diagnose early respiratory tract disease in bottlenose dolphins, Abstr., 27th Annual International Association for Aquatic Animal Medicine Conference, Chattanooga, TN, 112. Jones, J.M., 1990, Laboratory diagnosis of invasive candidiasis, Clin. Microbiol. Rev., 3: 32–45. 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. Kaplan, W., Goss, L.F., Ajello, L., and Ives, M.S., 1960, Pulmonary mucomycosis in a harp seal caused by Mucor pusillus, Mycopathol. Mycol. Appl., 12: 101–102. Kappe, R., Levitz, S., Harrison, T.S., Ruhnke, M., Ampel, N.M., and Just-Nubling, G., 1998, Recent advances in cryptococcosis, candidiasis, and coccidioidomycosis complicating HIV infection, Med. Mycol., 36(Suppl. 1): 207–215. Kwon-Chung, K.J., and Bennett, J.E., 1992, Blastomycosis, in Medical Mycology, Kwon-Chung, K.J., and Bennett, J.E. (Eds.), Lea & Febiger, Philadelphia, 248–279.
0839_frame_C17.fm Page 354 Tuesday, May 22, 2001 11:12 AM
354
CRC Handbook of Marine Mammal Medicine
Luna, B., Drew, R.H., and Perfect, J.R., 2000, Agents for treatment of invasive fungal infections, Otolaryngol. Clin. North Am., 33: 277–299. Maesaki, S., Kawamura, S., Hashiguchi, K., Hossain, M.A., Sasaki, E., Miyazaki, Y., Tomono, K., Tashiro, T., and Kohno, S., 1999, Evaluation of sandwich ELISA galactomannan test in samples of positive LA test and positive aspergillus antibody, Intern. Med., 38: 948–950. Magnussen, C.R., 1992, Disseminated Candida infection: Diagnostic clues, therapeutic options, J. Crit. Illness, 7: 513–522. McAdams, H.P., Rosado-de-Christenson, M., and Templeton, P.A., 1995, Thoracic mycoses from opportunistic fungi: Radiologic-pathologic correlation, Radiographics, 15: 271–286. Migaki, G., and Jones, S.R., 1983, Mycotic diseases in marine mammals, in Pathobiology of Marine Mammal Diseases, Vol. II, Howard, E.B. (Ed.) CRC Press, Boca Raton, FL, 1–25. Migaki, G., and Valerio, M.G., 1971, Lobo’s disease in an Atlantic bottlenose dolphin, J. Am. Vet. Med. Assoc., 159: 578–582. Migaki, G., Font, R.L., Kapanand, W., and Asper, E.D., 1978a, Sporotrichosis in a Pacific white-sided dolphin (Lagenorhynchus obliquidens), Am. J. Vet. Res., 39: 1916–1919. Migaki, G., Gunnels, R.D., and Casey, C.W., 1978b, Pulmonary cryptococcosis in an Atlantic bottlenose dolphin (Tursiops truncatus), Lab. Anim. Sci., 28: 603–606. Muller, J., 1994, Epidemiology of deep-seated, domestic mycoses, Mycoses, 37(Suppl 2): 1–7. Murnane, R.D., and Kinsen, M.J., 1998, Subacute and fulminating pulmonary cryptococcosis with dissemination in an Atlantic bottlenosed dolphin, Abstr., 29th Annual International Association for Aquatic Animal Medicine Conference, San Diego, CA, 43–45. Na, B.K., and Song, C.Y., 1999, Use of monoclonal antibody in diagnosis of candidiasis caused by Candida albicans: Detection of circulating aspartyl proteinase antigen, Clin. Diagn. Lab. Immunol., 6: 924–929. Nguyen, M.H., Peacock, J.E., and Morris, A.J., 1996, The changing face of candidemia: Emergence of non-Candida albicans species and antifungal resistance, Am. J. Med., 100: 617–623. Nicholls, J.M., Yuen, K.Y., and Tam, A.Y.C., 1993, Systemic fungal infections in neonates, J. Hosp. Med., 49: 420–427. Patterson, T.F., 1999, Role of newer azoles in surgical patients, J. Chemother., 11: 504–512. Poelma, F.G., de Vries, G.A., Blythe-Russell, E.A., and Luykx, H.F., 1974, Lobomycosis in an Atlantic bottlenosed dolphin in the Dolphinarium Harderwijk, Aquat. Mammals, 2: 11–15. Presterl, D., and Graninger, W., 1994, Efficacy and safety of fluconazole in the treatment of systemic fungal infections in pediatric patients. Multicentre study group, Eur. J. Clin. Microbiol. Infect. Dis., 13: 347–351. Reed, R.E., Migaki, G., and Cummings, J.A., 1972, Coccidioidomycosis in a California sea lion (Zalophus californianus), J. Wildl. Dis., 12: 372–375. Reef, V.B., 1991, Ultrasonographic evaluation, in Equine Respiratory Diseases, Beech, J. (Ed.), Lea & Febiger, Philadelphia, 69–79. Reidarson, T.H., and McBain, J., 1994, The use of LDH isoenzymes to differentiate three medical conditions in cetaceans, Abstr., 25th Annual International Association for Aquatic Animal Medicine Conference, Mystic, CT, 156. Reidarson, T.H., and McBain, J., 1995, The combined use of itraconazole and flucytosine in the treatment of chronic candida cystitis in a bottlenose dolphin (Tursiops truncatus), Abstr., 26th Annual International Association for Aquatic Animal Medicine Conference, Mystic, CT, 13. 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., 27th Annual International Association for Aquatic Animal Medicine Conference, Chattanooga, TN, 34. Reidarson, T.H., Griner, L., and McBain, J., 1998, Coccidioidomycosis in a bottlenose dolphin, J. Wildl. Dis., 34: 629–631. Reidarson, T.H., McBain, J.F., Dalton, L.M., and Rinaldi, M.G., 1999, Diagnosis and treatment of fungal infections in marine mammals, in Zoo and Wild Animal Medicine, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 478–485.
0839_frame_C17.fm Page 355 Tuesday, May 22, 2001 11:12 AM
Mycotic Diseases
355
Ribes, J.A., Vanover-Sams, C.L., and Baker, D.J., 2000, Zygomycetes in human disease, Clin. Microbiol. Rev., 13: 236–301. Rinaldi, M., 1989, Zygomycosis, Infect. Dis. Clin. North Am., 3: 19–41. Rippon, J.W., 1988, The pathogenic fungi and the pathogenic actinomycetes, in Medical Mycology, Vol. 3, W.B. Saunders, Philadelphia, 618–650. Robeck, T., and Dalton, L., in preparation, Treatment of a cutaneous, subcutaneous Apophysomyces elegans, a mucormycotic fungi, infection in a bottlenose dolphin (Tursiops truncatus) with the new antifungal agent, Nyotran. Robeck, T., Dalton, L., and Rinaldi, M., in preparation, 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. Soufleris, A.J., Klein, B.S., and Courtney, B.T., 1994, Utility of anti-WI-1 serological testing in the diagnosis of blastomycosis in Wisconsin residents, Clin. Infect. Dis., 19: 89–92. Stevens, D.A., and Lee, J.Y., 1997, Analysis of compassionate use itraconazole therapy for invasive aspergillosis by the NIAID Mycoses Study Group criteria, Arch. Intern. Med., 157: 1857–1862. 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. Symmers, W., 1983, A possible case of Lobo’s disease acquired in Europe from a bottlenose dolphin (Tursiops truncatus), Bull. Soc. Pathol. Exot., 76: 777–779. Taborda, P.R.O., Taborda, V.B.A., and McGinnis, M.R., 1999, Lacazia loboi gen. nov. comb. nov., the etiologic agent of lobomycosis, J. Clin. Microbiol., 37: 2031–2033. Thomas, N.J., Pappagianis, D., Creekmore, L.H., and Duncan, R.M., 1994, Coccidioidomycosis in southern sea otters, in Centennial Conference on Coccidioidomycosis, University of California, San Diego, 21–22. Townsend, F.I., Materese, F.J., and Sips, D.G., 1996, The use of liposomal amphotericin-B in the therapy of systemic zygomycosis, Abstr., 27th Annual International Association for Aquatic Animal Medicine Conference, Chattanooga, TN, 18. Vedros, N.A., Quinlivan, J., and Cranford, R., 1982, Bacterial and fungal flora of wild northern fur seals (Callorhinus ursinus), J. Wildl. Dis., 18: 447–456. Walsh, T.J., and Mitchell, T.G., 1991, Dimorphic fungi causing systemic mycoses, in Manual of Clinical Microbiology, Balows, A., Hausler, W.J., Herrmann, K.L. et al. (Eds.), American Society for Microbiology, Washington, D.C., 630–643. Wheat, J., 1996, Histoplasmosis in the acquired immuno deficiency syndrome, Curr. Top. Med. Mycol., 7: 7–18. Williamson, W.M., Lombard, L.S., and Getty, R.E., 1959, North American blastomycosis in a northern sea lion, J. Am. Vet. Med. Assoc., 135: 513–520. Wilson, T.M., Kierstead, M., and Long, J.R., 1974, Histoplasmosis in a harp seal, J. Am. Vet. Med. Assoc., 165: 815–817. Wünschmann, A., Siebert, U., and Weiss, R., 1999, Rhizopusmycosis in a harbor porpoise from the Baltic Sea, J. Wildl. Dis., 35: 569–573. Xu, J., Ramos, A.R., Vilgalys, R., and Mitchell, T.G., 2000, Clonal and spontaneous origins of fluconazole resistance in Candida albicans, J. Clin. Microbiol., 38: 1214–1220.
0839_frame_C17.fm Page 356 Tuesday, May 22, 2001 11:12 AM
0839_frame_C18 Page 357 Tuesday, May 22, 2001 11:13 AM
18 Parasitic Diseases Murray D. Dailey
Introduction Parasites have been known to cause major health problems in marine mammals since research scientists and veterinarians first began study of these animals. Researchers and veterinarians need to know: (1) how to remove and fix parasites for identification; (2) how to identify parasites; (3) which parasites affect the health of the animal; and (4) the best method to treat the marine mammal. This chapter will provide this information, host–parasite lists from each of the marine mammal groups, and resources to assist with identification. Because of the large number of parasites that have been reported from marine mammals, it is impossible to cover each parasite in detail here. Instead, this chapter will give new material and important examples of host– parasite associations, in addition to clinical signs, diagnosis, and treatment. The majority of life-cycles of marine mammal parasites—other than Filaroides (Parafilaroides) decorus, Otostrongylus circumlitus, and Uncinaria lucasi—have not been determined experimentally; thus, they will not be discussed here. This chapter is not meant to be a complete diagnostic field manual (the reader is referred below to references for identification purposes). It will, however, serve as a pragmatic guide to parasitic diseases of marine mammals.
Removal and Fixation of Parasites for Identification Parasites collected during necropsies must be handled correctly for identification and subsequent utilization in retrospective studies. A wealth of information about parasites can be gleaned if just minimal attention is paid to the preparation of specimens, whereas poor collection or fixation may make identification impossible. Preparation of parasitic organisms involves five stages: (1) removal from the host during necropsy, (2) killing, (3) fixing, (4) staining/ dehydrating, and (5) mounting. Complete field data, including host identification and collection sites, are also absolutely essential. To conduct parasitological analyses during necropsies of marine mammals, first perform an external examination for ectoparasites, including those found in the blowhole, nares, and ears, prior to opening the carcass. Pick up large parasites using forceps. If the parasites are small, use a toothpick or camel’s hair brush previously dipped in alcohol to collect the parasites; then transfer them to a vial containing preserving fluid. Swab the blowhole or nares, ears, and oral cavity, and store the swabs in vials containing preservatives or smear them onto glass slides. Collect gastrointestinal parasites by opening the stomach and intestines, then washing and gently scraping the mucosal surfaces in saline and placing the fluid in a container. This solution is
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
357
0839_frame_C18 Page 358 Tuesday, May 22, 2001 11:13 AM
358
CRC Handbook of Marine Mammal Medicine
then rinsed through a sieve with a mesh size of 150 µm to retain all macroparasites. Failure to scrape the mucosa gently (using gloved fingers or a tongue depressor) during washing may result in failure to collect smaller or more adherent parasites. If the infection is heavy, a representative subsample may be taken. To identify cestodes, a scolex (head), mature (middle), and gravid (end) segments are needed, as well as the approximate total length of the cestode. Parasites collected from the respiratory parenchyma must be teased free without breaking them. If only pieces can be collected, be sure to get the head and tail for identification. The auditory canals and sinuses of cetaceans undergoing necropsy must be opened and flushed for flukes of the genus Nasitrema and numerous genera of nematodes such as Stenurus. Fix protozoan parasites in Bouin’s fluid and store at 4°C (40°F); potassium dichromate in a 5% aqueous solution is best for Coccidia oocysts, as it allows them to sporulate. De-hemoglobinize (immerse in distilled water for 20 min) blood smears that may contain protozoa, and fix in 70% ethanol. Air-dry blowhole samples taken on a glass slide; then dip in absolute methanol for 1 min to fix, air-dry again, dip in 10% neutral buffered formalin for 2 min, air-dry, and store for staining. Leave cestodes, trematodes, and acanthocephalans in saline or tap water at 4°C (40°F) for several hours to allow them to relax and evacuate their eggs from the uterus; then remove them, and stretch them out in either a pan or petri dish depending upon their size. Open cestode cysts to evert the scolex. Flood them slowly with hot (60 to 70°C) (140 to 158°F) AFA (10 parts commercial formalin, 50 parts 95% ethanol, 2 parts glacial acetic acid, 40 parts distilled water), and let them sit for 1 to 2 hours. If the parasite is thick, fix it under slight pressure. If future work with any helminth specimen involves examination by electron microscopy (EM) or DNA or RNA work, prepare accordingly. For scanning electron microscopy (SEM), best results can be attained with 4% paraformaldehyde (phosphate buffered) and AFA solution. For transmission electron microscopy (TEM), fix specimens in gluteraldehyde (∼1 hour), rinse them in a phosphate buffer, and postfix in osmium tetroxide (∼1.5 hours). Rinse material for DNA or RNA work in saline, and freeze at –70°C, or preserve in 95% ethanol at 4°C. Drop living nematodes into steaming (not boiling) glycerin alcohol (1 part glycerin, 3 parts 95% ethanol), which will result in less shrinkage than alcohol alone. The heated solution causes the worms to straighten instantly and die in that position, thus avoiding the curled and distorted specimens obtained when using cold fixatives. Relax arthropods in Boardman’s solution (97 ml 20% ethanol, 3 ml ether) and transfer them to 70% ethanol for storage. Descriptions of clearing and mounting techniques are given in Dailey (1978). For further references on identification of marine mammal cestode, trematode, and nematode parasites, see Price (1932), Delyamure (1955), Schell (1985), Arvy (1982), Khalil et al. (1986), Margolis and Arai (1989), Margolis and Dailey (1972), Anderson (2000), and Commonwealth Institute of Helminthology Keys to the Nematode Parasites of Vertebrates (Anderson et al., 1974–1983). Duszynski et al. (1998) review coccidia of marine mammals, Heinrich (1999) the protozoa, and McDaniel (1979) mites and ticks. Advice may also be obtained from the following Web sites: http://www.museum.unl.edu/asp/ www.museum.unl.edu/asp/ascaris.med.tmd.ac.jp/ www.biosci.ohio-state.edu/~parasite/home.html www.oie.int www.pasteur.fr/helios.bto.ed.ac.uk/mbx/fgn/wow/parasites.html www.nhm.ac/zoology/hp-dat.htm dspace.dial.pipex.com/town/plaza/aan18/urls.htm
0839_frame_C18 Page 359 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
359
Treatment New drugs for treating parasites of domestic animals and humans are continually being developed because of the severe economic and health impacts that parasites can have, and because of the development of drug-resistance. A drug developed against a tapeworm of humans is likely to be effective in killing tapeworms of marine mammals, but the effects on the marine mammal may be very different from those on humans. Thus, caution is always warranted when utilizing a novel drug in a new host species. Many of the older and more widely used anthelmintics have now been used in marine mammals, although few have been tested rigorously to determine the extent of parasite removal. Some have severe side effects on the treated marine mammal, even causing death (see Chapter 31, Pharmaceuticals). Generally, praziquantel has been used against a variety of flatworms (cestodes and trematodes), fenbendazole and ivermectin against roundworms (nematodes), and ivermectin against ectoparasites. The protozoan infections of marine mammals are relatively novel, and treatments are more varied. As parasites of marine mammals are ubiquitous, and the effects on the host vary with the immune status of the host, supportive care is often more important than killing the parasites infecting the host (see Chapter 41, Seals and Sea Lions). In addition, the host’s inflammatory response to dead parasites may be more severe than that to live parasites, so treatment may exacerbate clinical signs. As so few successful treatments of marine mammal parasitic diseases have been published, the reader is advised to treat marine mammals cautiously, always to confer with other clinicians with previous experience, and to consult the literature. Dosages of anthelmintics that have been reported as used in marine mammals are given in Chapter 31, Pharmaceuticals.
Parasites of Cetacea Protozoa Ciliates
Haematophagus megapterae has been reported attached to the baleen plates of humpback (Megaptera novaeangliae), fin (Balaenoptera physalus), and blue (Balaenoptera musculus) whales (Woodcock and Lodge, 1921). It feeds on red blood cells, but is not considered pathogenic. Kyaroikeus cetarius was first described from the blowhole of a bottlenose dolphin (Tursiops truncatus), but has since been reported from blowholes of killer (Orcinus orca), false killer whales (Pseudorca crassidens), and belugas (Delphinapterus leucas) (Sneizek et al., 1995). It has also been found in skin lesions and lymph nodes. Chilodonella sp. (Figure 1) and another unclassified “helmet shaped” form have been routinely found in the blowhole mucus and skin scrapings of bottlenose dolphins from California (Arkush, pers. comm.). These ciliates are all considered opportunistic infections, and it is unknown at present whether they are pathogenic. Need for treatment is currently unclear. A number of unidentified ciliates have also been reported from the blowhole, lungs, lymph nodes, and skin of bottlenose dolphins (Howard et al., 1983; Dailey, 1985; Schulman and Lipscomb, 1997; respectively). Apicomplexans
Sarcocystis spp. and Toxoplasma gondii have been observed in a variety of marine mammal hosts, and within different tissues. In muscle, they appear to have little or no effect on the host, whereas in nervous tissue, they can lead to severe encephalitis (Dailey, 1985). Sarcocystis balaenopteralis was described from the skeletal muscle of a sei whale (Balaenoptera borealis) (Akao, 1970) and Sarcocystis sp. from a beluga, a northern right whale dolphin (Lissodelphis borealis), a pilot whale (Globicephala macrorhynchus), a striped dolphin (Stenella coeruleoalba), and a sperm whale
0839_frame_C18 Page 360 Tuesday, May 22, 2001 11:13 AM
360
CRC Handbook of Marine Mammal Medicine
FIGURE 1 Chilodonella sp. from skin scrapings of a bottlenose dolphin. (Photograph courtesy of S. Poynton.)
(Physeter macrocephalus) (De Guise et al., 1993; Cowan et al., 1986; Munday et al., 1978; Dailey and Stroud, 1978; Owens and Kakulas, 1968; respectively) (for figure see Dailey and Stroud, 1978). Toxoplasma gondii (Figure 2) has been reported from Atlantic bottlenose, Risso’s (Grampus griseus), striped and spinner (Stenella longirostris) dolphins (Cruickshank et al., 1990; Inskeep et al., 1990; Migaki et al., 1990; Di Guardo et al., 1995). Neither ante-mortem diagnosis nor treatment of protozoal encephalitis in cetaceans has been reported. Cystoisospora delphini, a coccidian, was described by Kuttin and Kaller (1996) as the cause of enteritis in bottlenose dolphins, although Duszynski et al. (1998) refer to it as an “unknown coccidia” of fish origin that was just passing through the dolphin. Flagellates
An unidentified species of flagellate (Chilomastix or Hexamita) was reported from the colon of a bowhead whale (Balaena mysticetus) by Heckmann et al. (1987). Kinetoplastid flagellates (family Bodonidae) have been reported from blowhole mucus of captive bottlenose dolphins and a stranded pygmy sperm whale (Kogia breviceps) (Figure 3) (Van Bonn, Sweeney, and Poynton, pers. comm.). Their clinical significance is unclear. Sarcodina
An Entamoeba sp. was described in the colon of a bowhead whale (Heckmann et al., 1987). The clinical significance of amoebae in cetaceans is yet to be determined.
0839_frame_C18 Page 361 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
361
FIGURE 2 Toxoplasma gondii bradyzoite (electron micrograph, original magnification ×5850). (University of Florida, College of Veterinary Medicine, courtesy of Claus Buergelt and Journal of the American Veterinary Medical Association.)
FIGURE 3 Kinetoplastid flagellate from Kogia breviceps blowhole mucus. (Photograph courtesy of S. Poynton.)
Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans) Gastrointestinal Tract
Nematodes of the family Anisakidae—Anisakis, Contracaecum, Pseudoterranova (= Phocanema, Terranova, Porrocaecum)—are composed of several complexes of sibling species (for comments on the systematics, see Paggi et al., 1991; Nascetti et al., 1993; Mattiucci et al., 1997). Members of these three genera are the most common nematodes of cetaceans, and are reported from most marine mammal species examined (although Contraceacum and Pseudoterranova are
0839_frame_C18 Page 362 Tuesday, May 22, 2001 11:13 AM
362
CRC Handbook of Marine Mammal Medicine
FIGURE 4 Esophageal types of anisakid worms (Anisakis, Phocanema/Pseudoterranova, Contracaecum).
usually only larval forms) (Dailey and Brownell, 1972). They can be distinguished from each other by esophageal morphology (Figure 4) and patent infections detected by characteristic ova or whole worms in feces (Figures 5 and 6). Mild infections rarely cause clinical signs, but heavy infections can result in gastritis and ulceration (Dailey, 1985; Smith, 1989). Prophylactic treatment with fenbendazole or ivermectin is performed routinely in captive dolphins to prevent accumulation of infection (Van Bonn, pers. comm.). Dailey (1985) lists ten genera of trematodes inhabiting the gastrointestinal system. Although these may cause irritation, they are not considered to cause health problems. Diagnosis of these worms is by detection of operculate eggs in the feces. Treatment has not been documented, although these parasites will probably be removed during treatment for cestode infections. Adult tapeworms in cetaceans are of two families (Tetrabothriidae and Diphyllobothriidae) and eight genera (Diplogonoporus, Diphyllobothrium, Hexagonoporus, Plicobothrium, Priapocephalus, Tetrabothrius, Trigonocotyle, and Strobilocephalus). Of these, only the last is considered pathogenic. Strobilocephalus triangularis penetrates the colon wall and forms necrotic ulcers (Dailey, 1985). Diagnosis of tapeworms is by detection of eggs or segments in the feces (Figure 7 is Diphyllobothrium). Most tapeworms are susceptible to praziquantel, which has been used in a variety of odontocetes (see Chapter 31, Pharmaceuticals). Two genera of acanthocephalans in the family Polymorphidae (Corynosoma and Bolbosoma) are found in marine mammals. Cetaceans are the primary host of Bolbosoma spp., whereas Corynosoma spp. may occur in cetaceans, but typically are found in pinnipeds (Dailey, 1985). Although acanthocephalans are not considered a serious disease problem, numerous abscesses caused by B. balanae were seen in a dead stranded gray whale (Eschrichtius robustus) (Dailey et al., 2000). Diagnosis is made by detection of spindle-shaped eggs in feces (Figure 8), but treatment has not been documented.
0839_frame_C18 Page 363 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
FIGURE 5 Anisakis sp. egg (original magnification × 400).
FIGURE 6 Contracaecum egg (original magnification ×400).
363
0839_frame_C18 Page 364 Tuesday, May 22, 2001 11:13 AM
364
CRC Handbook of Marine Mammal Medicine
FIGURE 7 Tapeworm egg (Diphyllobothrium sp.) (original magnification ×400).
FIGURE 8 Acanthocephalan (Bolbosoma sp.) egg (original magnification ×400).
0839_frame_C18 Page 365 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
365
FIGURE 9 Zalophotrema hepaticum egg (original magnification ×400).
Liver
Three genera of trematodes, Campula, Oschmarinella, and Zalophotrema, occupy the hepatic and pancreatic ducts of toothed whales, while Lecithodesmus infects baleen whales. Infection may lead to weight loss, decreased liver function, and predisposition to bacterial disease in chronic cases, or hepatic trauma due to migration of worms resulting in hepatitis, and death in acute cases (Zam et al., 1971). Diagnosis is dependent upon detection of eggs in feces (Figure 9). These parasites are probably sensitive to praziquantel, although treatment has not been documented. Respiratory System, Sinuses, and Brain
Trematodes of both the genera Hunterotrema and Nasitrema infect these sites in cetaceans, the latter genus (Nasitrema) being the most common in the brains/heads of small odontocetes. Nasitrema may be a significant cause of stranding in a variety of odontocetes. Nasitrema was the cause of parasitogenic eighth cranial neuropathy in a stranding of Risso’s dolphins (Morimitsu et al., 1992), encephalitis in a striped dolphin (O’Shea et al., 1991), and cerebral necrosis in stranded common dolphins (Delphinus delphis) (Dailey and Walker, 1978). Symptoms include loss of equilibrium, generalized central nervous signs, and head lashing. Diagnosis is by detection in mucus of eggs that are triangular in cross section (Figure 10). Treatment of bottlenose and common dolphins with praziquantel has been described (Gage, pers. comm.; see Chapter 31, Pharmaceuticals), although killing the parasites may exacerbate clinical signs. Nematodes infecting the lungs, pulmonary blood vessels, auditory spaces, and air sinuses are from two families (Pseudaliidae and Crassicaudidae), from which four genera (Halocercus, Pharurus, Pseudalius, and Stenurus) are the most common. Information on the lungworms of marine mammals has recently been comprehensively reviewed (Measures, 2001). Symptoms of lungworm infestation vary with host and parasite species, as well as with intensity
0839_frame_C18 Page 366 Tuesday, May 22, 2001 11:13 AM
366
CRC Handbook of Marine Mammal Medicine
FIGURE 10 Nasitrema sp. egg (original magnification ×400).
of infection. They include cough, dyspnea, lethargy, and potentially death (Moser and Rhinehart, 1993). Pseudaliids in the cranial sinus system of odontocetes have been suggested to cause stranding of the host, although negative effects on infected individuals have not been demonstrated (Geraci, 1979). Diagnosis is by detection of larvae from feces or blowhole mucus samples. Treatment of lungworms with ivermectin or fenbendazole has been successful (Chapter 31, Pharmaceuticals), although supportive care with antibiotics and mucolytics is also important. Crassicaudids have been reported from the air sinuses of nine genera of small odontocetes, where they have been implicated as the cause of bone erosion (Pascual et al., 2000). This erosion results from inflammation of the mucosa, purulent sinusitis, and osteitis (Raga et al., 1982). Their role in stranding is not clear (Delyamure, 1955; Geraci, 1979). Urogenital System
The family Crassicaudidae consists of two genera of large nematodes, Crassicauda and Placentonema. Members of the genus Crassicauda are commonly observed in mammary tissue, kidneys, and around the genitalia. The tails of adult worms extend into the renal calyxes or other openings, and eggs are released into the urine. The effect on individual animal health is unclear, but it has been suggested that these parasites may reduce populations of certain baleen whales (Lambertsen, 1986; 1992), and reduce reproductive success of Atlantic white-sided dolphins (Lagenorhynchus acutus) by decreasing milk production (Geraci et al., 1978). Placentonema gigantisma is a parasite of the sperm whale, and when present is thought to cause fetal death (Dailey, 1985). Diagnosis of crassicaudid infections is by detection of eggs or developing larvae in milk, urine, or tissue (Figure 11). Treatment has not been documented, although Lambertsen (1992) suggested using ivermectin by remote darting to treat fin whales with C. boopis. Connective Tissue
Two common larval cestodes (Phyllobothrium delphini and Monorygma grimaldii) are found free in the peritoneum, or in blubber and connective tissue (Dailey and Brownell, 1972; Norman, 1997). Agusti et al. (2000) speculate that two morphotypes, large and small, of a third
0839_frame_C18 Page 367 Tuesday, May 22, 2001 11:13 AM
367
CRC Handbook of Marine Mammal Medicine
FIGURE 11 Crassicauda sp. eggs (original magnification ×400).
larval type (Scolex pleuronectis), found in numerous marine organisms including cetaceans, are the precursors of these two parasites. Since these worms infect elasmobranch fishes as adults, infection of cetaceans may represent a “dead-end host” for these parasites, although the cetacean may be the natural intermediate host. No clinical signs or pathology are reported in association with infection, although intensity of infection may be heavy.
Ectoparasites Cetaceans are the host of several groups of external parasites (barnacles, copepods, and whale lice) that are primarily found on baleen whales. Whale lice in the family Cyamidae are the most numerous (26 species) (Dailey and Brownell, 1972). They feed on epidermal tissue, body fluids, and algal filaments. They require no intermediate host and are transmitted from whale to whale by contact. Clinical signs have not been associated with infection, and treatment is probably unnecessary.
Parasites of Pinnipeds Protozoa Apicomplexans
Although the coccidian Eimeria phocae (Figure 12) caused the deaths of captive harbor seals (Phoca vitulina) in Maine (Hsu et al., 1974) and Scotland (Munro and Synge, 1991), six other species of apicomplexans have been reported from Antarctic pinnipeds exhibiting no clinical signs (Drozda, 1987). Duszynski et al. (1998) suggest that the latter are incidental
0839_frame_C18 Page 368 Tuesday, May 22, 2001 11:13 AM
368
CRC Handbook of Marine Mammal Medicine
FIGURE 12 Eimeria phocae (original magnification ×400).
parasites of fish, and dispute their validity as marine mammal parasites. Munro and Synge (1991) suggest E. phocae may not affect seal health, unless the host is stressed through capture, handling, or changes in diet. They recommend a three-pronged approach to treatment. These are sulfonamides to interfere with coccidial metabolism, broad-spectrum antibiotics to contain secondary infection of the intestine, and rehydration via stomach tube to aid renal excretion of sulfonamides. Sarcocystis richardi has been reported from the harbor seal diaphragm (Hadwen, 1922). Sarcocystis spp. have been reported from California sea lions (Zalophus californianus), bearded seals (Erignathus barbatus), leopard seals (Hydrurga leptonyx), and ringed seals (P. hispida), as well as the Antarctic (Arctocephalus gazella) and northern (Callorhinus ursinus) fur seals (Mense et al., 1992; Bishop, 1979; Odening, 1983; Migaki and Albert, 1980; Baker and Doidge, 1984; Brown et al., 1974; respectively). As in cetaceans, Sarcocystis spp. in muscle do not appear to cause clinical signs. However, S. neurona was recently found in brain tissue associated with encephalitis in stranded harbor seals (LaPointe et al., 1998). Toxoplasma spp. have also been reported from cases of encephalitis in harbor seals and a northern fur seal (Van Pelt and Dieterich, 1973; Holshuh et al., 1985), as well as disseminated organ infection in two California sea lions (Lauckner, 1985). Serological methods of ante-mortem diagnosis are currently under evaluation, so that effective treatment can be determined (Chechowitz et al., 2000). Flagellates
Giardia sp. has been reported from adult harp (Pagophilus groenlandicus) and gray (Halichoerus grypus) seals, and one juvenile harbor seal in eastern Canada, ringed seals in the western Arctic region of Canada (Olson et al., 1997; Measures and Olson, 1999), and California sea lions from northern California (Deng et al., 2000). These pinnipeds may be potential reservoir hosts of giardiasis in these regions, but their effects on pinniped health are unknown.
0839_frame_C18 Page 369 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
369
Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans) Gastrointestinal Tract
Two families of nematodes infect the pinniped gastrointestinal tract, Ancylostomatidae (hookworms) and Anisakidae (large roundworms). Two hookworms of the genus Uncinaria (U. lucasi and U. hamiltoni) are thought to cause high mortality in young pinnipeds in which the adult worms cause hemorrhagic enteritis and anemia (Olsen, 1958; Keyes, 1965). Vertical transmission of U. lucasi occurs in northern fur seals (Lyons, 1994). Diagnosis is by detection of eggs in the feces (Figure 13). Once the pinniped host is over a certain age, hookworms fail to mature to produce ova; thus, fur seal pups may be anemic due to hookworm infection, but fail to show eggs in the feces, since the adult hookworms have been shed. Anisakids in pinnipeds are represented by the same genera that are found in cetaceans (Anisakis, Contracaecum, Pseudoterranova, and Phocascaris), with the latter three being more prevalent as adults, whereas Anisakis is most commonly found in its larval form. Intensity of infection within the stomachs of pinnipeds may be high with no apparent ill effects, although both larval and adult anisakids have been associated with clinical signs, including gastritis, gastric ulceration, enteritis, diarrhea, dehydration, anemia (Young and Lowe, 1969; Lauckner, 1985), and, rarely, gastric perforation (Ridgway et al., 1975). It is likely that both mechanical action and secretion of allergens play roles in nematode-associated gastric lesions (Lauckner, 1985). Contracaecum corderoi (previously known as (=) C. ogmorhini) has caused severe peritonitis and death in California sea lions resulting from perforated ulcers in the proximal duodenum (Fletcher et al., 1998). Diagnosis of anisakid infections is by detection of eggs in feces (see Figures 5 and 6). Hookworm infections of Steller sea lion (Eumatopias jubatus) pups have been treated successfully with fenbendazole (Panacur®, Hoechst, Somerville, NJ) at 50 mg/kg for
FIGURE 13 Hookworm egg (Uncinaria sp.) from Miroungsa angustirostris (original magnification ×400).
0839_frame_C18 Page 370 Tuesday, May 22, 2001 11:13 AM
370
CRC Handbook of Marine Mammal Medicine
4 days (D. Huff, pers. comm.). California sea lions, and harbor and northern elephant seals (Mirounga angustirostris) have been treated successfully with either fenbendaole at 10 mg/kg for 3 successive days, or ivermectin at 200 µg/kg twice, 10 days apart (Gage, Gulland, and Haulena, pers. comm.). Trematodes in the gastrointestinal system of pinnipeds belong primarily to ten genera, (Cryptocotyle, Galactosomum, Rossicotrema, Phagicola, Stictodera, Phocitrema, Pricetrema, Microphallus, Maritrema, and Ogmogaster), of which Pricetrema is reported most commonly. Intestinal trematodes may occur in massive numbers but are not considered pathogenic, although colitis is occasionally observed histologically in infected elephant seals (Lowenstine, pers. comm.). Treatment of California sea lions and elephant seals with a single dose of praziquantel at 10 mg/kg has been effective in removing eggs from feces (Gage, Gulland, and Haulena, pers. comm.). Numerous species of cestodes are found in pinnipeds, representing the same two families as in cetaceans (Diphyllobothriidae and Tetrabothriidae). Lauckner (1985) lists 19 and 15 species of the genus Diphyllibothrium from the Northern and Southern Hemispheres, respectively. Diphyllobothrium lanceolatum, D. pacificum, and D. tetrapterus are the most frequently reported species from captive pinnipeds. Cestode infections are seasonal, and can reach enormous intensities. Pathogenic effects of tapeworms in captive pinnipeds have been reported, although reports vary and the effects may occur only when large numbers obstruct the intestinal lumen (Lauckner, 1985). Diagnosis is by detection of eggs or segments in feces (see Figure 7). Rehabilitated pinnipeds (California sea lions and harbor and northern elephant seals) are commonly treated prophylactically with praziquantel at 10 mg/kg on 2 consecutive days (Gage, Gulland, and Haulena, pers. comm.) Acanthocephalans of the genus Corynosoma are cosmopolitan in pinnipeds, with C. obtuscens, C. strumosum (= C. semerme), C. villosum, and C. wegeneri (= C. hadweni) the most commonly encountered in captive animals. Bolbosoma spp. have been reported incidentally from pinnipeds. Diagnosis is by detection of eggs in feces (see Figure 8), but effects and treatment are poorly documented. Respiratory and Circulatory Systems
Species from three families of nematodes—the Filaroididae: Filaroides (Parafilaroides) spp.; Crenosomatidae: Otostrongylus circumlitus; and Filariidae: Acanthocheilonema (= Dipetalonema, Skrjabinaria) odendhali, A. spirocauda, and Dirofilaria immitis—infect these organs. The classification, ecology, and effects of these lungworms have recently been reviewed by Measures (2001). The life-cycles of both F. (P.) decorus and O. circumlitus involve fish as intermediate hosts, which are then eaten by weanling seals and sea lions (Dailey, 1970; Bergeron et al., 1997, Anderson, 2000). Filaroides (P.) spp. are cosmopolitan in both phocids and otariids, and frequently contribute to pneumonia in stranded animals (Lauckner, 1985). Their clinical significance, compared with that of concurrent bacterial infection, is often unclear, although F. (P.) decorus may be pathogenic alone in malnourished California sea lions (Gerber et al., 1993). Diagnosis of Filaroides (P.) spp. is by detection of first-stage larvae in feces or sputum. Treatment of California sea lions with ivermectin and concurrent prednisone or dexamethasone, in addition to antibiotic treatment, has been effective (Gage et al., 1993). Otostrongylus circumlitus is a parasite of the heart and lungs of seals (Measures, 2001). Host response to infection varies between species. Gulland et al. (1997) report this worm as the cause of death in 73 stranded, juvenile northern elephant seals, while Munro et al. (1992) found that it did not cause any inflammatory reaction in young harbor seals. Clinical signs in stranded elephant seals include anorexia, depression, dehydration, neutrophilia, disseminated intravascular
0839_frame_C18 Page 371 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
371
FIGURE 14 Otostrongylus circumlitis (large), Filaroides (P.) sp. (small), from Mirounga angustirostris (original magnification ×400).
coagulation, and death. Diagnosis of patent infections is by detection of first-stage larvae in feces or sputum (Figure 14). However, stranded northern elephant seals usually die during the prepatent period, so there is no method of diagnosis currently available (Gulland et al., 1997). Efficacy of treatment cannot be determined as no ante-mortem method of diagnosis exists. However, in harbor seals, treatment with both fenbendazole and ivermectin, accompanied by dexamethasone, antibiotics, and mucolytic agent treatment, has been effective (Haulena and Gulland, pers. comm.). When treating lungworm infections of pinnipeds, care must be taken to prevent sudden killing of parasites, as this can exacerbate the clinical signs. Thus, dexamethasone and antibiotic treatment is often initiated 3 days before giving the anthelmintic to debilitated pinnipeds (Gage, Gulland, and Haulena, pers. comm.). Acanthocheilonema spirocauda and A. odendhali are cosmopolitan parasites of both phocids and otariids found in the right ventricle of the heart and intermuscular fascia, respectively. Pathological changes associated with A. spirocauda infection can be considerable, whereas A. odendhali has no detectable effect on the host. Dirofilaria immitis (canine heartworm) has also been reported from captive pinnipeds maintained in endemic areas (Lauckner, 1985), with symptoms and pathology similar to those caused by A. spirocauda. Clinical signs associated with heartworm are anorexia, dyspnea, coughing, and erratic breathing. Diagnosis is by detection of microfilariae in blood smears. Although they are much alike, the microfilariae of these three species can be differentiated by their size, with the smallest A. odendhali (231 to 249 × 3.5 µm), followed by A. spirocauda (225 to 250 × 4.4 µm) and D. immitis (285 to 290 × 5.5 µm). Preventive treatment of captive California sea lions in D. immitis-endemic areas with ivermectin during the mosquito season is recommended (see Chapter 41, Seals and Sea Lions). As
0839_frame_C18 Page 372 Tuesday, May 22, 2001 11:13 AM
372
CRC Handbook of Marine Mammal Medicine
transmission of A. spirocauda is suspected to be via the seal louse (Echinophthirius horridus), but has not been demonstrated experimentally, control of ectoparasite infections may reduce the likelihood of heartworm transmission (Geraci et al., 1981). Liver, Biliary System, and Pancreas
Only trematodes infect these organs in pinnipeds. They consist of two families (Campulidae, Opistorchiidae) and five genera (Orthosplanchnus, Zalophotrema, Opistorchis, Metorchis, Pseudamphistomum). Of these, the most commonly reported is Z. hepaticum from the California sea lion, harbor seal, and northern elephant seal. Prevalence increases with age in California sea lions, and many older animals have thickened biliary ducts, probably as a consequence of infection (Lowenstine, pers. comm.). Diagnosis is by detection of eggs in feces (see Figure 9), and treatment of California sea lions with praziquantel at 10 mg/kg has been successful (Gage, Gulland, and Haulena, pers. comm.). Connective Tissue
Worms found in connective tissue other than A. odenhali (see above) are the larval cestode, Phyllobothrium delphini, and the nematode, Trichinella nativa. These are incidental findings, with the former reported from the southern hemisphere (sea lion, fur seal, and leopard seal) and the latter from walrus (Odobenus rosmarus), bearded, and ringed seals in the Arctic.
Ectoparasites Mites (Demodex and Sarcoptes spp.) and lice (Anoplura spp.) are ectoparasites of pinnipeds, and most infections are species specific (Lauckner, 1985). The significance of these parasites depends upon the numbers infesting the host. Heavy burdens of lice can cause irritation, alopecia, and anemia, although because they are often secondary to debilitation, it may be difficult to determine the exact cause of the anemia (i.e., other parasites and/or malnutrition may also cause anemia). The lice can be observed with the naked eye, and are readily treated with ivermectin, dichlorvos, or disophenol systemically, or topical rotenone louse powder. Demodicosis, also characterized by hyperkeratosis, alopecia, and pruritus, has been observed in California sea lions and northern fur seals (Gulland and Spraker, pers. comm.). Diagnosis is based on histological detection in biopsies, or deep skin scrapings; treatment with amitraz has been effective (Sweeney, pers. comm.). Mites in the nares, nasopharynx, airways, and lungs of phocids belong to the genus Halarachne, whereas otariids and walrus (Odobenus spp.) are infested with members of the genus Orthohalarachne. Clinical signs range from sneezing and mucus discharge with nasal infections, to emphysema and lung infections. Transmission is by direct spread of larvae from animal to animal. Diagnosis is by detection of larval mites in sputum and nasal exudate. Treatment with ivermectin at 200 µg/kg twice, 2 weeks apart is usually effective.
Parasites of Sirenia Protozoa—Apicomplexans Toxoplasma gondii has caused death due to meningoencephalitis in the Florida manatee (Trichechus manatus) (Buergelt and Bonde, 1983). Coccidians, including Eimeria trichechi are reported from the Amazonian manatee (T. inunguis) (Lainson et al., 1983), and E. manatus and E. nodulosa from the Florida manatee (Upton et al., 1989). Their significance to sirenian health is unknown. Treatment has not been documented.
0839_frame_C18 Page 373 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
373
Helminths (Nematodes, Trematodes) The predominant parasites of sirenians are monostome trematodes (except for one species, Nudacotyle undicola), and they are exclusive to sirenians (Dailey et al., 1988). Nine genera (Chiorchis, Indosolenorchis, Lankatrema, Lankatremoides, Rhabdiopoeus, Schizamphistoma, Solenorchis, Taprobonella, and Zygocotyle) may inhabit the stomach, pyloric cecum, and intestine in massive numbers. Some (Lankatrema) produce lesions in the stomach and form cystic cavities in the mucosa. Three additional genera of trematodes (Opisthotrema, Chocleotrema, Pulmonicola) inhabit the nasal passages, Eustachian tubes, airways, and lungs, and a fourth genus (Labicola) occurs in the upper lip. Diagnosis is by detection of eggs with polar filaments in feces or sputum. Two genera of nematodes reported from sirenians are Heterocheilus and Paradujardina. Prophylactic anthelmintic treatment is routinely performed on captive manatees, but not on those released back into the natural environment (see Chapter 43, Manatees). Both copepods (Harpacticus) and barnacles (Chelonibia, Platylepas, and Balanus) have been reported from sirenia, without any associated disease (Lauckner, 1985).
Parasites of Sea Otters Protozoa—Apicomplexans Toxoplasma gondii (see Figure 2) and Sarcocystis neurona have been reported from only the California population of sea otters (Enhydra lutris nereis), in which prevalence is relatively high, and infection may be asymptomatic or severe, causing encephalitis (Thomas and Cole, 1996; Cole et al., 2000; Lindsay et al., 2000). Clinical signs vary from none to severe, generalized neurological signs. Methods for serological diagnosis and treatment are currently under investigation (Chechowitz et al., 2000; Murray, pers. comm.).
Helminths (Nematodes, Trematodes, Cestodes, Acanthocephalans) The helminth fauna of sea otters consists primarily of species acquired indirectly from pinnipeds or birds through ingestion of shared intermediate hosts. There are differences between the parasites of the Californian and Alaskan (Enhydra lutris kenyoni) populations of sea otters. Larval Anisakis spp. and adult Pseudoterranova azaras and P. decipiens are reported from only the Alaskan population (Margolis et al., 1997), as are intestinal perforations by larval nematodes (P. decipiens) resulting in death (Rausch, 1953). Six species of trematodes (Microphallus pirum, M. nicolli, Nanophyetes sp., Phocitrema fusiforme, Plenosoma minimum, and Pricetrema zalophi) infect the intestine, with one species (Orthosplanchnus fraterculus), found in the gall bladder. Only the two species of Microphallus and P. minimum are observed in the California population (Dailey, pers. obs.). All three species of cestodes (Diplogonoporus tetrapterus, Diplogonoporus sp., and Pyramicocephalus phocarum) are reported but only from Alaskan otters. The acanthocephalan Corynosoma enhydra is the only helminth of otters that is exclusive to this host species. Corynosoma stromosum, C. villosum, and three species of Profilicollis (= Polymorphus) are shared with pinnipeds and birds, respectively. Prevalence of mortality due to intestinal perforations by Profilicollis spp. is relatively high in California sea otters (Thomas and Cole, 1996). Diagnosis of acanthocephalan infection and the peritonitis associated with perforation of the intestinal tract is difficult, because death of the otter usually occurs before the acanthocephalans become patent. Success of treatment is thus difficult to determine, so effective treatment is still being assessed (Haulena and Murray, pers. comm.).
0839_frame_C18 Page 374 Tuesday, May 22, 2001 11:13 AM
374
CRC Handbook of Marine Mammal Medicine
Nasal mites of the genus Halarachne are common from otters in Alaska and California, but do not cause obvious clinical disease.
Parasites of Polar Bears Remarkably few parasites have been reported in polar bears (Ursus maritimus). Garner et al. (1997) report a fatal case of hepatic sarcocystosis in a captive polar bear. Two species of nematodes, Baylisascaris transfug and B. multipapillata, are reported in polar bears. These ascaridoid worms use small mammals as intermediate hosts, in which they are reported to cause neurological disease (Wallach and Boever, 1983; Fowler, 1986). Once these nematodes are established in enclosures of captive polar bears, it is very difficult to rid the enclosure completely of these parasites, since the ova can remain viable in the environment for as long as years. The clinical signs are variable, depending upon intensity of infection, and range from loose stools to diarrhea, rough hair coat, and, in extreme cases, severe weight loss and intestinal obstruction, leading to death. Trichinella nativa appears to have a sylvatic cycle in the Arctic. The larvae have been found in the muscle and fascia of polar bears, where they are generally considered incidental findings, and do not cause overt disease (Pozio et al., 1990; Nozais et al., 1996). When signs do occur, they generally are muscular pain and eosinophilia. At times, there can be central nervous system involvement. There are three species of cestodes reported in polar bears, Diphyllobothrium latum, Bothriocephalus spp., and Taenia ursi-maritimus (Leiby and Dyer, 1970). Diagnosis is by detection of proglottids or eggs in the feces.
Acknowledgments The author thanks all those who contributed to this chapter, specifically, Kristen Arkush, Bob Bonde, Cathy Bonde, Claus Buergelt, Malia Dailey, Don Duszynski, Laurie Gage, Frances Gulland, Marty Haulena, Antje Heinrich, David Lindsey, Jim McBain, Sarah Poynton, Toni Raga, Carol Stack, Jay Sweeney, Pam Tuomi, and Steve Upton. Also thanked are Dave Huff, Jay Sweeney, Linda Lowenstine, Mike Murray, and Bill Van Bonn for personal communications, and Lena Measures, Gene Lyons, and Sharon Tolliver for their reviews of this chapter.
References Agusti, C., Aznar, F.J., Montero, F.E., and Raga, J.A., 2000, The ontogeny of Scolex pleuronectis (Tetraphyllidea) in the striped dolphin, Stenella coerulealba, Abstr., 28th Meeting of the European Association of Aquatic Medicine, Benidorm, Spain, 11–15 March. Akao, S., 1970, A new species of Sarcocystis in the whale Balaenoptera borealis, J. Protozool., 17: 290–294. Anderson, R.C., 2000, Nematode Parasites of Vertebrates, Their Development and Transmission, C.A.B. International, New York, 592 pp. Anderson, R.C., Chabaud, A.G., and Wilmott, S., 1974–1983, Commonwealth Institute of Helminthology Keys to the Nematode Parasites of Vertebrates, Commonwealth Agricultural Bureaux, Farnham Royal, Bucks, U.K. Arvy, L., 1982, Phoresies and parasitism in cetaceans: A review, Invest. Cetacea, 14: 233–335. Baker, J.R., and Doidge, D.W., 1984, Pathology of the Antarctic fur seal (Arctocephalus gazella) in South Georgia, Br. Vet. J., 140: 210–219.
0839_frame_C18 Page 375 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
375
Bergeron, E., Measures, L.M., and Haut, J., 1997, Experimental transmission of Otostrongylus circumlitus (Raillet, 1899) (Metastrongyloidea: Crenosomatidae), a lungworm of seals in eastern arctic Canada, Can. J. Zoo., 75: 1364–1371. Bishop, L., 1979, Parasite related lesions in a bearded seal, Erignathus barbatus, J. Wildl. Dis., 15: 285– 293. Brown, R.J., Smith, A.W., and Keyes, M.C., 1974, Sarcocystis in the northern fur seal, J. Wildl. Dis., 10: 53. Buergelt, C.O., and Bonde, R.K., 1983, Toxoplasmic meningoencephalitis in a West Indian manatee, J. Am. Vet. Med. Assoc., 183: 1294–1296. Chechowitz, M.A., Sverlow, K., Crosbie, P.R., Packham, A., Gardner, I., Barr, B.C., Lowenstine, L.J., Gulland, F., Jessup, D., and Conrad, P., 2000, Protozoal brain infections in harbor seals (Phoca vitulina) and sea otters (Enhydra lutris nereis) in California, an update, in Proceedings of the American Association of Zoo Veterinarians and International Association for Aquatic Animal Health, New Orleans, Sept. 17–21, 345. Cole, R.A., Lindsay, D.S., Howe, D.K., Roderick, C.L., Dubey, J.P., Thomas, N.J., and Baeten, L.A., 2000, Biological and molecular characterizations of Toxoplasm gondii strains obtained from southern sea otters (Enhydra lutris nereis), J. Parasitol., 86: 526–530. Cowan, D.F., Walker, W.A., and Brownell, R.L. Jr., 1986, Pathobiology of small cetaceans stranded along southern California beaches, in Research on Dolphins, Bryden, M.M., and Harrison, R. (Eds.), Clarendon Press, Oxford, 323–367. Cruickshank, J.J., Hainea, D.M., Palmer, N.C., and St. Aubin, D.J., 1990, Cysts of a toxoplasma-like organism in an Atlantic bottlenose dolphin, Can. Vet. J., 31: 213–215. Dailey, M.D., 1970, The transmission of Parafilaroides decorus (Nematoda: Metastrongylidae) in the California sea lion (Zalophus californianus), in Proceedings of the Helminthologic Society of Washington, 37: 215–222. Dailey, M.D., 1978, Preparation of parasites for identification and cataloging, J. Zoo Anim. Med., 9: 13–15. Dailey, M.D., 1985, Diseases of mammals: Cetacea, in Diseases of Marine Animals, Vol. 4, Pt. 2, Kinne, O. (Ed.), Biologische Anstalt Helgoland, Hamburg, chap. 7, 805–847. Dailey, M.D., and Brownell, R.L., Jr., 1972, A checklist of marine mammal parasites, in Mammals of the Sea, Biology and Medicine, Ridgway, S. (Ed.), Charles C Thomas, Springfield, IL, 528–589. 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. Dailey, M.D., and Walker, W.A., 1978, Parasitism as a factor in single strandings of southern California cetaceans, J. Parasitol., 64: 593–596. Dailey, M.D., Vogelbein, W., and Forrester, D.J., 1988, Moniligerum blairi n. g., n. sp. and Nudacatyle undicola n. sp. (Trematoda: Digenea) from the West Indian manatee, Trichechus manatus L., Syst. Parasitol., 11: 159–163. Dailey, M.D., Gulland, F.M., Lowenstine, L., Silvagni, P., and Howard, D., 2000, Prey, parasites and pathology associated with the mortality of a juvenile gray whale (Eschrichitus robustus) stranded along the northern California coast, Dis. Aquat. Organisms, 42: 111–117. De Guise, S., Lagace, A., Girard, C., and Beland, P., 1993, Intramuscular Sarcocystis in two beluga whales and an Atlantic white-sided dolphin from the St. Lawrence estuary, Quebec, Canada, J. Vet. Diagn. Invest., 5: 296–300. Delyamure, S.L., 1955, Helminthofauna of Marine Mammals (Ecology and Phylogeny) [in Russian], Skrjabin, K.I. (Ed.), Izdatel’stvo Akademii Nauk SSR, Moscow, Translated by Israel Program for Scientific Translations, Jerusalem, 1968, 522 pp. Deng, M.Q., Peterson R.P., and Cliver, D.O., 2000, First findings of Cryptosporidium and Giardia in California sea lions (Zalophus californianus), J. Parasitol., 86: 490–494. DiGuardo, G., Agrimi, U., Morelli, L., Cardeti, G., Terracciano, G., and Kennedy, S., 1995, Post-mortem investigations on cetaceans found stranded on the coasts of Italy between 1990 and 1993, Vet. Res., 136: 439–442.
0839_frame_C18 Page 376 Tuesday, May 22, 2001 11:13 AM
376
CRC Handbook of Marine Mammal Medicine
Drozda, J., 1987, Oocysts of six new Coccidomorpha species from pinnipeds of King George Island (South Shetlands, Antarctic), Acta Protozool., 26: 263–266. Duszynski, D.W., Upton, S.J., and Couch, L., 1998, Coccidia (Eimeriidae) of marine mammals (cetacea, pinnipeds, sirenia), in Coccidia of the World, NSF grant PEET DEB 9521687. Fletcher, D., Gulland, F.M.D., Haulena, M., Lowenstine, L.J., and Dailey, M., 1998, Nematode-associated gastrointestinal perforations in stranded California sea lions (Zalophus californianus), in International Association for Aquatic Animal Medicine 29th Annual Conference Proceedings, San Diego, CA, 59. Fowler, M.E., 1986, Ursidae, in Zoo and Wild Animal Medicine, W.B. Saunders, Philadelphia, 811 pp. Gage, L.J., Gerber, J.A., Smith, D.M., and Morgan, L.E., 1993, Rehabilitation and treatment success rate of California sea lions (Zalophus californianus) and northern fur seals (Callorhinus ursinus) stranded along the central and northern California coast, 1984–1990, J. Zoo Wildl. Med., 24: 41–47. Garner, M.M., Barr, B.C., Peckham, A.E., Marsh, A.E., Burek-Huntington, K.A., Wilson, R.K., and Dubey, J.P., 1997, Fatal hepatic sarcocystosis in two polar bears (Ursus maritimus), J. Parasitol., 83: 523–526. Geraci, J.R., 1979, The role of parasites in marine mammal strandings along the New England Coast, in Biology of Marine Mammals: Insights through Strandings, Geraci, J.R., and St. Aubin, D.J. (Eds.), U.S. Department of Commerce, NTIS Report PB-293-890, pp. 85–91. Geraci, J.R., Dailey, M.D., and St. Aubin, D.J., 1978, Parasitic mastitis in the Atlantic white-sided dolphin, Lagenorhynchus acutus, as a probable factor in herd productivity, J. Fish. Res. Board Can., 35: 1350– 1355. Geraci, J.R., Fortin, J.F., St. Aubin, D.J., and Hicks, B.D., 1981, The seal louse, Echinophthirius horridus, an intermediate host of the seal heartworm, Dipetalonema spirocauda (Nematoda), Can. J. Zool., 59: 1457–1459. Gerber, J.A., Roletto, J., Morgan, L.E., Smith, D.M., and Gage, L., 1993, Findings in pinnipeds stranded along the central and northern California coast, 1984–1990, J. Wildl. Dis., 29: 423–433. Gulland, F.M.D., Beckman, K., Burek, K., Lowenstine, L., Werner, L., Spraker, T., Dailey, M., and Harris, E., 1997, Nematode (Otostrongylus circumlitus) infestation of northern elephant seals (Mirounga angustirostris) stranded along the central California coast, Mar. Mammal Sci., 13: 446–459. Hadwen, S., 1922, Cyst-forming protozoa in reindeer and caribou, and a sarcosporidian parasite of the seal (Phoca richardsi), J. Am. Vet. Med. Assoc., 61: 374–382. Heckman, R.A., Jensen, L.A., Warnack, R.G., and Coleman, B., 1987, Parasites of the bowhead whale, Balaena mysticetus, Great Basin Nat., 47: 355–372. Heinrich, A.B., 1999, Protozoans from Cetaceans: Morphology, Taxonomy, and Clinical Significance, Thesis für Diplomarbeit im Fach Zoologie, Christian-Albrechts-Universität zu Kiel. Holshuh, H.J., Sherrod, A.E., Taylor, C.R., Andrews, B.F., and Howard, E.B., 1985, Toxoplasmosis in a feral northern seal, J. Am. Vet. Med. Assoc., 187: 1229–1230. Howard, E.B., Britt, J.O., Jr., and Matsumoto, G.K., 1983, Parasitic diseases, in Pathobiology of Marine Mammal Diseases, Vol. 1, Howard, B. (Ed.), CRC Press, Boca Raton, FL, 119–232. Hsu, C.K., Melby, E.C., and Altman, N.H., 1974, Eimeria phocae sp. n. from the harbor seal (Phoca vitulina concolor), J. Parasitol., 60: 399–402. Inskeep, W.H., Gardiner, C.H., Harris, R.K., Dubey, J.P., and Goldston, R.T., 1990, Taxoplasmosis in Atlantic bottle-nosed dolphins (Tursiops truncatus), J. Wildl. Dis., 26: 377–382. Keyes, M.C., 1965, Pathology of the northern fur seal, J. Am. Vet. Med. Assoc., 147: 1090–1095. Khalil, L.F., Jones, A., and Bray, R.A. (Eds.), 1986, Keys to the Cestode Parasites of Vertebrates, 1986, International Institute of Parasitology, CAB International, Hertfordshire, U.K. Klumov, S.K., 1963, Food and helminth fauna of whalebone whales (Mystacoceti) in the main whaling regions of the world oceans, Trudy Inst. Okeanol., 71: 94–194 [Fisheries and Research Board of Canada Translation Series 589]. Kuttin, E.S., and Kaller, A., 1996, Cystoisospora delphini n. sp. enteritis in a bottlenosed dolphin (Tursiops truncatus), Aquat. Mammals, 22: 57–60.
0839_frame_C18 Page 377 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
377
Lainson, R., Naiff, R.D., Best, R.C., and Shaw, J.J., 1983, Eimeria trichechi n. sp. from the Amazonian manatee, Trichechus inunguis (Mammalia: Sirenia), Syst. Parasitol., 5: 287–289. Lambertsen, R., 1986, Disease of the common fin whale: Crassicaudosis of the urinary system, J. Mammal., 67: 353–366. Lambertsen, R., 1992, Crassicaudosis: A parasitic disease threatening the health and population recovery of large baleen whales, Rev. Sci. Tech. Off. Int. Epiz., 11: 1131–1141. LaPointe, J.M., Duignan, P.J., March, A.E., Gulland, F.M., Barr, B.C., Naydan, D.K., Kang, D.P., Farman, C.A., Burek, K.A., and Lowenstine, L.J., 1998, Meningoencephalitis due to a Sarcocystis neuronalike protozoan in Pacific harbor seals (Phoca vitulina richardsi), J. Parasitol., 84: 1184–1189. Lauckner, G., 1985, Diseases of Mammalia: Pinnipeda, in Diseases of Marine Animals, Vol. IV, Pt. 2, Kinne, O. (Ed.), Biologische Anstalt Helgoland, Hamburg, chap. 5, 683–793. Leiby, P.D., and Dyer, W.G., 1970, Cyclophyllidean tapeworms of wild carnivora, in Parasitic Diseases of Wild Mammals, Davis, J.W., and Anderson, R.C. (Eds.), Iowa State Press, Ames, 332–343. Lindsay, D.S., Thomas, N.J., and Dubey, J.P., 2000, Biological characterisation of Sarcocystis neurona isolated from a southern sea otter (Enhydra lutris nereis), Int. J. Parasitol., 30: 617–624. Lyons, E.T., 1994, Vertical transmission of nematodes: Emphasis on Uncinaria lucasi in northern fur seals and Strongyloides westeri in equids, J. Helminthol. Soc. Wash., 61: 169–178. Margolis, L., and Arai, H.P., 1989, Parasites of marine mammals, in Synopsis of the Parasites of Vertebrates of Canada, Kennedy, M.J. (Ed.), Alberta Agriculture, Edmonton, Alberta, Canada, 26 pp. Margolis, L., and Dailey, M.D., 1972, Revised annotated list of parasites from sea mammals caught off the West Coast of North America, U.S. Department of Commerce, National Oceanic and Atmospheric Administration Technical Report, NMFS SSRF-647, 23 pp. Margolis, L., Groff, J.M., Johnson, S.C., McDonald, T.E., Kent, M.L., and Blaylock, R.B., 1997, Helminth parasites of sea otters (Enhydra lutris) from Prince William Sound, Alaska: Comparison with other populations of sea otters and comments on the origin of their parasites, J. Helminthol. Soc. Wash., 64: 161–168. Mattiucci, S., Nascetti, G., Cianchi, R., Paggi, L., Andoino, P., Margolis, L., Bratley, J., Webb, S., D’Amelio, S., Orecchia, P., and Bullini, L., 1997, Genetic and ecological data on the anisakis simplex complex, with evidence for a new species (nematoda, ascaridoidea, anisakidae), J. Parasitol., 83: 401–416. McDaniel, B., 1979, How to Know the Mites and Ticks, W.C. Brown, Dubuque, IA, 335 pp. Measures, L.N., 2001, Lungworms of marine mammals, in Parasitic Diseases of Wild Mammals, 2nd ed., Samuel, W.M., Pybus, M.J., and Kocan, A.A. (Eds.), The Iowa State University Press, Ames, 559 pp. Measures, L.N., and Olson, M., 1999, Giardiasis in pinnipeds from Eastern Canada, J. Wildl. Dis., 35: 779–782. Mense, M.G., Dubey, J.P., and Homer, B.L., 1992, Acute hepatic necrosis associated with a sarcocystislike protozoa in a sea lion (Zalophus californianus), J. Vet. Diagn. Invest., 4: 486–491. Migaki, G., and Albert, T.F., 1980, Sarcosporidiosis in the ringed seal, J. Am. Vet. Med. Assoc., 177: 917– 918. Migaki, G., Sawa, T.R., and Dubey, J.P., 1990, Fatal disseminated toxoplasmosis in a spinner dolphin (Stenella longirostris), Vet. Pathol., 27: 463–464. Morimitsu, H., Kawano, K., Torihara, K., Kato, E., and Koono, M., 1992, Histopathology of eighth cranial nerve of mass stranded dolphins at Goto Islands, Japan, J. Wildl. Dis., 28: 656–658. Moser, M., and Rhinehart, H., 1993, The lungworm, Halocercus spp. (Nematoda: Pseudallidae) in cetaceans from California, J. Wildl. Dis., 29: 507–508. Munday, B.L., Mason, R.W., Hartley, W.F., Presidente, R.J.A., and Abendorf, D., 1978, Sarcocystis and related organisms in Australian wildlife, J. Wildl. Dis., 14: 417–433. Munro, R., and Synge, B., 1991, Coccidiosis in seals, Vet. Rec., 129: 179–180. Munro, R., Ruos, H., Corwell, C., and Gilmour, J., 1992, Disease conditions affecting common seals (Phoca vitulina) around the Scottish mainland, September–November 1988, Sci. Total Environ., 115: 67–82.
0839_frame_C18 Page 378 Tuesday, May 22, 2001 11:13 AM
378
CRC Handbook of Marine Mammal Medicine
Nascetti, G., Cianchi, R., Mattiucci, S., D’Amelia, S., Orecchia, P., Paggi, L., Brattey, J., Berland, B., Smith, J.W., and Bullini, L., 1993, Three sibling species within Contracaecum osculatum (Nematoda, Ascaridida, Ascaridoidea) from the Atlantic arctic-boreal region, reproductive isolation and host preferences, Int. J. Parasitol., 23: 105–120. Norman, R.J. de B., 1997, Tetraphyllidean cysticerci in the peritoneal cavity of the common dolphin, J. Wildl. Dis., 33: 891–895. Nozais, J.P., Mannevy, B., and Danis, M., 1996, Two cases of trichinosis from polar bear (Thalarctos maritimus) meat, Med. Mal. Infect., 26: 732–733. Odening, K., 1983, Sarkozysten einer antarktischen Robbe, Angew. Parasitol., 24: 197–200. Olsen, O.W., 1958, Hookworms, Uncinaria lucasi Stiles, 1901, in fur seals, Callorhinus ursinus (Linn.), on the Pribilof Islands, in Transactions of 23rd North American Wildlife Conference, Wildlife Management Institute, Washington, D.C., 152–175. Olson, M.E., Roach, P.D., Stabler, M., and Chan, W., 1997, Giardiasis in ringed seals from the western Arctic, J. Wildl. Dis., 33: 646–648. O’Shea, T.J., Homer, B.L., Greiner, E.C., and Layton, W.A., 1991, Nasitrema sp. associated encephalitis in a striped dolphin (Stenella coeruleoalba) stranded in the Gulf of Mexico, J. Wildl. Dis., 27: 706–709. Owens, C.G., and Kakulas, B.A., 1968, Sarcosporidosis in sperm whale, Aust. J. Sci., 31: 46–47. Paggi, L., Nascetti, G., Cianchi, R., Orecchia, P., Mattiucci, S., D’Amelia, S., Berland, B., Brattey, J., Smith, J.W., and Bullini, L., 1991, Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the North Atlantic and Norwegian and Barents Seas, Int. J. Parasitol., 21: 195–212. Pascual, S., Abollo, E., and Lopez, A., 2000, Elemental analysis of cetacean skull lesions associated with nematode infections, Dis. Aquat. Organisms, 42: 71–75. Pozio, K.V., Mortelmans, D.J., and Demeurichy, W., 1990, Characterization of trichinella isolate from polar bear, Ann. Soc. Belg. Med. Trop., 70: 131–135. Price, E.W., 1932, The trematode parasites of marine mammals, Proc. U.S. Nat. Mus., 81: 1–68. Raga, J.A., Casinos, A., Filella, S., and Raduan, M.A., 1982, Notes on cetaceans of the Siberian coasts. V. Crassicauda grampicola Johnston & Mawson, 1941 (Nematoda) cause of injuries in the pterygoids of some specimens of Grampus griseus, Saugetierk. Mitt., 30(4): 315–318. Rausch, R.L., 1953, Studies on the helminth fauna of Alaska XIII. Disease in the sea otter, with special reference to helminth parasites, Ecology, 34: 584–604. Ridgway, S., Geraci, J.R., and Medway, W., 1975, Diseases of pinnipeds, Rapp. P. V. Reun. Cons. Int. Explor. Mer, 169: 327–337. Schell, S.C., 1985, Trematodes of North America North of Mexico, University Press of Idaho, Moscow. Schmidt, G.D., 1986, CRC Handbook of Tapeworm Identification, CRC Press, Boca Raton, FL, 673 pp. Schulman, F.Y., and Lipscomb, T.P., 1997, Ciliate dermatitis in dolphins that died during the 1987–88 Atlantic bottlenose dolphin morbilliviral epizootic, Vet. Pathol., 34: 505. Smith, J.W., 1989, Ulcers associated with larval Anisakis simplex B (Nematoda: Ascaridoidea) in the forestomach of harbour porpoises Phocoena phocoena (L.), Can. J. Zool., 67: 2270–2276. 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. Eukar. Micro., 42: 260–268. Thomas, N.J., and Cole, R.A., 1996, Biology and status of the southern sea otter 6. The risk of disease and threats to the wild population, University of Michigan, School of Natural Resources and Environment, Endangered Species Update, 13: 23–27. Upton, S.J., Odell, D.K., Bossart, G.D., and Walsh, M.T., 1989, Description of two new species of Eimeria (Apicomplexa: Eimeriidae) from the Florida manatee, Trichechus manatus (Sirenia: Trichechidae), J. Protozool., 36: 87–90. Van Pelt, R.W., and Dieterich, R.A., 1973, Staphylococcal infection and toxoplasmosis in a young harbor seal, J. Wildl. Dis., 9: 258–262.
0839_frame_C18 Page 379 Tuesday, May 22, 2001 11:13 AM
Parasitic Diseases
379
Wallach, J.D., and Boever, W.J., 1983, Ursidae, in Disease of Exotic Animals: Medical and Surgical Management, W.B. Saunders, Philadelphia, 1159 pp. Woodcock, H.M., and Lodge, O., 1921, Protozoa: I Parasitic Protozoa, British Antarctic (“Terra Nova”) Expedition, 1910, Natural History Report, Zoology, 6: 1–24. Yablokov, A.V., Bel’kovich, V.M., and Borisov, V.I., 1972, Whales and Dolphins, Part 1. Kity i. Del’finy, Nauka, Moscow [translated from Russian by the U.S. Department of Commerce, Springfield, VA, 1974]. Young, P.C., and Lowe, D., 1969, Larval nematodes from fish of the subfamily Anisakinae and gastrointestinal lesions in mammals, J. Comp. Pathol., 79: 301–313. Zam, S.G., Caldwell, D.K., and Caldwell, M.C., 1971, Some endoparasites from small odontocete cetaceans collected in Florida and Georgia, Cetology, 2: 1–11.
0839_frame_C18 Page 380 Tuesday, May 22, 2001 11:13 AM
0839_frame_C19 Page 381 Tuesday, May 22, 2001 11:15 AM
IV Pathology of Marine Mammals
0839_frame_C19 Page 382 Tuesday, May 22, 2001 11:15 AM
0839_frame_C19 Page 383 Tuesday, May 22, 2001 11:15 AM
19 Clinical Pathology Gregory D. Bossart, Thomas H. Reidarson, Leslie A. Dierauf, and Deborah A. Duffield
Introduction Over the past decade, an information explosion has occurred in the field of veterinary clinical pathology. This is partially due to impressive technological advances in diagnostic equipment and analytical techniques, which are influencing and contributing to our knowledge of marine mammal clinical pathology. Marine mammal clinical laboratory medicine has also advanced. Baseline hematological, serum analyte, microbiological, cytological, and other specialized clinicopathological parameters have been established. During the last 10 years, new molecular techniques for disease diagnosis have also emerged. Today, the combination of clinical, laboratory, and other diagnostic data leads efficiently and accurately to diagnostic plans and subsequent treatment regimes. Clinical laboratory data and other diagnostic testing greatly complement the clinical examination of the patient. The correct evaluation of the health status of a marine mammal is dependent upon the integration of patient history, physical examination, and the results of diagnostic testing. In 1967, Wills and Halsted stated, “A physician who depends on the laboratory to make his diagnosis is probably inexperienced; one who says that he does not need a laboratory is uninformed. In either instance, the patient is in danger” (Coles, 1986). This statement is also applicable to marine mammal medicine. Selection and evaluation of diagnostic tests are basic to clinical decision making. In human medicine, 50 to 60 of the hundreds of readily available diagnostic tests account for 70% of the results generated by the modern hospital clinical laboratory (Borer, 1992). This implies that common diseases are evaluated using common tests, and that diagnoses are not usually made after only one test. However, a small number of well-chosen laboratory tests may help confirm a clinical suspicion, and/or rule out other candidates in the differential diagnosis.
Abnormalities and Artifacts Abnormal laboratory measurements are defined as those that occur outside the normal reference range. Major uncertainties that can be particularly troublesome in marine mammal medicine are the quality of data used to generate normal values, the variability inherent in various test mechanics, and variability of physiological systems of marine mammals. Determining whether borderline values represent normal physiological variation or disease can be challenging for marine mammal clinicians. Some indecision can be alleviated by using reference ranges adjusted by species, age, sex, husbandry practices (samples obtained from trained
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
383
0839_frame_C19 Page 384 Tuesday, May 22, 2001 11:15 AM
384
CRC Handbook of Marine Mammal Medicine
animals vs. those from animals that were captured or restrained), reproductive status, and laboratory methodology. It is imperative to collect and handle specimens properly for accurate clinical interpretation. Artifactual results from incorrectly handled specimens can result in incorrect differential diagnoses, especially if the clinician is unaware or ignorant of the presence of the artifact. The specific effects of improper handling of blood samples and various conditions present in blood samples are detailed under each serum analyte below. In general, for serum collection, set the red top tube (or other designated serum-tube variations) in an upright position and wait for clot retraction (approximately 10 min). The serum or plasma should be separated and removed from the cells within 15 to 20 min of collection, and kept refrigerated or frozen at or below −20°C. Prolonged contact of erythrocytes with serum decreases serum glucose concentrations at a rate of ∼10% per hour. Most biochemical parameters are stable in serum or plasma stored at 4°C (39.2°F) for at least 24 hours. Effects of storage on blood parameters vary with the anticoagulant used. For example, hematocrits in blood samples from harp seals (Pagophilus groenlandicus) collected into ethylene- diamine tetraacetate (EDTA) and stored for 7 days gradually increased over time, whereas hematocrits in samples collected into sodium heparin remained unchanged (Geraci and Engelhardt, 1974). In blood samples collected from harbor seals (Phoca vitulina), moderate to severe hemolysis significantly increased serum lactate dehydrogenase (LDH) activity and total protein, albumin, calculated globulin, and total bilirubin levels, whereas it decreased creatinine (Morgan et al., 1998). In bottlenose dolphins ( Tursiops truncatus), moderate hemolysis caused statistically significant increases in the concentrations of serum iron, LDH, potassium, and uric acid, and a decrease in creatinine. Severe hemolysis resulted in statistically significant increases in these analytes, as well as increases in alanine aminotransferase (ALT), calcium, globulins, albumin, and total protein, and decreases in total bilirubin and γ-glutamyltransferase (GGT) (Morgan et al., 1999). A study of clinically healthy young northern elephant seals (Mirounga angustirostris) reported erroneously elevated white blood cell counts due to automated count error, bone marrow contamination, poorly made blood smears, and faulty staining techniques (Goldstein et al., 1998). The effects of stress on clinical pathology in various marine mammal species are summarized in Chapter 13, Stress.
Blood Collection Sampling Equipment and Processing Phlebotomy and associated equipment should be preassembled and easily accessible prior to use. This equipment should include: 1. 2. 3. 4. 5.
Mechanical and/or chemical restraint equipment Sterile needles (of varying sizes and gauges, including butterfly sets) Sterile syringes of various sizes Povidone-iodine solution (spray bottle or swabs) and isopropyl alcohol Blood collection tubes (check expiration dates of anticoagulant tubes) a. EDTA or heparin vacutainer tubes for complete blood counts (CBC) b. Serum chemistries-serum separator tubes for all species except cetaceans; fibrin-containing vacutainer tubes for cetaceans c. Coagulation profiles-citrate containing vacutainer tubes 6. Pencil/indelible ink pen for tube labeling 7. Laboratory submission forms
0839_frame_C19 Page 385 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
385
In general, either apply alcohol or perform a standard surgical scrub before phlebotomy or any injection, to avoid the potential for iatrogenic abscessation. The veterinarian must be familiar with collection protocols for the type of test desired, and provide all pertinent information on the laboratory sample submission form.
Blood Collection Sites Cetaceans
See Figure 1 and Chapter 40, Cetaceans. 1. 2. 3. 4.
Caudal vascular bundle of flukes Peduncle Dorsal fin vein Pectoral flipper vein
The central tail, dorsal fin, and pectoral flipper veins are periarterial venous retia; consequently, samples are typically mixed venous/arterial blood. The dorsal fin vein is an excellent venipuncture site for chronically ill animals when multiple samplings are required, as it is more easily held above water and the animal may resist less. A 1.0-in., 18- to 22-gauge needle (or butterfly set) for calves and juveniles and a 1.0- to 1.5-in., 18- to 20-gauge needle (or butterfly set) for adults are recommended. The caudal peduncle site requires a long needle (1.5 to 3.5 in.). Otariids
See Figure 2 and Chapter 41, Seals and Sea Lions. 1. 2. 3. 4.
Caudal gluteal vein Interdigital veins of hind flipper Precaval vein Jugular vein
A 1.0-in., 18- to 20-gauge needle (or a butterfly set) for pups or juveniles and a 1.5- to 2.0-in., 18to 20-gauge needle for adults are recommended. The caudal gluteal vein is used routinely in California sea lions (Zalophus californianus), but can be hard to locate (see Chapter 41, Seals and Sea Lions). The interdigital vessels are often visible, but may be more readily visualized by use of a tourniquet or warm bag under the flipper. The precaval vein is the route of choice for emergency administration of drugs, but is not recommended for routine venipuncture. The precava on the sea lion forms by the bilateral fusion of the jugulars about midpoint on the manubrium; the ventral aspect of this venous sinus actually attaches to the dorsal aspects of the first two sternabrae. The sinus is about a finger’s width (on each side) wider than the sternum between the first two ribs. The jugular vein is not palpable, and the external jugulars are smaller than the internal jugular. Phocids
See Figure 3 and Chapter 41, Seals and Sea Lions. 1. Epidural vertebral vein 2. Interdigital veins of hind flipper
The needle length and gauge are the same as those used with otariids. Odobenids
See Chapter 42, Walruses. 1. Epidural vertebral sinus, as in phocids 2. Interdigital veins of the hind flipper
0839_frame_C19 Page 386 Tuesday, May 22, 2001 11:15 AM
386
CRC Handbook of Marine Mammal Medicine
PAVR -periarterial venous rete
Each artery is relatively deep and surrounded by veins PAVR (deep, along midline)
Blubber
3 Superficial dorsal fin vv. 2
Superficial caudal peduncle vv.
Rommel 2000 Pelvic vestige 4
Superficial flipper vv.
Superficial fluke vv.
PAVR (deep)
PAVR, (shallow, dorsal & ventral)
Muscle
Bone Note that the vascular bundle is arteriovenous
1
Caudal vascular bundle Chevron canal
Aorta Heart
Vena cava Chevron bones
FIGURE 1 Veins used for blood collection in the bottlenose dolphin. (© S.A. Rommel. Used with permission of the illustrator.)
0839_frame_C19 Page 387 Tuesday, May 22, 2001 11:15 AM
387
Clinical Pathology
Dorsal
L. Lateral Neural spine, Sa2
Caudal
1 gluteal v.
Neural spine, Sa2
Caudal
lliac crest
Calcaneus Patella
1 gluteal v. Calcaneus
lliac crest
Patella
2 Interdigital vv.
Nerve plexus Nerve plexus
L. lnguinal lymph node
L. Flipper L. Testis
L. Testis Ventral
L. lnguinal lymph node
L. atrium Cartilagenous sternal ribs (cut)
L. ventricle
L. External 3 jugular v.
Pericardium
Manubrium
3
R. External jugular v.
4
Cranial vena cava
Sternum
Rommel 2000
FIGURE 2 Veins used for blood collection in the California sea lion. (© S.A. Rommel. Used with permission of the illustrator.)
Manatees (Trichechus manatus)
See Figure 4 and Chapter 43, Manatees. 1. Brachial vascular bundle (preferred site) 2. Caudal vascular bundle (or fluke)
A 1.0-in., 20- to 22-gauge needle (or butterfly set) for calves and a 1.0- to 1.5-in., 18- to 20-gauge needle (or butterfly set) for adults are recommended. Blood collected from the brachial vascular bundle will be mixed arteriovenous blood. Sea Otters (Enhydra lutris)
See Figure 5 and Chapter 44, Sea Otters. 1. Femoral vein (medial aspect at the proximal one third of femur) 2. Saphenous vein 3. Jugular vein
A 1.0-in., 19- to 20-gauge needle can be used for animals of all ages.
0839_frame_C19 Page 388 Tuesday, May 22, 2001 11:15 AM
388
CRC Handbook of Marine Mammal Medicine
Neural spine Vertebral body
Epidural sinus 1
Epidural 1 sinus
Spinal cord
Lumbar vertebrae 3-4 Calcaneus
L.lumbar vena cava
Abdominal-wall plexus Saphenousfemoral
Gluteal anastomosis
Calcaneus
Inguinal plexus
Pudendo epigastric anastomosis
Saphenous femoral vein (surrounding saphenous artery)
Plantar 2 network
Rommel 2000
FIGURE 3 Veins used for blood collection in the harbor seal. (© S.A. Rommel. Used with permission of the illustrator.)
0839_frame_C19 Page 389 Tuesday, May 22, 2001 11:15 AM
389
Clinical Pathology
Lateral nerve
1
vein
median nerve ulnar nerve
Humerus
H
ulnar nerve median nerve
vascular bundle
vascular bundle Lateral nerves
Radius
vein
vein
Rommel 2000
nerves
U R
median nerve Lateral
Ulna
vascular bundle
1
vein
nerves
nerves
U
R
Lateral
vein
nerves U
vein
1
vascular bundle
nerves
vein nerves
R
vascular bundle
1
muscle
nerves Note that vascular bundles are arterio-venous.
Caudal vascular bundle
2
Aorta
Heart
Rommel 2000 Vertebra
Chevron bones
Muscle
Chevron canal
FIGURE 4 Veins used for blood collection in the manatee. (© S.A. Rommel. Used with permission of the illustrator.)
0839_frame_C19 Page 390 Tuesday, May 22, 2001 11:15 AM
390
CRC Handbook of Marine Mammal Medicine
Ventral
3 L. External jugular v.
Tibia Catilagenous sternal ribs
Public Symphysis
L. Femur
L. Mandibular lymph nodes
L. lliac crest
L. Mandibular gland
R. Calcaneus
Mandible Hyoid venous arc R. Angle of mandible
Heart R. Femoral v.
R. External Tip of auditory sternum meatus
1 R. Saphenous v.
Rommel 2000
2
FIGURE 5 Veins used for blood collection in the sea otter. (© S.A. Rommel. Used with permission of the illustrator.)
Polar Bears (Ursus maritimus) 1. Cephalic vein 2. Femoral vein
A 1.5-in., 18- to 19-gauge needle can be used for animals of all ages (see Chapter 45, Polar Bears).
Hematology (CBC) Normal ranges for hematology and serum biochemistry parameters from some marine mammal species regularly examined by veterinarians are presented in Tables 1 through 3. Unless otherwise noted, the hematology and chemistry values are from clinically normal, healthy animals in good body condition. Because of the large number of values for individual cetaceans from SeaWorld, ranges were determined by calculating 25 to 75% quartiles around individual medians. All other ranges were calculated as ±2 standard deviations around the means. Values outside this range were eliminated, means and standard deviations were then recalculated, and new ranges were determined. Table 4 provides references on blood parameters in some other species. Clinicians must recognize that the blood values provided in this chapter remain guidelines; baselines still need to be established for each individual marine mammal before being confident that any particular value is abnormal. Additionally, captivity demands physiological adjustments that may be reflected in clinical laboratory values. The degree to which these adjustments affect blood values needs further study. Nothing can replace the process of establishing semiannual baseline values for each individual animal during regular, routine, and scheduled physical examinations. The ability to diagnose disease and monitor responses to treatment is tied to deviations from an individual animal’s own baseline data.
0839_frame_C19 Page 391 Tuesday, May 22, 2001 11:15 AM
391
Clinical Pathology
TABLE 1 Ranges of Hematology and Biochemistry Values from Nine Species of Free-Ranging and Captive Cetaceans
Parameter 6
3
RBC (10 /mm ) Hb (g/dl) HCT (%) MCV (fl) MCH (pg) MCHC (g/dl) 3 3 Platelets (10 /mm ) Reticulocytes (%) nRBC ESR (at 60 min) Leukocytes/µl Neutrophil (band) Neutrophil (mature) Lymphocyte Monocyte Eosinophil Basophil Serum proteins (g/dl) Albumin (g/dl) Globulin (g/dl) Glucose (mg/dl) BUN (mg/dl) Creatinine (mg/dl) Bilirubin T/D (mg/dl) Cholesterol (mg/dl) Alkaline phos (U/l) ALT (U/l) AST (U/l) GGT (U/l) CK (U/l) LDH (U/l) Calcium (mg/dl) Phosphorus (mg/dl) Sodium (mEq/l) Potassium (mEq/l) Chloride (mEq/l) Iron (mcg/dl) Fibrinogen (mg/dl)
Captive Commerson’s Dolphins (Cephalorhynchus a commersoni) (n = 10, sample = 196)
Captive, after Rehabilitation, Common Dolphins (Delphinus a delphis) (n = 2, sample = 44)
Captive (n = 13, sample = 216)
Free-Ranging (n, sample = 145)
4.3–5.5 15.0–19.0 43–53 94–104 33–37 34–36 120–250 0.6–2.4 0–3 0 4000–8000 0 1150–3250 1260–2420 150–270 890–2200 0 5.6–7.0 3.4–4.0 2.0–3.3 80–130 33–43 0.5–0.9 0.1–0.2/N.D. 130–200 90–290 40–140 160–300 28–50 130–300 300–500 8.0–9.5 3.5–6.0 154–159 3.5–4.6 118–123 120–230 150–250
4.6–4.9 16.1–19.4 46–55 100–114 35–40 34–36 55–100 0.8–1.4 0 0 4570–4900 0 2590–4150 380–850 120–350 620–1280 0 6.3–7.3 3.9–4.7 1.8–3.0 91–119 22–46 0.9–1.3 0.1–0.9/N.D. 130–200 202–580 49–84 191–236 37–44 c N.D. 354–568 8.8–9.6 2.8–5.3 152–159 4.0 120–121 184–270 N.D.
3.0–3.4 19.0–22.0 50–60 163–185 59–66 36–38 60–130 0.3–0.8 0–1 0–9 5000–9500 0 2580–5520 1100–4150 220–780 90–640 0 5.7–7.3 4.1–4.7 1.6–2.8 84–124 47–59 1.2–1.6 0.1/N.D. 170–260 100–220 3–10 45–80 16–36 80–180 100–220 9.1–10.6 4.5–5.8 153–159 3.5–4.1 111–120 195–380 70–130
3.5–3.7 21.2–21.9 58–60 159–164 58–61 36–37 N.D. N.D. N.D. N.D. 9200–10900 0 3700–4400 3600–4700 370–530 2400–3300 0 7.9–8.2 4.1–4.3 3.7–4.0 108–114 52–55 0.3–3.0 0.2–0.4/N.D. 174–194 166–211 7–15 70–83 15–18 149–175 218–574 10.4–10.8 7.9–8.3 162–165 4.5–4.8 113–114 438–551 N.D.
Belugas a (Delphinapterus leucas) a
b
(Continued)
Evaluation of Erythrocytes Indices Erythrocyte or red blood cell (RBC) parameters in blood are quantified by cell counting (RBCs/µl), determining blood hemoglobin (Hb) content (g/dl), and hematocrit (HCT) or packed cell volume (PCV) as a percentage. Essentially all Hb is present within RBCs, so the RBC count, HCT, and Hb content complement each other. All RBC indices must be properly assessed to evaluate abnormalities
0839_frame_C19 Page 392 Tuesday, April 9, 2002 1:40 PM
392
CRC Handbook of Marine Mammal Medicine
TABLE 1 Ranges of Hematology and Biochemistry Values from Nine Species of Free-Ranging and Captive Cetaceans (continued)
Parameter 6
3
RBC (10 /mm ) Hb (g/dl) HCT (%) MCV (fl) MCH (pg) MCHC (g/dl) 3 3 Platelets (10 /mm ) Reticulocytes (%) nRBC ESR (at 60 min) Leukocytes/µl Neutrophil (band) Neutrophil (mature) Lymphocyte Monocyte Eosinophil Basophil Serum proteins (g/dl) Albumin (g/dl) Globulin (g/dl) Glucose (mg/dl) BUN (mg/dl) Creatinine (mg/dl) Bilirubin T/D (mg/dl) Cholesterol (mg/dl) Alkaline phos (U/l) ALT (U/l) AST (U/l) GGT (U/l) CK (U/l) LDH (U/l) Calcium (mg/dl) Phosphorus (mg/dl) Sodium (mEq/l) Potassium (mEq/l) Chloride (mEq/l) Iron (mcg/dl) Fibrinogen (mg/dl)
Captive Pilot Whales (Globicephala a macrorhynchus) (n = 2, sample = 74)
Captive Pacific White-Sided Dolphins (Lagenorhynchus a obliquidens) (n = 9, sample = 373)
Captive Killer Whales a (Orcinus orca) (n = 19, sample = 1761)
Captive, False Killer Whales (Pseudorca a crassidens) (n = 5, sample = 81)
3.3–3.7 15.1–16.0 43–45 123–129 43–46 34–36 70–90 0.7–1.2 0 16–52 4720–6500 0 2930–4360 660–2080 190–460 240–870 0 5.3–6.0 2.9–3.3 2.2–3.0 98–106 46–55 2.0–2.4 0.1/N.D. 187–288 143–243 26–69 170–317 39–41 55–80 425–505 7.8–8.4 4.3–4.8 153–154 3.7–4.2 118–119 108–179 280–445
4.5–5.3 17–20 47–57 90–98 32–36 35–37 100–150 0.8–2.5 0–2 0 3000–7000 0 1250–3730 390–1390 80–240 720–1910 0 5.8–6.8 3.0–3.8 2.4–3.0 90–130 30–43 0.7–1.1 0.1–0.2/N.D.* 100–175 200–570 30–90 180–270 25–70 80–150 350–550 7.8–8.8 3.0–6.0 153–158 3.3–3.8 112–120 120–240 163–240
3.5–4.3 13.5–15.5 40–46 105–115 35–40 34–36 120–230 0.7–2.5 0 0–2 4000–8000 0 2380–8080 520–1850 140–420 10–160 0 5.5–7.5 3.0–3.7 2.0–3.4 110–135 30–50 0.8–2.0 0.1–0.2/N.D. 140–280 100–700 10–40 35–60 8–25 60–230 280–400 8.0–9.5 5.0–7.0 154–158 3.5–4.5 115–125 50–130 170–330
3.4–4.6 13.7–17.6 39–51 112–119 40–42 34–36 78–150 0.5–0.8 0–1 3–29 5000–9000 0 2280–5040 990–2490 120–400 410–1540 0 5.6–6.6 3.5–3.9 2.2–2.8 94–134 32–43 1.0–2.1 0.1/N.D. 170–400 380–700 6–16 130–230 25–46 59–143 260–370 7.6–8.8 4.4–6.4 152–157 3.7–4.4 120–124 100–200 230–320
adequately. Modern electronic cell counters may calculate the HCT using the measured RBC count and mean cell volume (MCV). It is important for the clinician to remember that with marine mammal blood, the RBC count, MCV, and HCT are accurate only if measured using a calibrated electronic cell counter to accommodate for the variably sized RBCs of marine mammals; Hb content is measured spectrophotometrically by the cyanmethemoglobin method. Because of trapped plasma, the HCT determined by centrifugation of blood in a microhematocrit tube may be 1 to 3% higher than when determined electronically and with RBC size-adjusted counts. It is nevertheless recommended that the microhematocrit method for HCT determination
0839_frame_C19 Page 393 Tuesday, April 9, 2002 1:40 PM
393
Clinical Pathology
TABLE 1 Ranges of Hematology and Biochemistry Values from Nine Species of FreeRanging and Captive Cetaceans (continued) Bottlenose Dolphins a (Tursiops truncatus) a
Parameter 6
3
RBC (10 /mm ) Hb (g/dl) HCT (%) MCV (fl) MCH (pg) MCHC (g/dl) 3 3 Platelets (10 /mm ) Reticulocytes (%) nRBC ESR (at 60 min) Leukocytes/µl Neutrophil (band) Neutrophil (mature) Lymphocyte Monocyte Eosinophil Basophil Serum proteins (g/dl) Albumin (g/dl) Globulin (g/dl) Glucose (mg/dl) BUN (mg/dl) Creatinine (mg/dl) Bilirubin T/D (mg/dl) Cholesterol (mg/dl) Alkaline phos (U/l) ALT (U/l) AST (U/l) GGT (U/l) CK (U/l) LDH (U/l) Calcium (mg/dl) Phosphorus (mg/dl) Sodium (mEq/l) Potassium (mEq/l) Chloride (mEq/l) Iron (mcg/dl) Fibrinogen (mg/dl)
c
Captive (n = 38, sample = 1150)
Free-Ranging (n, sample = 36)
3.0–3.7 13.5–15.5 38–44 115–135 38–48 34–36 80–150 1.0–2.3 1–4 4–17 5000–9000 0 3230–4850 840–1660 140–350 530–1020 0 6.0–7.8 4.3–5.3 1.3–2.5 90–170 42–58 1.0–2.0 0.1–0.2/N.D. 150–260 300–1300 28–60 190–300 30–50 100–250 350–500 8.5–10.0 4.0–6.0 153–158 3.2–4.2 113–125 120–340 170–280
3.1–4.0 12.7–15.5 37–47 111–127 36–43 32–35 92–217 N.D. 0 N.D. 5600–12400 0 2540–6140 520–2420 80–610 740–4530 0–30 6.4–8.8 2.9–3.7 3.1–5.5 62–139 45–72 1.0–2.1 0.1–0.4/N.D. 137–235 51–610 9–33 133–318 17–31 N.D. 324–538 8.2–9.4 3.2–7.2 151–158 3.2–4.4 108–118 74–176 N.D.
Stranded Juvenile Gray Whale (Eschrichtius a robustus) (n = 1, sample = 14) 3.0–4.0 13.0–16.0 39–47 129–142 43–48 33–34 60–304 0–2 0 48–100 2700–10710 0–30 1670–9250 300–1120 40–910 0–30 0 4.0–7.0 3.0–4.0 1.0–3.0 47–147 21–75 1.0–2.0 0–0.2/N.D. 136–1470 1263–3017 3–12 41–113 2–52 107–255 120–584 8.0–11.0 3.7–9.0 146–154 4.0–5.0 106–115 54–328 277–517
a
Data contributed by SeaWorld clinical laboratories. Data contributed by D. St. Aubin and S. De Guise. c Data contributed by R. Wells and H. Rhinehart, Sarasota Dolphin Research Program, Chicago Zoological Society. N.D. = not determined. b
be done for each clinical sample, because of the potential for electronic cell counter error. The microhematocrit method is simple and quick, and if done by centrifugation, can also indicate the total leukocyte count (buffy coat height) and the presence of hyperbilirubinemia, lipemia, and hemolysis. In addition, a plasma protein determination can be made from the microhematocrit tube plasma, providing important clinical information from the simultaneous interpretation of the HCT and plasma protein concentration (see Serum Proteins below).
3
RBC (10 /mm ) Hb (g/dl) HCT (%) MCV (fl) MCH (pg) MCHC (g/dl) 3 3 Platelets (10 /mm ) Reticulocytes (%) nRBC ESR (at 60 min) Leukocytes/µl Neutrophil (band) Neutrophil (mature) Lymphocyte Monocyte Eosinophil Basophil Serum proteins (g/dl) Albumin (g/dl) Globulin (g/dl) Glucose (mg/dl) BUN (mg/dl) Creatinine (mg/dl) Bilirubin T/D (mg/dl) Cholesterol (mg/dl) Alkaline phos (U/l) ALT (U/l) AST (U/l) GGT (U/l)
6
Parameter
Rehabilitated Weanling Northern Elephant Seals (Mirounga angustirostris)b (n, sample = 25) 2.6–3.3 16.2–24.2 46–61 170–185 63–74 37–41 355–652 N.D. N.D. N.D. 10600–28500 0–527 7176–20714 2385–7630 400–3400 100–795 0–160 6.6–9.9 3.2–3.9 3.1–6.1 128–189 33–76 0.2–0.8 0.1–0.6/0.1–0.2 100–347 108–454 18–65 39–117 22–136
Free-Ranging Pup Steller Sea Lions (Eumetopias jubatus)a (n, sample = 25)
N.D.d 10.6–19.5 31–45 N.D. N.D. N.D. N.D. N.D. N.D. N.D. 5400–28760 3985–17972 4130–18560 1320–6900 50–2480 45–1255 0–130 5.6–6.9 3.5–4.5 1.7–2.6 138–232 8–29 0.4–0.8 0.1–0.4/N.D. 147–272 87–206 16–68 13–51 32–83 3.1–3.4 14.9–19.2 43–52 135–153 48–56 35–37 100–262 0.2 0 31–107 5300–9510 0–11 2945–6680 995–2070 210–660 210–590 0 6.5–7.1 2.2–3.1 3.7–4.6 103–111 37–45 0.9–1.3 0.1–0.2/N.D. 162–276 25–67 15–55 43–152 109–443
Captive Pacific Walrus (Odobenus rosmarus)c (n = 3, sample = 42) 3.6–5.4 13.8–21.0 32–58 92–114 33–44 33–41 373–1164 N.D. 1–2 N.D. 5900–24610 60–340 3420–13960 700–3570 70–1900 70–1860 100–1060 6.5–9.0 2.7–3.7 2.7–6.4 101–186 34–60 0.3–1.1 0.2–0.8/0–0.2 168–399 52–224 30–95 50–159 8–23
Rehabilitated Weanlingb (n, sample = 42) 4.1–5.5 17.0–23.5 48–66 102–135 36–48 35–37 155–555 0.1–0.9 0–1 0–45 4800–14250 0–140 3430–1030 550–3830 160–1250 50–810 0–120 6.1–8.8 2.5–3.3 3.2–6.0 101–172 28–53 0.5–2.5 0.1–0.5/N.D. 166–351 11–176 8–75 29–168 3–15
Captive Adultc (n = 33, sample = 184)
Harbor Seals (Phoca vitulina)
TABLE 2 Ranges of Hematology and Biochemistry Values in Four Species of Free-Ranging and Captive Pinnipeds
2.9–4.0 12–18 34–50 94–115 33–41 31–39 208–720 N.D. 1.0–2.5 N.D. 9780–22800 0–680 5140–16680 1180–8380 0–1600 0–1470 0 7.7–10.7 2.9–4.0 4.1–7.4 62–135 20–88 0–1.4 0.1–1.1/0–0.1 102–269 9–175 4–92 13–166 2–157
Stranded Weanling (n = 73, sample = 201)
b
c
3.7–5.3 13–22 38–59 97–116 33–42 34–37 158–355 0.1–0.6 0.0–1.3 0–76 3400–11380 0–10 2120–8220 370–3190 45–830 0–630 0–35 6.1–8.5 2.7–3.6 3.1–5.6 71–203 14–38 1.1–2.6 0.1–0.4/N.D. 165–507 34–175 19–71 12–66 20–123
Captive Adult (n = 23, sample = 54)
California Sea Lions (Zalophus californianus)
0839_frame_C19 Page 394 Tuesday, May 22, 2001 11:15 AM
394 CRC Handbook of Marine Mammal Medicine
N.D. N.D. 10.2–12.1 5.3–9.1 141–151 3.8–5.2 102–109 N.D. N.D.
N.D. 223–542 10.2–12.8 6.6–9.9 143–154 4.7–6.1 98–107 34–249 N.D.
b
Data contributed by T. Loughlin. Data contributed by The Marine Mammal Center clinical laboratory. c Data contributed by SeaWorld clinical laboratories. d N.D. = not determined.
a
CK (U/l) LDH (U/l) Calcium (mg/dl) Phosphorus (mg/dl) Sodium (mEq/l) Potassium (mEq/l) Chloride (mEq/l) Iron (mcg/dl) Fibrinogen (mg/dl)
21–80 236–340 8.8–9.3 4.4–6.4 146–151 4.1–4.6 112–116 98–147 250–358
N.D. 339–1303 8.7–11.8 5.9–9.8 148–158 4.1–6.0 99–111 53–451 N.D.
34–4572 167–1000 7.8–11.1 1.9–7.4 147–158 3.1–17.3 96–116 49–218 120–324
N.D. 350–1684 8.5–11.2 4.9–8.9 143–162 4.1–5.7 101–115 57–219 N.D.
77–2351 333–1054 8.8–11.5 1.8–7.8 149–156 3.7–5.0 105–113 72–185 151–321
0839_frame_C19 Page 395 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology 395
0839_frame_C19 Page 396 Tuesday, May 22, 2001 11:15 AM
396
CRC Handbook of Marine Mammal Medicine
TABLE 3 Ranges of Hematology and Biochemistry Values in Free-Ranging Manatees, Polar Bears, and Sea Otters
Parameter 6
3
RBC (10 /mm ) Hb (g/dl) HCT (%) MCV (fl) MCH (pg) MCHC (g/dl) 3 3 Platelets (10 /mm ) Reticulocytes (%) nRBC ESR (at 60 min) Leukocytes/µl Neutrophil (band) Neutrophil (mature) Heterophil Lymphocyte Monocyte Eosinophil Basophil Serum proteins (g/dl) Albumin (g/dl) Globulin (g/dl) Glucose (mg/dl) BUN (mg/dl) Creatinine (mg/dl) Bilirubin T/D (mg/dl) Cholesterol (mg/dl) Alkaline phos (U/l) ALT (U/l) AST (U/l) GGT (U/l) CK (U/l) LDH (U/l) Calcium (mg/dl) Phosphorus (mg/dl) Sodium (mEq/l) Potassium (mEq/l) Chloride (mEq/l) Iron (mcg/dl) Fibrinogen (mg/dl)
Florida Manatee (Trichechus manatus a latirostris) (n, sample = 23)
Polar Bears b (Ursus maritimus) (n, sample = 18–151)
Juvenile (n, sample = 35–37)
Adult (n, sample = 61–62)
2.4–3.4 9.8–13.2 30–40 122–149 38–46 30–33 195–412 0–4 0 7.0–8.0 4000–11800 0 d N.D. 960–8590 960–8590 0–1020 0 0 6.2–8.6 3.6–5.9 2.6–2.7 56–117 6.4–16.0 0.4–2.1 0–0.1/N.D. 107–328 64–183 6–30 5–28 39–64 79–302 94–372 10.1–12.2 3.0–8.0 142–157 4.2–6.6 90–103 50–199 N.D.
5.40–8.24 12.9–17.5 36–53 62–73 21–26 33–37 107–513 N.D. N.D. N.D. 3300–10,800 175–575 1955–6405 0 640–2095 370–1210 130–420 25–85 4.9–9.4 2.9–4.6 2.0–5.7 61–168 3.9–64.7 0.6–2.7 0–0.2/N.D. 151–638 0–31 8–29 16–90 16–176 54–384 N.D. 5.4–10.7 4.3–7.6 133–148 3.1–5.2 88–114 N.D. N.D.
4.6–6.3 14.6–22.3 44–57 95–118 32–37 32–35 N.D. N.D. N.D. N.D. 6100–14400 0–240 3200–10,220 0 1340–5230 0–490 0–230 N.D. 5.3–9.4 2.0–3.5 N.D. 60–191 25.0–86.0 0.4–3.2 N.D. 115–382 31–312 68–366 N.D. N.D. N.D. N.D. 6.4–11.8 3.5–13.0 141–160 4.1–5.6 N.D. N.D. N.D.
4.4–6.2 13.6–21.8 40–66 97–118 32–40 32–37 N.D. N.D. N.D. N.D. 6700–14400 0–260 4110–11,820 0 1510–5400 0–420 0–320 N.D. 4.9–7.8 2.4–3.7 N.D. 67–161 31.0–90.0 0.5–1.8 N.D. 198–288 57–259 71–248 N.D. N.D. N.D. N.D. 6.4–10.8 4.5–10.5 140–159 4.0–5.5 N.D. N.D. N.D.
Sea Otters (Enhydra lutris)
c
a
Data contributed by G. Bossart and D. Odell. Data contributed by M. Cattet and N. Caulkett. c Data contributed by T. Williams. d N.D. = not determined. b
Red blood cell size and Hb concentrations are increased in most marine mammals compared with terrestrial mammals (Reidarson et al., 2000). For example, the hooded seal’s (Cystophora cristata) RBCs are 60% larger and have a 30% greater Hb concentration than human RBCs (Clausen and Ersland, 1969); Sirenian RBCs are larger than those of the bottlenose dolphin
0839_frame_C19 Page 397 Tuesday, May 22, 2001 11:15 AM
397
Clinical Pathology
TABLE 4 Additional References for Normal Hematology and Clinical Chemistries in Marine Mammals Listed by Species (in alphabetical order) Species
Reference Cetaceans (baleen)
Bowhead whale (Balaena mysticetus)
Heidel et al., 1996 Cetaceans (toothed)
Beluga (Delphinapterus leucas)
St. Aubin and Geraci, 1979; Medway and Geraci, 1986; Cornell et al., 1988 Medway and Geraci, 1964; 1965; 1986; Ridgway, 1972; Englehardt, 1979; Asper et al., 1990; Morgan et al., 1999 Ridgway, 1972
Bottlenose dolphin (Tursiops truncatus)
Boutu (Inia geoffrensis) Dall’s porpoise (Phocoenoides dalli) Killer whale (Orcinus orca)
Ridgway, 1972
Pacific white-sided dolphin (Lagenorhynchus obliquidens) Pilot whale (Globicephala sp.)
Ridgway, 1972; Medway and Geraci, 1986; Cornell, 1993 Ridgway, 1972; Medway and Geraci, 1986 Ridgway, 1972; Medway and Geraci, 1986
Pinnipeds (otariids) Australian sea lion (Neophoca cinerea)
Cargill et al., 1979; Needham et al., 1980
California sea lion (Zalophus californianus)
Hubbard, 1968; Ridgway, 1972; Englehardt, 1979; Roletto and Dougherty, 1983; Medway and Geraci, 1986; Roletto, 1993 Lander et al., 2000
Guadalupe fur seal (Arctocephalus townsendi) Northern fur seal (Callorhinus ursinus) Steller sea lion (Eumetopias jubatus)
Hubbard, 1968; Hunter and Madin, 1978 Hubbard, 1968; Castellini et al., 1993; Rea et al., 1998 Pinnipeds (phocids)
Baikal seal (Pusa sibirica) Crabeater seal (Lobodon carcinophagus)
Ronald and Kay, 1983; Morgan et al., 1998 Tyler, 1960; Seal et al., 1971 (Continued)
0839_frame_C19 Page 398 Tuesday, May 22, 2001 11:15 AM
398
CRC Handbook of Marine Mammal Medicine
TABLE 4 Additional References for Normal Hematology and Clinical Chemistries in Marine Mammals Listed by Species (in alphabetical order) (continued) Species
Reference
Gray seal (Halichoerus grypus)
Greenwood et al., 1971; Medway and Geraci, 1986; Schweigert, 1993; Hall, 1998 Hubbard, 1968; Ridgway, 1972; Englehardt, 1979; McConnell and Vaughan, 1983; Roletto and Dougherty, 1983; Kuiken, 1985; Medway and Geraci, 1986; Williams et al., 1994; de Swart et al., 1995 Ronald et al., 1969; Vallyathan et al., 1969; Geraci, 1971; Worthy and Lavigne, 1982 Banish and Gilmartin, 1988
Harbor seal (Phoca vitulina)
Harp seal (Pagophilus groenlandicus)
Hawaiian monk seal (Monachus schauinslandi) Hooded seal (Cystophora cristata) Northern elephant seal (Mirounga angustirostris)
Clausen and Ersland, 1969 Hubbard, 1968; Ridgway, 1972; Englehardt, 1979; Costa and Ortiz, 1982; Roletto and Dougherty, 1983; Wickham et al., 1990; Roletto, 1993; Gulland et al., 1996; Beckmen et al., 1997 Bryden and Linn, 1969; Seal et al., 1971; Lane et al., 1972 Lenfant et al., 1970
Southern elephant seal (Mirounga leonina) Ribbon seal (Phoca fasciata) Ringed seal (Phoca hispida) Weddell seal (Leptonychotes weddellii)
Geraci and Smith, 1975; Geraci et al., 1979 Kooyman and Drabek, 1968; Schumacher et al., 1992 Manatees
Florida manatee (Trichechus manatus latirostris) Antillean manatees (Trichechus manatus manatus)
White et al., 1976; Medway and Geraci, 1986; Walsh and Bossart, 1999 Converse et al., 1994
0839_frame_C19 Page 399 Tuesday, May 22, 2001 11:15 AM
399
Clinical Pathology
TABLE 4 Additional References for Normal Hematology and Clinical Chemistries in Marine Mammals Listed by Species (in alphabetical order) (continued) Species
Reference Polar Bears
Ursus maritimus
Seal et al., 1967; Lee et al., 1977 Sea Otters
California sea otter (Enhydra lutris)
Williams and Pulley, 1983; Wickham et al., 1990; Williams et al., 1994
(White et al., 1976); RBC size and blood volume are not directly proportional to body weight in marine mammals (Ridgway et al., 1984), as in some terrestrial vertebrates. Sea otter RBC, Hb, and HCT values are similar to those in pinnipeds and cetaceans, reflecting adaptations to the marine environment (Williams and Pulley, 1983). Free-ranging polar bears have lower Hb and HCT than captive polar bears (Lee et al., 1977) When compared with terrestrial mammals, marine mammals have the lowest overall RBC numbers (Lenfant, 1969). However, the RBC indices, heart weights, and blood volumes of marine mammals are generally higher, as adaptations for the additional oxygen-carrying capacity necessary for a diving animal (Ridgway, 1972). RBC counts are higher in neonates than in adult marine mammals; RBC counts decrease as the animal grows and learns to dive (Bryden and Linn 1969; Geraci, 1971; Roletto and Dougherty, 1983). RBC counts decrease and MCV increases at ∼5 to 8 weeks of age in elephant seals and harbor seals, corresponding to exposure to water, swimming, and diving (Roletto and Dougherty, 1983). RBC parameters of captive harbor seals are lower than in free-ranging harbor seals, presumably because of the lack of deep water diving in captivity (McConnell and Vaughan, 1983). Lower red cell counts were observed in wild harbor seals when their diet shifted from clupeids to alternative prey (Thompson et al., 1997).
Anemia Absolute anemia is a decrease in RBC mass and HCT; Hb and RBC counts are usually below reference ranges. The hydration status of the animal is critical, because anemia can be masked by concomitant dehydration. Alternatively, overhydration can produce a relative anemia, which is characterized by low RBC parameters with normal total RBC mass. The classification of anemia can aid in the determination of disease etiology and prognosis. Anemia can occur from blood loss, increased RBC destruction, or decreased RBC production. Factors that help categorize anemia include RBC morphology, reticulocyte counts, RBC indices, serum iron measurements, and bone marrow evaluation. A useful clinical approach is to determine if an anemia is regenerative (evidence of bone marrow erythropoiesis) or nonregenerative (no evidence of bone marrow erythropoiesis). Increased polychromasia is usually present in regenerative anemias. Polychromasia indicates the presence of polychromatophilic RBCs, which are reticulocytes that stain bluish-red as a result of the combined
0839_frame_C19 Page 400 Tuesday, May 22, 2001 11:15 AM
400
CRC Handbook of Marine Mammal Medicine
presence of Hb and ribosomes. Polychromasia is usually absent or slight in most healthy marine mammals. Therefore, the presence of increased numbers of polychromatophilic RBCs in anemic marine mammals generally indicates an increased percentage of reticulocytes and a regenerative anemia. Increased anisocytosis (variation in RBC diameters) due to different populations of cells and the presence of nucleated RBCs, including rubricytes and metarubricytes, can also be observed with regenerative anemias. However, circulating nucleated RBCs may also be present in nonanemic disorders. A good example of this is the frequent presence of nucleated RBCs with no anemia associated with chronic inflammatory disease in manatees. The presence of reticulocytosis (polychromasia) indicates anemia resulting from blood loss or increased RBC destruction. In general, hemolytic anemias produce more pronounced reticulocytosis than does hemorrhage, with a lag response time of about 4 days. Anemias with no increase in, or low numbers of, reticulocytes (no or minimal polychromasia) are classified as poorly regenerative, or nonregenerative, anemias. Reticulocyte counts are given as a percentage of total RBCs. The reticulocyte count would therefore be higher in an anemic animal than in a normal animal, if the absolute number of reticulocytes in the circulation were the same. Thus, it is often recommended that the reticulocyte count be corrected by using this formula: Corrected Reticulocyte Count (%) = [ actual HCT/Mean normal HCT* ] × uncorrected reticulocyte count (%)
(1)
Anemia can also be classified and correlated to potential pathological etiologies by RBC indices, MCV, and mean corpuscular hemoglobin concentration (MCHC). Anemias related to RBC size are macrocytic (increased MCV), normocytic (normal MCV), and microcytic (decreased MCV). Anemias related to RBC Hb concentrations are normochromic (normal MCHC) or hypochromic (decreased MCHC). Hyperchromic (high MCHC) anemias are artifacts. Classification of Anemia by RBC Indices Normocytic, Normochromic 1. 2. 3. 4. 5. 6. 7. 8.
Acute hemolysis before sufficient time has elapsed for reticulocyte response Mild hemorrhage that does not stimulate reticulocyte response Chronic inflammation, common particularly in dolphins and manatees Chronic renal disease, common in California sea lions Endocrine deficiencies Aplastic and hypoplastic bone marrow Lead intoxication Vitamin B12 deficiency
Macrocytic, Hypochromic 1. Regenerative anemia with marked reticulocytosis, consistent with regenerative response (a favorable prognostic indicator); pattern observed in recovering animals that lost a large volume of blood following traumatic injury Macrocytic, Normochromic 1. Regenerative anemia (e.g., recovery from hemorrhage) 2. Folate deficiency (rare; however, in marine mammals, some drugs, particularly sulfamethoxazole, have antifolate-like activity, which can produce life-threatening anemias; McBain, 1984; Townsend, 1999) *For a particular marine mammal species.
0839_frame_C19 Page 401 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
401
Microcytic, Normochromic, or Hypochromic 1. Chronic iron deficiency (time of induction is months in adults and weeks in nursing animals) 2. Anemia of chronic disease (usually normocytic)
Nonregenerative anemias without an accompanying leukopenia or thrombocytopenia suggest a bone marrow abnormality affecting erythroid cells. A bone marrow aspirate and/or biopsy is recommended in these cases to assess bone marrow function. The prognosis in these cases is guarded. In most marine mammal species, regenerative anemias can be seen with acute blood loss (e.g., traumatic injury, hemorrhagic gastritis, gastroenteritis) or secondary to various chronic disease states (Townsend and Petro, 1998). The cause of anemia secondary to chronic disease is multifactorial and poorly understood in marine mammals. Abnormalities that can contribute to this type of anemia include low serum iron, the production of inflammatory mediators that inhibit erythropoiesis, and shortened RBC life spans, presumably secondary to inflammatory oxidants. Nonregenerative anemia in marine mammals is uncommon. Examples include anemia secondary to chronic renal disease in California sea lions with leptospirosis (presumably due to lack of erythropoietin), sulfamethoxazole-induced fatal pancytopenia in cetaceans, and bone marrow neoplasia (e.g., metastatic squamous cell carcinoma in a bottlenose dolphin, and immunoblastic malignant lymphoma and other lymphoproliferative disorders in various dolphin species) (McBain, 1984; Bossart et al., 1997; Townsend, 1999; Manire and Rhinehart, 2000).
Evaluation of Leukocytes Marine mammal leukocytes, or white blood cells (WBCs), are classified like those of other mammals as either polymorphonuclear (PMN) or mononuclear leukocytes. The PMN leukocytes are also referred to as granulocytes, because of their cytoplasmic granules. These granules represent lysosomes, which contain hydrolytic enzymes and other antibacterial compounds. Most marine mammal granulocytes include neutrophils, eosinophils, and basophils. Granulocytes constitute the cellular components of the innate or nonspecific immune system (see Chapter 12, Immunology). A detailed discussion of lymphocyte subsets and new diagnostic immunological tests for marine mammals can be also found in Chapter 12.
Neutrophils or Heterophils Traditional cytochemical techniques have demonstrated that leukocyte morphology and relative cell numbers in healthy cetaceans and sirenians are similar to terrestrial mammals, with some notable exceptions (Bossart, 1995; Reidarson et al., 2000). The neutrophils of most marine mammals have round to oval granules that either do not stain or stain a pale pink, with routine Romanowsky-type blood stains. However, the sirenians have neutrophils with distinct-staining pleomorphic, round to oval, red to pink granules (Bossart and Bigger, 1994). These cells have been termed heterophils or heterophilic granulocytes, and they stain positive for myeloperoxidase (similar to neutrophils in humans, dogs, cats, and horses, but heterophils of rabbits and birds) (Kiehl and Schiller, 1994). Converse et al. (1994) demonstrated that this segmented cell with eosinophilic granules more closely resembles the typical mammalian neutrophil.
Eosinophils In manatees, eosinophils are the least numerous circulating granulocytes. This differs from the bottlenose dolphin, in which eosinophils are the second most common granulocyte (Bossart, 1995). It appears that the manatee may not have a clearly distinct eosinophil population.
0839_frame_C19 Page 402 Tuesday, May 22, 2001 11:15 AM
402
CRC Handbook of Marine Mammal Medicine
In some cases, with routine staining, only subtle differences are present between the manatee’s neutrophilic granulocyte and eosinophilic granulocyte (Bossart, 1995); the significance of this is unknown and deserves further study. Bottlenose dolphins typically have higher eosinophil numbers than most other mammals (see Table 1) (Asper et al., 1990; Reidarson et al., 2000). The functions of eosinophils are not completely understood in mammals. Eosinophils accumulate in tissues in response to chemoattractants generated in response to parasites. Eosinophil peroxidase released from granules interacts with hydrogen peroxide generated from the respiratory burst and halide ions (Meyer and Harvey, 1998). This complex, combined with other oxygen metabolites and the major basic protein released from their secondary granules, gives eosinophils bactericidal and parasiticidal effects (Clark and Kaplan, 1975; Roth and Kaeberle, 1981; Jain, 1986). The high number of circulating, pulmonary, and gastrointestinal eosinophils in dolphins may represent an enhanced role of the eosinophil in bactericidal and parasiticidal effects.
Basophils In marine mammals, basophil granulocytes are rare. In one study with bottlenose dolphins (n = 180) and manatees (n = 56), basophils were never observed (Bossart, 1995).
Monocytes and Lymphocytes Marine mammal mononuclear leukocytes consist of monocytes and lymphocytes. Monocytes are usually larger than lymphocytes with a nuclear-to-cytoplasmic ratio of ≤1:1. Monocyte nuclei are pleomorphic with round and kidney-shaped nuclei typical forms. In blood smears prepared using EDTA, artifacts that look like cytoplasmic granules may be observed. Lymphocytes have high nuclear-to-cytoplasmic ratios and generally have monomorphic, round nuclei with coarsely clumped chromatin. Most lymphocytes of marine mammals reside within lymphoid organs (lymph nodes, thymus, spleen, and/or bone marrow), and low numbers circulate in the blood. One exception to this is the manatee. Manatees have high circulating total lymphocyte numbers compared with cetaceans or pinnipeds (Bossart, 1995). This pattern is similar to the manatee’s evolutionary relative, the elephant seal (Silva and Kuruwita, 1993). T- and B-lymphocytes have been described in some cetaceans and sirenians (see Chapter 12, Immunology). The total number and differential count of WBCs vary considerably among marine mammal species. Monitoring the dynamic shifts of the relative and absolute WBC numbers in these species often provides more useful diagnostic and prognostic information than the total WBC count alone.
Leukocytes and Age Older cetaceans tend to have higher numbers of total leukocytes (Englehart, 1979). The neutrophilto-lymphocyte ratio tends to be higher at birth than later in life in some cetaceans and manatees. In domestic animals, this is thought to be due to increased endogenous blood cortisol levels at birth that cause circulating neutrophils to increase and lymphocytes to decrease. In bottlenose dolphins, exogenous glucocorticoid administration parallels the changes seen in terrestrial mammals, including neutrophilia, lymphopenia, eosinopenia, hyperglycemia, and elevated levels of insulin, triglycerides, and liver-specific enzymes (Reidarson and McBain, 1999). Many of these changes were also observed in wild belugas (Delphinapterus leucas) (St. Aubin and Geraci, 1989) (see Chapter 10, Endocrinology; Chapter 13, Stress).
0839_frame_C19 Page 403 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
403
Leukocytes and Disease In marine mammals, peripheral blood leukocyte dynamics in various disease states follow a typical mammalian pattern, with some distinct species-specific exceptions. In one study, acutely ill bottlenose dolphins with bacterial infections had leukocytoses that were two to three times the WBC reference range (Bossart, 1995). This leukocytosis involved an absolute mature neutrophilia and eosinophilia, without the presence of band neutrophils. The dolphins in this study demonstrated an apparently adequate innate immunological response to bacterial infection. When appropriate antibiotic therapy was initiated, many of these dolphins survived the illness with eventual complete recovery. Alternatively, bottlenose dolphins that had terminal overwhelming, Gram-negative bacterial endotoxemias had profound leukopenia, with absolute neutropenia, lymphopenia, and eosinopenia, without the presence of band neutrophils. An absolute neutrophilia with the presence of many band neutrophils (a regenerative left shift) may occur in diseased bottlenose dolphins and killer whales (Orcinus orca). This indicates a bone marrow response to the demand for neutrophils (usually a good prognostic indicator). Regenerative left shifts are common, and have been reported in adenoviral hepatitis in California sea lions (Dierauf et al., 1981). Care must be taken in the identification of band neutrophils, as variations in interpretations often exist among laboratory technical staff. Variations of as much as 7% have been seen in values reported from different laboratories, and within the same laboratory, using different technicians. Band cells are not typically seen in healthy killer whales, manatees, and sea otters (Cornell et al., 1981; Williams and Pulley, 1983). Additionally, care must be taken in interpreting absolute neutrophilias in marine mammals. Although pathological neutrophilia occurs with inflammation and infectious disease, physiological neutrophilia can occur with epinephrine and corticosteroid release during exercise, excitement, or stressful situations (see Chapter 13, Stress). Inherited neutrophil effects occur in some marine mammals, although they are rare. Chediak– Higashi syndrome has been reported in a killer whale (Taylor and Farrell, 1973). The syndrome is characterized by the presence of enlarged membrane-bound granules in many cell types, including neutrophils. Neutrophils from affected animals exhibit reduced mobility and defective phagocytic and/or bactericidal responses. Monocytosis may occur in marine mammals in both acute and chronic inflammatory conditions. A pronounced absolute monocytosis has been observed in a bottlenose dolphin recovering from morbillivirus bronchopneumonia, in two bottlenose dolphins recovering from orthopedic jaw surgery to correct traumatic fractures (Bossart, unpubl. data), and in a bottlenose dolphin recovering from Aspergillus fumigatus pneumonia (Reidarson et al., 1998). Circulating total leukocyte numbers of manatees appear to be less responsive than those of other mammals (Walsh and Bossart, 1999). A severe leukocytosis in a manatee is only ∼25,000 3 cells/mm . More important diagnostic information can be obtained in manatees by observing shifts in the leukocyte differential (see Chapter 43, Manatees).
Serum Analytes and Enzymes Glucose, Lipids, and Pancreatic Enzymes Fasting levels of blood glucose in marine mammals are normally higher than those in domestic animals (≥100 mg/dl) (Ridgway, 1972). This may be a response to animal restraint, handling, and/or venipuncture, leading to release of endogenous glucocorticoids. Medway and Geraci (1986) postulated that elevated blood glucose levels were a response to increased central nervous
0839_frame_C19 Page 404 Tuesday, May 22, 2001 11:15 AM
404
CRC Handbook of Marine Mammal Medicine
system requirements for glucose during diving. Additionally, higher glucose levels may fuel muscles for diving (Vallyathan et al., 1969). A diet high in protein and fat may also contribute to higher glucose levels (Greenwood et al., 1971). Hyperglycemia is also seen with diabetes mellitus, obesity, hyperpituitarism, hyperthyroidism, and endogenous or exogenous glucocorticoids (St. Aubin and Geraci, 1989; Reidarson and McBain, 1999). Blood glucose levels in neonatal marine mammals, particularly harbor seals, bottlenose dolphins, and manatees, must be monitored regularly (see Chapter 30, Intensive Care). Polar bears experience a rise in blood glucose levels from October to November, yet no diurnal fluctuations occur. This seasonal fluctuation may be due to decreasing ambient temperatures in the fall or greater activity as ice floes begin to form (Lee et al., 1977). Glucose stress, excitement, anoxia diabetes mellitus obesity hyperpituitarism/hyperthyroidism endogenous glucocorticoids (stress, adrenal disease) severe systemic disease neoplasia hyperinsulinism malnutrition, starvation fulminating hepatic necrosis hepatic cirrhosis glycogen storage disease
Prolonged contact of serum with metabolically active erythrocytes decreases glucose values. Glucose values can be increased with lipemia, hemolysis, and severe azotemia. Some drugs or hormones can increase serum or plasma glucose levels either artifactually or physiologically (e.g., corticosteroids, phenytoin, estrogen, progesterone, thyroxine, asparaginase, β-adrenergics, furosemide, glucose, heparin, methimazole, phenothiazines, and theophylline). Food intake can also cause a glucose increase. Drugs that can decrease serum or plasma glucose levels either physiologically or artifactually include acetaminophen, aspirin, insulin, ascorbic acid, and propanolol. Commercially available portable blood glucose meters and color reagent test strips provide blood glucose concentrations reasonably close to those obtained with reference methods. Some devices are more accurate than others; thus, the clinician must be aware that in some cases, the use of results from these devices may lead to erroneous clinical decisions (Cohn et al., 2000).
Total Cholesterol and Triglycerides Altered lipid metabolism can accompany hepatic disease. Since cholesterol and triglycerides are typically eliminated through the formation of bile acids, increases in these two compounds can develop with cholestatic disease. In bottlenose dolphins and false killer whales (Pseudorca crassidens), fatal necrotizing pancreatitis can result in increases in cholesterol and/or triglycerides (Bossart, unpubl. data). Postprandial pinnipeds and cetaceans have lipemic sera high in triglycerides (Nelson, 1970; Geraci et al., 1979). Orphaned neonatal harbor seals (Roletto and Dougherty, 1983), California sea lions (Ridgway, 1972), manatees, and bottlenose dolphins (Bossart, unpubl. data) can have low
0839_frame_C19 Page 405 Tuesday, May 22, 2001 11:15 AM
405
Clinical Pathology
triglyceride and/or cholesterol levels, probably as a result of prolonged inanition. Pregnant harbor seals may have higher triglyceride levels than nonpregnant seals (Kuiken, 1985). Triglycerides and Cholesterol postprandial acute pancreatitis diabetes mellitus hypothyroidism nephrotic syndrome cholestatic hepatic disease hyperadrenocorticism prolonged inanition malabsorption hyperthyroidism
Triglyceride values can vary widely with dietary change. Lipemia and icterus have variable effects on both triglyceride and cholesterol levels. The anticoagulants oxalate and fluoride can raise cholesterol levels. Drugs that can raise cholesterol and/or triglycerides include corticosteroids, estrogen, phenytoin, and phenothiazines, whereas dipyrone, thyroxine, ascorbic acid, asparaginase, and heparin can decrease them.
Amylase, Lipase, and Trypsin-Like Immunoreactivity Amylase and lipase are the enzymes measured most frequently as markers of acute pancreatitis in small domesticated animals. However, amylase and lipase have pancreatic and extrapancreatic sources (Williams, 1994). The presence of these enzymes in serum from extrapancreatic organs, particularly the liver and kidneys, makes it difficult to diagnose pancreatitis based solely on observing elevated serum levels of amylase and lipase (Archer et al., 1997). Additionally, in dogs, low amylase and lipase activities cannot be used to rule out pancreatitis because their activities are not necessarily high in all animals with disease (Braun et al., 1997). Six cases of acute necrotizing pancreatitis in marine mammals have been observed (Bossart, unpubl. data). These cases were diagnosed post-mortem by histological examination of the pancreas in Atlantic bottlenose dolphins and false killer whales. Two of the bottlenose dolphin calves had high ante-mortem amylase and lipase values (>1000 IU/l). The remaining animals with acute pancreatitis had ante-mortem serum amylase and lipase values within normal reference ranges. Pancreatitis in older cetaceans has been diagnosed by observing pale, greasy feces with leukocytosis and increased serum amylase levels (Sweeney and Ridgway, 1975); however, this study did not provide actual amylase values. Therefore, if acute pancreatitis is suspected in marine mammals, it is prudent to determine the values of both amylase and lipase with a greater than twofold increase in serum activity considered suggestive of acute pancreatitis. Normal, healthy, neonatal and juvenile manatees have high levels of amylase and lipase compared with other marine mammals (Bossart, unpubl. data). No correlation has been observed between these levels and pancreatic disease, nor are serum amylase levels increased in manatees with prerenal azotemia. Chronic pancreatitis typically is caused by recurrent acute or subacute pancreatitis in domestic animals; neither amylase nor lipase levels would be expected to change because organ fibrosis limits the ability of the organ to produce digestive enzymes. Recently, a radioimmunoassay
0839_frame_C19 Page 406 Tuesday, May 22, 2001 11:15 AM
406
CRC Handbook of Marine Mammal Medicine
technique to measure trypsinogen (measured as trypsin-like immunoreactivity or TLI) has been developed and validated for the dog and cat (Lucena et al., 1999). Trypsinogen is specific for the pancreas (Carro and Williams, 1989), so variations in serum TLI can be used to help diagnose pancreatitis (Lucena et al., 1999). In dogs, cats, and humans, TLI has become the cornerstone for the diagnosis of exocrine pancreatic insufficiency (EPI) because of the simplicity and high sensitivity and specificity of the test; a decrease in TLI is diagnostic of EPI. However, the TLI test is species specific, as there is minimal or no cross-reactivity of the test among species, due to antibody specificity used in the test. Two bottlenose dolphins with suspected EPI were tested with the antibody to canine TLI (Stelling, pers. comm.). The TLI values in these two cases were below test ranges in six healthy bottlenose dolphins. However, the significance of these findings was unclear because of the species-specific nature of this test and the lack of known antibody cross-reactivity between canine and dolphin. Lipase and Amylase pancreatic inflammation, necrosis, and neoplasia pancreatic duct obstruction chronic renal insufficiency slight increase with glucocorticoids (lipase only) intestinal perforations
Increases occur with hemolysis and decreases with lipemia (amylase only) and the anticoagulants EDTA, oxalate, citrate, and fluoride (amylase only). Exogenous corticosteroids can increase lipase and decrease amylase. Increases can also occur with sulfonamides, tetracycline, and estrogen (amylase only).
Markers of Hepatobiliary System Disorders Primary hepatic disease and secondary reactions of the liver to extrahepatic disease can cause identical serum abnormalities. The secondary hepatic response is not uncommon in marine mammals, and frequently causes the clinician a diagnostic dilemma. Serum hepatic enzyme tests in terrestrial mammals are grouped into those that indicate hepatocellular injury or repair— alanine aminotransferase (ALT), aspartate aminotransferase (AST), glutamate dehydrogenase (GLDH), sorbitol dehydrogenase (SDH), and lactate dehydrogenase (LDH)—and those that reflect increased enzyme production stimulated by retained bile or drug induction—alkaline phosphatase (ALP) and γ-glutamyltransferase (GGT). Similar enzyme patterns occur in marine mammals with a few exceptions.
Alanine Aminotransferase (ALT or SGPT) In some terrestrial mammals, ALT is a useful indicator of acute diffuse hepatocellular injury, with the magnitude of serum increase crudely reflecting the number of injured hepatocytes. ALT, produced by the liver, appears to be liver specific in bottlenose dolphins, Pacific white-sided dolphins (Lagenorhynchus obliquidens), and pilot whales (Globicephala spp.) (Bossart, 1984; Bossart et al., 1990; 1991). ALT is likely liver-specific in pinnipeds, but not in manatees. The sirenians, as nonruminant herbivores, appear to be similar to horses and ruminants, all of which have insignificant levels of hepatic ALT. Besides hepatocellular necrosis, rises in ALT are associated with parasitism, neoplasia, and hepatic and muscular trauma. In polar bears, ALT fluctuates seasonally (Cattet, pers. comm.).
0839_frame_C19 Page 407 Tuesday, May 22, 2001 11:15 AM
407
Clinical Pathology
ALT (SGPT) liver disease (species-specific), including: infection (particularly hepatic necrosis) parasitism neoplasia trauma ↓
hepatic cirrhosis (species-specific)
Hemolysis, lipemia, corticosteroids, anabolic steroids, acetaminophen, aspirin, ibuprofen, phenylbutazone, sulfonamides, phenobarbital, phenytoin, primidone, barbiturates, azathioprine, asparaginase, cimetidine, salicylates, and thiacetarsamide can all elevate ALT.
Aspartate Aminotransferase (AST or SGOT) In terrestrial mammals, a variety of tissues, particularly skeletal muscle and liver, have high AST activity. This is also the trend in marine mammals. In some cetaceans with hepatic disease, serum AST activity will return to normal more rapidly than serum ALT activity. This may be due to differences in plasma half-lives and cellular location. Persistent mild to moderate increases of serum ALT or AST activity suggest a “smoldering” inflammatory process, such as chronic–active hepatitis. AST values in porpoises are comparatively higher in those animals with higher metabolic rates (Ridgway, 1972). In seals, AST may not be a useful test for liver damage because handling and restraint may produce muscle damage leading to elevated enzyme levels (Medway and Geraci, 1986; Reidarson et al., 1998). In sirenians, elevations of AST are seen with massive skeletal muscle damage secondary to boat collisions (Bossart, unpubl. data). Primary hepatic disease is rare in manatees, so correlations between high AST and liver disease in this species are difficult. In sea otters, baseline AST levels may be three times higher than values for other mustelids (Williams and Pulley, 1983). With overexertion in sea otters, AST is elevated, apparently as a result of skeletal muscle damage, rather than primary hepatic disease. In polar bears, AST fluctuates seasonally (Cattet, pers. comm.). AST (SGOT) liver disease (species-specific), including: hepatocellular necrosis neoplasia parasites (particularly flukes) skeletal muscle damage heart disease ↓
hepatic cirrhosis (species-specific)
Lipemia, hemolysis, and ketonemia may increase AST. Corticosteroids and salicylates can increase AST levels, due to a physiological hepatic change. Metronidazole can decrease AST.
Sorbitol Dehydrogenase (SDH) and Glutamate Dehydrogenase (GLDH) In most terrestrial mammals, SDH and GLDH are located primarily in the liver and are elevated following acute hepatic insult. The hepatic activity of these enzymes in marine mammals is unknown, but evaluation of these enzymes may be diagnostically valuable. Because convenient
0839_frame_C19 Page 408 Tuesday, May 22, 2001 11:15 AM
408
CRC Handbook of Marine Mammal Medicine
methodologies for running these tests are unavailable, the use of these enzymes is at present limited.
Lactate Dehydrogenase (LDH) LDH is located in numerous body tissues, including liver, kidney, pancreas, intestine, heart, brain, and skeletal muscle. Serum LDH consists of five isoenzymes, separable by electrophoresis, LDH1 through LDH5. The widespread tissue distribution of LDH limits its popular diagnostic use in domestic animal veterinary medicine. In some species, isoenzyme patterns reflect specific tissue damage (Levinson and Hobbs, 1994; Patel et al., 1994; Drent et al., 1996; Lossos et al., 1997). Different clinical conditions in cetaceans, such as intramuscular injections, azole therapy, and trauma, produce distinctly differing LDH isoenzyme patterns, so LDH isoenzyme patterns may provide useful corroboration with other clinical findings when diagnostic modalities are limited (Reidarson et al., 1999a). In stranded cetaceans, total serum LDH (nonsubdivided) concentrations can be elevated. Elevation also occurs with some pulmonary infections (Bossart and Trimm, 1993) and following trauma. In sea otters, LDH levels are higher than in other mustelids, but not as high as those in cetaceans and pinnipeds (Williams and Pulley, 1983). LDH skeletal muscle necrosis or degeneration intramuscluar injections pulmonary disease (species-specific) hepatic, myocardial, central nervous system, or intestinal disease
Increases in serum LDH concentrations occur with severe hemolysis and when there is a delay in separation of serum from cells.
-Glutamyltransferase (GGT) GGT is an enzyme located on the cellular membranes of many organs, including the liver, heart, skeletal muscle, kidneys, and pancreas. In the liver, GGT is associated with epithelial cells of the biliary system. Renal tubular epithelial cells also have high GGT activity. Acute tubular injury results in an increase in the activity of GGT in the urine (but not the serum) in marine mammals (Medway and Geraci, 1986). Therefore, the measurement of urine GGT may be a useful indicator of acute tubular necrosis, such as that associated with the use of aminoglycoside antibiotics. In most terrestrial mammals, normal hepatic GGT activity is minimal, but can become markedly elevated in serum subsequent to increased production stimulated by impaired bile flow. This also appears to occur with cholestasis and biliary tract disease in some species of dolphins (Bossart, 1984; Bossart et al., 1990). Hepatic cirrhosis in bottlenose dolphins can result in sustained serum GGT levels (>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.
0839_frame_C19 Page 409 Tuesday, May 22, 2001 11:15 AM
409
Clinical Pathology
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
0839_frame_C19 Page 410 Tuesday, May 22, 2001 11:15 AM
410
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C19 Page 411 Tuesday, May 22, 2001 11:15 AM
411
Clinical Pathology
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
0839_frame_C19 Page 412 Tuesday, May 22, 2001 11:15 AM
412
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 413 Tuesday, May 22, 2001 11:15 AM
413
Clinical Pathology
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.
0839_frame_C19 Page 414 Tuesday, May 22, 2001 11:15 AM
414
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 415 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
415
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.
0839_frame_C19 Page 416 Tuesday, May 22, 2001 11:15 AM
416
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 417 Tuesday, May 22, 2001 11:15 AM
417
Clinical Pathology
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).
0839_frame_C19 Page 418 Tuesday, May 22, 2001 11:15 AM
418
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 419 Tuesday, May 22, 2001 11:15 AM
419
Clinical Pathology
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.
0839_frame_C19 Page 420 Tuesday, May 22, 2001 11:15 AM
420
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C19 Page 421 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
421
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.
0839_frame_C19 Page 422 Tuesday, May 22, 2001 11:15 AM
422
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 423 Tuesday, May 22, 2001 11:15 AM
423
Clinical Pathology
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.
0839_frame_C19 Page 424 Tuesday, May 22, 2001 11:15 AM
424
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 425 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
425
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.
0839_frame_C19 Page 426 Tuesday, May 22, 2001 11:15 AM
426
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C19 Page 427 Tuesday, May 22, 2001 11:15 AM
427
Clinical Pathology
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
0839_frame_C19 Page 428 Tuesday, May 22, 2001 11:15 AM
428
CRC Handbook of Marine Mammal Medicine 3
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,
0839_frame_C19 Page 429 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
429
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).
0839_frame_C19 Page 430 Tuesday, May 22, 2001 11:15 AM
430
CRC Handbook of Marine Mammal Medicine
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.
References Archer, F.J., Kerr, M.E., and Houston, D.M., 1997, Evaluation of three pancreatic specific protein assays, TLI, PASP and CA 19-9 for diagnosis of canine pancreatitis, Zentralbl. Veterinärmed. [A], 44: 109–114. Asper, E.D., Cornell, L., Duffield, D.A., Odell, D.K., Joseph, B.E., Stark, B.I., and Perry, C.A., 1990, Hematology and serum chemistry values in bottlenose dolphins, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 479–485. Banish, L.D., and Gilmartin, W.G., 1988, Hematology and serum chemistry of the young Hawaiian monk seal (Monachus schauinslandi), J. Wildl. Dis., 24: 225. Bateman, S.W., Matthews, K.A., Abrams-Ogg, A.C.G., Lumsden, J.H., Johnstone, I.B., Hillers, T.K., and Foster, R.A., 1999, Diagnosis of disseminated intravascular coagulation in dogs admitted to an intensive care unit, J. Am. Vet. Med. Assoc., 215: 798–804. Beckman, K.B., Lowenstine, L.J., Newman, J., Hill, J., Hanni, K., and Gerber, J., 1997, Clinical and pathological characterization of northern elephant seal skin disease, J. Wildl. Dis., 33: 438–449. Bonner, W.N., and Fogden, S.C.L., 1974, A note on blood typing seals, Rapp. P. V. Reun. Cons. Int. Explor. Mer, 161: 439. Borer, W.Z., 1992, Selection and use of laboratory tests, in Applied Laboratory Medicine, Tietz, N.W. (Ed.), Saunders, Philadelphia, 1–5. Bossart, G.D., 1984, Suspected acquired immunodeficiency in an Atlantic bottlenose dolphin with chronic-acute hepatitis and lobomycosis, J. Am. Vet. Med. Assoc., 185: 1413–1414. Bossart, G.D., 1995, Immunocytes of the Atlantic Bottlenose Dolphin (Tursiops truncatus) and West Indian Manatee (Trichechus manatus latirostris): Morphologic Characterizations and Correlations between Healthy and Disease States and Under Free-Ranging and Captive Conditions, Dissertation, Florida International University, Miami, FL, 183 pp. Bossart, G.D., and Bigger, C.H., 1994, Cellular components of the nonspecific, humoral and cell-mediated immune system in the peripheral blood of the West Indian manatee (Trichechus manatus latirostris): Cellular populations in health and disease, in Proceedings of the First International Manatee and Dugong Research Conference, 148 pp. Bossart, G.D., and Dierauf, L.A., 1990, Marine mammal clinical laboratory medicine, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 1–52. Bossart, G.D., and Eimstad, E., 1988, Erysipelothrix vesicular glossitis in a killer whale (Orcinus orca), J. Zoo Anim. Med., 19: 42–47. Bossart, G.D., and Schwartz, J.C., 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., and Trimm, M.A., 1993, Neuromuscular electrical stimulation therapy for stranded short-finned pilot whales (Globicephala macrorhynchus) with rhabdomyolysis and skeletal muscle atrophy resulting in scoliosis, in Proceedings of the 10th Biennial Conference on the Biology of Marine Mammals, Chicago, 63 pp. Bossart, G.D., Brawner, T.A., Cabal, C., Kuhns, M., Eimstadt, E.A., Caron, J., Trimm, M., and Bradley, P., 1990, Hepatitis B-like infection in a Pacific white-sided dolphin (Lagenorhynchus obliquidens), J. Am. Vet. Med. Assoc., 196: 127–130.
0839_frame_C19 Page 431 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
431
Bossart, G.D., Odell, D.K., Walsh, M.T., Lynch, J.D., Beusse, D.O., Friday, R., and Young, G.G., 1991, The histopathologic findings in a mass stranding of pilot whales (Globicephala macrorhynchus), in Marine Mammal Strandings in the United States, NOAA/NMFS, 98: 85–89. Bossart, G.D., Cray, C., Solarzano, J.L., Decker, S.F., Cornell, L.H., and Altman, N.H., 1996, Cutaneous papovaviral-like papillomatosis in a killer whale (Orcinus orca), Mar. Mammal Sci., 12: 274–281. Bossart, G.D., Ewing, R., Herron, A.S., Cray, C., Decker, S., Alexander, J., and Altman, N.H., 1997, Immunoblastic malignant lymphoma in dolphins: Histologic, ultrastructural and immunohistochemical features, J. Vet. Diagn. Invest., 9: 454–458. Braun, J.P., Garnier, F., and Medaille, C.H., 1997, Exploration biochimique du pancreas exocrine chez le chien: Une revue, Prat. Med. Chir. Anim. Comp., 32: 15–17. Bryden, M.M., and Linn, G.H.K., 1969, Blood parameters of the southern elephant seal (Mirounga leonina) in relation to diving, Comp. Biochem. Physiol., 28: 139. Cargill, C.F., Needham, D.J., and Judson, G.J., 1979, Plasma biochemical values of clinically normal Australian sea lions (Neophoca cinerea), J. Wildl. Dis., 14: 105–110. Carro, T., and Williams, D.A., 1989, Relationship between dietary protein concentration and serum trypsin-like immunoreactivity in dogs, Am. J. Vet. Res., 50: 2105–2107. Castellini, M.A., Davis, R.W., Loughlin, T.R., and Williams, T.M., 1993, Blood chemistries and body condition of Steller sea lion pups at Marmot Island, Alaska, Mar. Mammal Sci., 9: 202–208. Clark, R.A., and Kaplan, A.P., 1975, Eosinophil leukocytes: Structure and function, Clin. Hematol., 4: 635–642. Clausen, G., and Ersland, A., 1969, The respiratory properties of the blood of the bladdernose seal (Cystophora cristata), Respir. Physiol., 7: 1–6. Cohn, L.A., McCaw, D.L., Tate, D.J., and Johnson, J.C., 2000, Assessment of five portable blood glucose meters, a point-of-care analyzer, and color test strips for measuring blood glucose concentration in dogs, J. Am. Vet. Med. Assoc., 216: 198–202. Coles, E.H., 1986, Veterinary Clinical Pathology, W.B. Saunders, Philadelphia, chaps. 1 (1–8) and 10 (203–213). Converse, L.S., Fernandes, P.S., MacWilliams, P.S., and Bossart, G.D., 1994, Hematology, serum chemistry and morphometric reference values for Antillean manatees (Trichechus manatus manatus), J. Zoo Wildl. Med., 25: 423–431. Cornell, L.H., 1983, Hematology and clinical chemistry values in the killer whale, Orcinus orca, J. Wildl. Dis., 19: 259–264. Cornell, L.H., Duffield, D.A., and Antrim, J.E., 1981, Clinical application of hematological values in captive cetaceans, in Annual Proceedings of the American Association of Zoo Veterinarians, 80. Cornell, L.H., Duffield, D.A., Joseph, B.E., and Stark, B., 1988, Hematology and serum chemistry of the beluga (Delphinapterus leucas), J. Wildl. Dis., 24: 220–224. Costa, D.P., and Ortiz, C.L., 1982, Blood chemistry homeostasis during prolonged fasting in the northern elephant seal, Am. J. Physiol., 242: R591–595. Cotter, S.M., and Blue, J., 1985, The bone marrow, in Textbook of Small Animal Surgery, Slatter, D.H. (Ed.), W.B. Saunders, Philadelphia, 1201–1210. de Swart, R., Ross, P.S., Vedder, L.J., Boink, F.B.T.J., Reijnders, P.J.H., Mulduer, P.G.H., and Osterhaus, A.D.M.E., 1995, Haematology and clinical chemistry values for harbour seals (Phoca vitulina) fed environmentally contaminated herring remain within normal ranges, Can. J. Zool., 73: 2035–2043. Dierauf, L.A., Lowenstine, L.J., and Jerome, C., 1981, Viral hepatitis (adenovirus) in a California sea lion, J. Am. Vet. Med. Assoc., 179: 1194–1197. Dierauf, L.A., Dougherty, S.A., and Baker, B. 1984, Neonatal hyperbilirubinemia in harbor seals (Phoca vitulina richardsi), J. Zoo Anim. Med., 15: 55–59. Dover, S., McBaen, J., and Little, K., 1993, Serum alkaline phosphatase as an indicator of nutritional status in cetaceans, in Annual Proceedings of the International Association for Aquatic Animal Medicine, CD-ROM Archive.
0839_frame_C19 Page 432 Tuesday, May 22, 2001 11:15 AM
432
CRC Handbook of Marine Mammal Medicine
Drent, M., Cobben, N.A., Henderson, R.F., Wouters, E.F., and Dieijen-Visser, M., 1996, The usefulness of lactate dehydrogenase and its isoenzymes as indicators of lung damage or inflammation, Eur. Respir. J., 9: 1736–1742. Ehrhart, N., Dernell, W.S., Hoffman, W.E., Weigel, R.M., Powers, B.E., and Winthrow, S.J., 1998, Prognostic importance of alkaline phosphatase activity in serum from dogs with appendicular osteosarcoma: 75 cases (1990–1996), J. Am. Vet. Med. Assoc., 213: 1002–1005. Elin, R.J., 1994, Magnesium: The fifth but forgotten electrolyte, Clin. Chem., 102: 616–618. Englehardt, F.R., 1979, Haematology and plasma chemistry of captive pinnipeds and cetaceans, Aquat. Mammals, 7: 11–20. Fenwick, B.W., Schroeder, J.P., and Smith, J.E., 1988, Decreases in serum iron concentrations as an indicator of acute phase responses in dolphins: Clinical considerations, in Proceedings of the International Association for Aquatic Animal Medicine, 19: 71. Fothergill, M.B., Schwegman, C.A., Garratt, P.A., Govender, A., and Robertson, W.D., 1991, Serum alkaline phosphatase—Changes in relation to state of health and age of dolphins, Aquat. Mammals,17: 71–75. Geraci, J.R., 1971, Functional hematology of the harp seal (Pagophilus groenlandicus), Physiol. Zool., 44: 162–170. Geraci, J.R., 1972, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618–621. Geraci, J.R., and Engelhardt, F.R., 1974, The effects of storage time, temperature, and anticoagulants on harp seal, Phoca groenlandica, hemograms: A simulated field study, Physiol. Zool., 47: 22–28. Geraci, J.R., and Medway, W., 1973, Simulated field blood studies in the bottlenose dolphin, Tursiops truncatus, effects of stress on some hematologic plasma chemical parameters, J. Wildl. Dis., 9: 29–33. Geraci, J.R., and Smith, T.G., 1975, Functional hematology of ringed seals in the Canadian Arctic, J. Fish. Res. Board Can., 32: 2559–2564. Geraci, J.R., St. Aubin, D.J., and Smith, T.G., 1979, Influence of age, condition, sampling time, and method in plasma chemical constituents in free-ranging ringed seals, Phoca hispida, J. Fish. Res. Board Can., 36: 1278–1282. Goldstein, T., Johnson, S.P., Werner, L.J., Nolan, S. and Hilliard, B.A., 1998, Causes of erroneous white blood cell counts in clinically healthy northern elephant seals (Mirounga angustirostris), J. Zoo Wildl. Med., 29: 408–412. Greenwood, A.G., Ridgway, S.H. and Harrison, R.J., 1971, Blood values in young grey seals, J. Am. Vet. Med. Assoc., 159: 571–578. Gulland, F.M.D., Werner, L., O'Neill, S., Lowenstine, L., Trupkiewitz, J., Smith, D., Royal, B., and Strubel, L., 1996, Baseline coagulation assay values for northern elephant seals (Mirounga angustirostris) and disseminated intravascular coagulation in the species, J. Wildl. Dis., 32: 536–540. Gulland, F.M.D., Beckmen, K., Burek, K., Lowenstine, L., Werner, L., Spraker, T., Dailey, M., and Harris, E., 1997, Nematode (Otostrongylus circumlitus) infestation of northern elephant seals (Mirounga angustirostris) stranded along the California coast, Mar. Mammal Sci., 13: 446–459. Hall, A.J., 1998, Blood chemistry and hematology of gray seal (Halichoerus grypus) pups and from birth to postweaning, J. Zoo. Wildl. Med., 29: 401–407. Heidel, J.R., Philo, L.M., Albert, T.F., Andreasen, C.B., and Stang, B.V., 1996, Serum chemistry of bowhead whales (Balaena mysticetus), J. Wildl. Dis., 32: 75–79. Hubbard, R.C., 1968, Husbandry and laboratory care of pinnipeds, in The Behavior and Physiology of Pinnipeds, Harrison, R.J., Hubbard, R.C., Peterson, R.S., Rice, C.E., and Schusterman, R.J. (Eds.), Appleton-Century-Crofts, New York, 299–358. Hunter, L., and Madin, S.H., 1978, Clinical blood values of northern fur seal, Callorhinus ursinus, II, Comparison of fresh versus frozen serum, J. Wildl. Dis., 14: 116–119. Jain, N.C., 1986, Schlam’s Veterinary Hematology, Lea & Febiger, Philadelphia, 677–729. Kaneko, J.J., 1980, Serum proteins and the dysproteinemias, in Clinical Biochemistry of Domestic Animals, Kaneko, J.J. (Ed.), Academic Press, New York, 97–118.
0839_frame_C19 Page 433 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
433
Kiehl, A.R., and Schiller, C.A., 1994, A study of manatee leukocytes using peroxidase stain, Vet. Clin. Pathol., 23: 50–53. Kooyman, G.L., and Drabek, C.M., 1968, Observations on milk, blood, and urine constituents of the Weddell seal, Physiol. Zool., 41: 187–194. Kuiken, T., 1985, Influences of diet, gestation and age on haematology and plasma chemistry of the harbour seal, Phoca vitulina, Aquat. Mammals, 11: 40. Lander, M.E., Gulland, F.M.D., and DeLong, R.L., 2000, Satellite tracking a rehabilitated Guadalupe fur seal (Arctocephalus townsendi), Aquat. Mammals, 26: 137–142. Lane, R.A.B., Morris, R.J.H., and Sheedy, J.W., 1972, A hematological study of the southern elephant seal (Mirounga leonina), Comp. Bichem. Physiol., 42A: 841–850. Lee, J., Ronald, K., and Oritsland, N.A., 1977, Some blood values of wild polar bears, J. Wildl. Manage., 41: 520–526. Lenfant, C., 1969, Physiological properties of blood of marine mammals, in The Biology of Marine Mammals, Anderson, H.T. (Ed.), Academic Press, New York, 95–116. Lenfant, C., Johansen, K., and Tottance, J.D., 1970, Gas transport and oxygen storage capacity in some pinnipeds and the sea otter, Respir. Physiol., 9: 277–286. Levinson, S.S., and Hobbs, G.A., 1994, Usefulness of various lactate dehydrogenase isoenzyme-1 profiles after myocardial infarction, Am. Clin. Lab. Sci., 24: 364–370. Lewis, J.H., Bayer, W.L., and Szeto, I.L.F., 1969, Coagulation factor 12 deficiency in the porpoise (Tursiops truncatus), Comp. Biochem. Physiol., 31: 667–671. Lossos, I.S., Breuer, R., Intrator, O., and Sonenblick, M., 1997, Differential diagnosis of pleural effusion by lactate dehydrogenase isoenzyme analysis, Chest, 111: 648–651. Lucena, R., Ginel, P.J., Novales, M., and Molleda, J.M., 1999, Effects of dexamethasone administration on serum trypsin-like immunoreactivity in healthy dogs, Am. J. Vet. Res., 60: 1357–1359. MacDonald, M.K., 1981, Increased enzyme activity in rapidly growing northern elephant seal pups, in Proceedings of the 4th Biennial Conference on the Biology of Marine Mammals, 43. Manire, C., and Rhinehart, H.L., 2000, Use of human recombinant erythropoietin for the treatment of nonregenerative anemia in a rough-toothed dolphin (Steno bredanensis), J. Zoo Wildl. Med., 31: 157–163. Manire, C.A., Rhinehart, H.L., Colbert, D.E., and Smith, D.R., 1999, Experimentally induced serum and urinary creatinine elevations in captive West Indian manatees, in Proceedings of the 13th Biennial Conference on the Biology of Marine Mammals, 114. McBain, J., 1984, Sulfamethoxazole toxicity in three killer whales, in Proceedings of the International Association for Aquatic Animal Medicine, 15: 38. McBain, J., 1996, Clinical pathology interpretation in delphinidae with emphasis on inflammation, in Proceedings of the American Association of Zoo Veterinarians, Puerto Vallarta, Mexico, 308. McBain, J., and Reidarson, T.H., 1994, A case of renal failure in a Pacific pilot whale (Globicephala macrorhynchus) and attempted therapy, in Proceedings of the International Association for Aquatic Animal Medicine, 25: 23. McBain, J., and Reidarson, T.H., 1998, Long term rehabilitation of a gray whale (Eschrichtius robustus) calf, in Proceedings of the International Association for Aquatic Animal Medicine, 29: 61. McConnell, L.C., and Vaughan, R.W., 1983, Some values in captive and free-ranging harbor seals (Phoca vitulina), Aquat. Mammals, 10: 9–13. Medway, W., 1976, Some studies on the blood of the Florida manatee, Comp. Biochem. Physiol., 55A: 413–415. Medway, W., and Geraci, J.R., 1964, Hematology of the bottlenose dolphin (Tursiops truncatus), Am. J. Physiol., 207: 1367–1370. Medway, W., and Geraci, J.R., 1965, Blood chemistry of the bottlenose dolphin (Tursiops truncatus), Am. J. Physiol., 209: 169–172. Medway, W., and Geraci, J.R., 1986, Clinical pathology of marine mammals, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 791 pp. Medway, W., Schryver, H.F., and Bell, B., 1966, Clinical jaundice in porpoises, J. Am. Vet. Med. Assoc., 149: 891–895.
0839_frame_C19 Page 434 Tuesday, April 9, 2002 1:38 PM
434
CRC Handbook of Marine Mammal Medicine
Meyer, D.J., and Harvey, J.W., 1998, Veterinary Laboratory Medicine: Interpretation and Diagnosis, W.B. Saunders, Philadelphia, chaps. 3 (43–82) and 7 (157–186). Miller, W.G., Adams, L.G., Ficht, T.A., Cheville, N.F., Payeur, J.P., Harley, D.R., House, C., and Ridgway, S.H., 1999, Brucella-induced abortions and infection in bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med., 30: 100–110. Morgan, L., Kumaresan, S., Thomas, C., and MacWilliams, P., 1998, Hematology and chemistry reference values for free-ranging harbor seals (Phoca vitulina) and the effects of hemolysis on chemistry values of captive seals, J. Zoo Wildl. Med., 29: 394–400. Morgan, L.W., Van Bonn, W., Jensen, E.D., and Ridgway, S.H., 1999, Effects of in vitro hemolysis on serum biochemistry values of the bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 30: 70–75. Needham, D.J., Cargill, C.F., and Sheriff, D., 1980, Haematology of the Australian sea lion, Neophoca cinerea, J. Wildl. Dis., 16: 103–109. Nelson, G.I., 1970, The lipid composition of the blood of marine mammals, I. Young elephant seals, Mirounga leonina and harp seals, Pagophilus groenlandicus, Comp. Biochem. Physiol., 34: 109–115. Patel, P.S., Adharyu, S.G., and Balar, D.B., 1994, Serum lactate dehydrogenase and its isoenzymes in leukemia patients: Possible role in the diagnosis and treatment monitoring, Neoplasm, 41: 55–59. Poelstra, K., Bakker, W.W., Klok, P.A., Hardonk, M.J., and Meijer, D.K.F., 1997, A physiologic function for alkaline phosphatase: Endotoxin detoxification, Lab. Invest., 76: 319–327. Rea, L.D., Castellini, M.A., Fadely, B.S., and Loughlin, T.R., 1998, Health status of young Alaska sea lion pups (Eumetopias jubatus) as indicated by blood chemistry and hematology, Comp. Biochem. Physiol. A Mol. Integrative Physiol., 120A: 617–623. Reidarson, T.H., and McBain, J., 1999, Hematology, biochemical and endocrine effects of dexamethasone on bottlenose dolphins (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 65–66. Reidarson, T.H., Harrell, J.H., Rinaldi, M.G., and McBain, J., 1998, Bronchoscopic and serologic diagnosis of Aspergillus fumigatus pulmonary infection in a bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 451–455. Reidarson, T.H., McBain, J., and Dalton, L.M., 1999a, Lactate dehydrogenase isoenzyme patterns in cetaceans, J. Zoo Wildl. Med., 30: 228–234. Reidarson, T.H., McBain, J., Dalton, L.M., and Rinaldi, M.G., 1999b, Diagnosis and treatment of fungal infections in marine mammals, in Zoo and Wild Animal Medicine 4, Fowler, M.E., and Miller, E.R. (Eds.), W.B. Saunders, Orlando, FL, 478–485. Reidarson, T.H., Duffield, D., and McBain, J., 2000, Normal hematology of marine mammals, in Schalm’s Veterinary Hematology, 5th ed., Feldman, B.F., Zinkl, J.G., and Jain, N.C. (Eds.), Lippincott Williams & Wilkins, Philadelphia, 1164–1173. Reidarson, T.H., McBain, J., and Yochem, P., in press, Medical and nutritional aspects of a California gray whale (Eschrichtius robustus) rehabilitation, Aquat. Mammals. Ridgway, S.H., 1972, Homeostasis in the aquatic environment, in Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 590–747. Ridgway, S.H., Bowers, C.A., Miller, D., Schultz, M.I., Jacobs, C.A., and Dooley, C.A., 1984, Diving and blood oxygen in the white whale, Can. J. Zool., 62: 2349–2351. Robinson, A.J., Dropatkin, M., and Aggeler, P.M., 1969, Hageman factor (12) deficiency in marine mammals, Science, 166: 1420. Roletto, J., 1993, Hematology and serum chemistry values for clinically healthy and sick pinnipeds, J. Zoo Wildl. Med., 24: 145–157. Roletto, J., and Dougherty, S.A., 1983, Hematologic changes in young growing phocids (Phoca vitulina and Mirounga angustirostris), Abstr., Proceedings of the International Association for Aquatic Animal Medicine, 14: 29. Ronald, K., and Kay, J., 1983, Haematology and plasma chemistry of captive Baikal seals, Aquat. Mammals, 9: 83–87.
0839_frame_C19 Page 435 Tuesday, May 22, 2001 11:15 AM
Clinical Pathology
435
Ronald, K., Foster, M.E., and Johnson, E., 1969, The harp seal, Pagophilus groenlandicus, II. Physical blood properties, Can. J. Zool., 47: 461–468. Rosonke, B.J., Brown, S.R., Tornquist, S.J., Snyder, S.P., Garner, M.M., and Blythe, L.L., 1999, Encephalomyelitis associated with a Sarcosystis neurona-like organism in a sea otter, J. Am. Vet. Med. Assoc., 215: 1839–1842. Roth, J.A., and Kaeberle, M.L., 1981, Isolation of neutrophils and eosinophils from the peripheral blood of cattle and comparison of functional activities, J. Immunol. Methods, 45: 153–156. St. Aubin, D.J., and Geraci, J.R., 1989, Adaptive changes in hematologic and plasma chemical constituents in captive beluga whales (Delphinapterus leucas), Can. J. Fish. Aquat. Sci., 46: 796–803. Saito, H., Poon, M., Goldsmith, G.H., Ratnoff, O.D., and Arnason, U., 1976, Studies in the blood clotting and fibrinolytic system in the plasma from a sei whale, Proc. Soc. Exp. Biol. Med., 152: 503. Schumacher, U., Rauh, G., Plotz, J., and Welsch, U., 1992, Basic biochemical data on blood from Antarctic Weddell seals (Leptonychotes weddelli): Ions, lipids, enzymes, serum proteins and thyroid hormones, Comp. Biochem. Physiol., 102: 449–451. Schweigert, F.J., 1993, Effects of fasting and lactation on blood chemistry and urine composition in the grey seal (Halichoerus grypus), Comp. Biochem. Physiol., 105: 353–357. Seal, U.S., Swain, W.R., and Erickson, A.W., 1967, Hematology of the Ursidae, Comp. Biochem. Physiol. B, 22: 451. Seal, U.S., Erickson, A.W., Siniff, D.B., and Cline, D.R., 1971, Blood chemistry and protein polymorphisms in three species of Antarctic seals (Lobodon carcinophagus, Leptonychotes weddelli, and Mirounga leonia), in Antarctic Pinnipedia, Burt, W.H. (Ed.), American Geophysical Union, Baltimore, MD, 181–192. Silva, I.D., and Kuruwita, V.Y., 1993, Hematology, plasma and serum biochemistry values in free-ranging elephants (Elephas maximus cylonicus) in Sri Lanka, J. Zoo Wildl. Med., 24: 434–439. Slappendal, R.J., 1988, Disseminated intravascular coagulation, Vet. Clin. North Am. Small Anim. Pract., 18: 169–184. Stetter, M., Mangold, B., Miller, M., Weber, M., and Capobianco, J., 1999, Successful treatment of lead toxicosis in a bottlenose dolphin (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 146–147. Sweeney, J.C., 1974, Common diseases of pinnipeds, J. Am. Vet. Med. Assoc., 165: 805–810. Sweeney, J.C., and Ridgway, S.H., 1975, Common diseases of small cetaceans, J. Am. Vet. Med. Assoc., 167: 533–540. Taylor, R.F., and Farrell, R.K., 1973, Light and electron microscopy of peripheral blood neutrophils in a killer whale affected with Chediak-Higashi syndrome, Fed. Proc., 32: 822. Thompson, P.M., Tollit, D.J., Corpe, H.M., Reid, R.J., and Ross, H.M., 1997, Changes in haematological parameters in relation to prey switching in a wild population of harbour seals, Functional Ecol., 11: 743–750. Townsend, F.I., 1999, Medical management of stranded small cetaceans, in Zoo and Wild Animal Medicine, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 485–493. Townsend, F.I., and Petro, S., 1998, Significant panhypoproteinemia and regenerative anemia secondary to duodenitis in rough-tooth dolphins (Steno bredanensis), in Proceedings of the International Association for Aquatic Animal Medicine, 29: 157. Tyler, J.C., 1960, Erythrocyes and hemoglobin in the crabeater seal, J. Mammal., 41: 527. Vallyathan, N.V., George, J.C., and Ronald, K., 1969, The harp seal, Pagophilus groenlandicus V. Levels of haemoglobin, iron, certain metabolites and enzymes in the blood, Can. J. Zool., 47: 1193–1196. Walsh, M., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., Murphy, D., and Innis, S.M., 1999, Pneumotosis intestinalis in orphan manatees (Trichechus manatus), diagnosis, pathological findings and potential therapy, in Proceedings of the International Association for Aquatic Animal Medicine, 30: 1.
0839_frame_C19 Page 436 Tuesday, May 22, 2001 11:15 AM
436
CRC Handbook of Marine Mammal Medicine
White, J.R., Harkness, D.R., Isaacks, R.E., and Duffield, D.A., 1976, Some studies on the blood of the Florida manatee (Trichechus manatus latirostris), Comp. Biochem. Physiol. A, 55: 413–417. Wickham, J.J., Costa, D.P., and Elsner, R., 1990, Blood rheology of captive and free-ranging northern elephant seals and sea otters, Can. J. Zool., 68: 375–338. Williams, D.A., 1994, Diagnosis and management of pancreatitis, J. Small Anim. Pract., 35: 445–449. Williams, T.D., and Pulley, L.T., 1983, Hematology and blood chemistry in the sea otter (Enhydra lutris), J. Wildl. Dis., 19: 44–50. Williams, T.M., Antonelis, G.A., and Balke, J., 1994, Health evaluation, rehabilitation, and release of oiled harbor seal pups, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 227–241. Worthy, G.A.J., and Lavigne, D.M., 1982, Changes in blood properties of fasting and feeding harp seal pups, Phoca groenlandica after weaning, Can. J. Zoo., 60: 586–592.
0839_frame_C20.fm Page 437 Tuesday, May 22, 2001 11:50 AM
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).
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
437
0839_frame_C20.fm Page 438 Tuesday, May 22, 2001 11:50 AM
438
CRC Handbook of Marine Mammal Medicine
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,
0839_frame_C20.fm Page 439 Tuesday, May 22, 2001 11:50 AM
Cetacean Cytology
439
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
0839_frame_C20.fm Page 440 Tuesday, May 22, 2001 11:50 AM
440
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C20.fm Page 441 Tuesday, May 22, 2001 11:50 AM
441
Cetacean Cytology
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
0839_frame_C20.fm Page 442 Tuesday, May 22, 2001 11:50 AM
442
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C20.fm Page 443 Tuesday, May 22, 2001 11:50 AM
Cetacean Cytology
443
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).
0839_frame_C20.fm Page 444 Tuesday, May 22, 2001 11:50 AM
444
CRC Handbook of Marine Mammal Medicine
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).
0839_frame_C20.fm Page 445 Tuesday, May 22, 2001 11:50 AM
Cetacean Cytology
445
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
0839_frame_C20.fm Page 446 Tuesday, May 22, 2001 11:50 AM
446
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C20.fm Page 447 Tuesday, May 22, 2001 11:50 AM
Cetacean Cytology
447
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.
0839_frame_C20.fm Page 448 Tuesday, May 22, 2001 11:50 AM
448
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C21.fm Page 449 Tuesday, May 22, 2001 11:50 AM
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
449
0839_frame_C21.fm Page 450 Tuesday, May 22, 2001 11:50 AM
450
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C21.fm Page 451 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols 451
0839_frame_C21.fm Page 452 Tuesday, May 22, 2001 11:50 AM
452
CRC Handbook of Marine Mammal Medicine
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.)
0839_frame_C21.fm Page 453 Tuesday, May 22, 2001 11:50 AM
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).
0839_frame_C21.fm Page 454 Tuesday, May 22, 2001 11:50 AM
454
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C21.fm Page 455 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols
455
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.
0839_frame_C21.fm Page 456 Tuesday, May 22, 2001 11:50 AM
456
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C21.fm Page 457 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols
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).
0839_frame_C21.fm Page 458 Tuesday, May 22, 2001 11:50 AM
458
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C21.fm Page 459 Tuesday, May 22, 2001 11:50 AM
459
Gross Necropsy and Specimen Collection Protocols
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
0839_frame_C21.fm Page 460 Tuesday, May 22, 2001 11:50 AM
460
CRC Handbook of Marine Mammal Medicine
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
Gross Necropsy and Specimen Collection Protocols
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
0839_frame_C21.fm Page 461 Tuesday, May 22, 2001 11:50 AM
461
0839_frame_C21.fm Page 462 Tuesday, May 22, 2001 11:50 AM
462
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C21.fm Page 463 Wednesday, May 23, 2001 10:45 AM
Gross Necropsy and Specimen Collection Protocols 463
0839_frame_C21.fm Page 464 Tuesday, May 22, 2001 11:50 AM
464
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C21.fm Page 465 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols
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
0839_frame_C21.fm Page 466 Wednesday, May 23, 2001 10:46 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C21.fm Page 467 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols
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
0839_frame_C21.fm Page 468 Wednesday, May 23, 2001 10:47 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C21.fm Page 469 Tuesday, May 22, 2001 11:50 AM
Gross Necropsy and Specimen Collection Protocols
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.
0839_frame_C21.fm Page 470 Tuesday, May 22, 2001 11:50 AM
470
CRC Handbook of Marine Mammal Medicine
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.
0839_frame_C22.fm Page 471 Wednesday, May 23, 2001 10:47 AM
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
Bossart et al., 1998
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)
0839_frame_C22.fm Page 472 Tuesday, May 22, 2001 11:51 AM
CRC Handbook of Marine Mammal Medicine
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
Bossart et al., 1998
Shlosberg et al., 1997
Bossart et al., 1998
Shlosberg et al., 1997
0839_frame_C22.fm Page 473 Tuesday, May 22, 2001 11:51 AM
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)
0839_frame_C22.fm Page 474 Tuesday, May 22, 2001 11:51 AM
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C22.fm Page 475 Tuesday, May 22, 2001 11:51 AM
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)
0839_frame_C22.fm Page 476 Tuesday, May 22, 2001 11:51 AM
476 CRC Handbook of Marine Mammal Medicine
0839_frame_C22.fm Page 477 Tuesday, May 22, 2001 11:51 AM
Toxicology
477
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).
0839_frame_C22.fm Page 478 Tuesday, May 22, 2001 11:51 AM
478
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C22.fm Page 479 Tuesday, May 22, 2001 11:51 AM
Toxicology
479
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)
0839_frame_C22.fm Page 480 Tuesday, May 22, 2001 11:51 AM
480
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C22.fm Page 481 Tuesday, May 22, 2001 11:51 AM
Toxicology
481
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).
0839_frame_C22.fm Page 482 Tuesday, May 22, 2001 11:51 AM
482
CRC Handbook of Marine Mammal Medicine
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 <80% lipid than 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
0839_frame_C22.fm Page 483 Tuesday, May 22, 2001 11:51 AM
Toxicology
483
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,
0839_frame_C22.fm Page 484 Wednesday, May 23, 2001 10:48 AM
484
CRC Handbook of Marine Mammal Medicine
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
0839_frame_C22.fm Page 485 Tuesday, May 22, 2001 11:51 AM
Toxicology
485
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 <1 ppm in recent times). trans-Nonachlor is typically a major contributor to total chlordane compound residues in blubber. The insecticide toxaphene is a mixture of chlorinated norbornane derivatives or polychlorinated camphenes. These compounds have been reported from marine mammal tissues from all over the globe (O’Shea et al., 1980; de Boer and Wester, 1993; Jarman et al., 1996a; Vetter et al., 1999). Toxaphene-related compounds are deposited over great distances by atmospheric transport and occur in low concentrations in blubber of marine mammals from remote areas of the Arctic. In some northern latitudes they can occur at equivalent or greater concentrations than sumDDTs or PCBs (which are typically low in such areas) (Muir et al., 1988a; 1990; 1992b; Zhu and Norstrom, 1993; Wade et al., 1997). The insecticide HCH occurs in various isomers in marine mammal tissues. The pure γ-isomer of HCH is also referred to as benzene hexachloride (BHC) or lindane. These compounds can appear in low concentrations (usually <1.0 ppm) in blubber (Duinker et al., 1989; Simmonds et al., 1994; Jarman et al., 1996a; Muir et al., 1996a,b; Tanabe et al., 1996; Vetter et al., 1999). Other organochlorine insecticide residues reported infrequently and at low concentrations in marine mammals are kepone, mirex, endosulfan, and the fungicide or the organochlorine by-product hexachlorobenzene (HCB) and related compounds (often reported as total chlorobenzenes, CBz) (O’Shea et al., 1980; Muir et al., 1990; 1996a,b; Woodley et al., 1991; Bratton et al., 1993; de Kock et al., 1994; Kuehl et al., 1994; Aono et al., 1997). Because of the herbivorous food habits of sirenians and the low biomagnification of organochlorines in plants, only a few organochlorine pesticide residues, other than DDE, have been reported from blubber of sirenians. Dieldrin, HCB, and lindane have been found, but only at very low concentrations (Miyazaki et al., 1979; O’Shea et al., 1984; Ames and Van Vleet, 1996). Sea otters (Enhydra lutris) do not have blubber, but analyses of livers have revealed modest concentrations of DDT and metabolites (varying with location) and low levels of dieldrin, HCB, HCHs, and chlordane compounds (Jarman et al., 1996b; Estes et al., 1997; Nakata et al., 1998). Polar bears have a wide range of organochlorine pesticides and metabolites in liver and adipose tissues, but in low concentrations (Muir et al., 1988b; Norstrom et al., 1988; 1998; Norheim et al., 1992; Letcher et al., 1998). These include DDT and metabolites, dieldrin, HCH isomers, HCB, and other chlorobenzenes. Chlordane compounds and metabolites (oxychlordane, nonachlors, heptachlor epoxide, photoheptachlor, others) are particularly predominant. There are complex patterns in the relative concentrations of organochlorines in polar bears, which vary by location, sex, and age, and can differ from those found in cetaceans or pinnipeds.
Polychlorinated Biphenyls PCBs are a group of up to 209 congeners produced by the industrial chlorination of biphenyls. They once had wide applications in industry, including use in electrical transformers, capacitors, hydraulics, heat transfer systems, plastics, and inks. Most industrialized nations banned or suspended their production in the 1970s and 1980s due to recognition of their widespread occurrence as environmental contaminants, and heavy publicity of accidental mass poisonings of human foods. Despite reductions in manufacturing, the products and systems in which
0839_frame_C22.fm Page 486 Tuesday, May 22, 2001 11:51 AM
486
CRC Handbook of Marine Mammal Medicine
PCBs were originally employed continue to hold enormous quantities of these substances, and are slowly degrading and releasing them to the environment. Thus, the amounts that reach the oceans will continue to increase with time (Tanabe, 1988). PCBs are very lipophilic and are quickly incorporated into marine food webs; they have become some of the most common and abundant environmental contaminants reported in tissues of marine mammals. There is wide variance among individual congeners in their toxicity to laboratory mammals and persistence in marine mammal tissues. PCBs (primarily administered as commercial mixtures) are well known to produce toxic effects (including lethality) in laboratory exposures of select species of mammals. Pathological effects with greatest potential impacts on mammal populations center around reproductive impairment, which PCBs are known to cause through alteration of menstrual cycles, embryo absorption, abortion, still births, and impaired growth and survival of young (Allen et al., 1974; 1980; Allen and Barsotti, 1976; Barsotti et al., 1976; Aulerich and Ringer, 1977; Bleavins et al., 1980; 1982; Aulerich et al., 1985; Linzey, 1987; 1988; Kihlström et al., 1992; O’Hara and Rice, 1996). Dosage required to produce toxic effects varies widely, even among closely related species, and in some mammals toxic effects have not been seen (although the numbers of experiments are limited). Earlier demonstrations of toxicity may have been confounded by the presence of even more toxic contaminants, such as polychlorinated dibenzodioxins (PCDDs) and dibenzofurans (PCDFs), within experimentally administered PCB formulations. No experimental determinations of effects of PCBs in isolation have been established for marine mammals (a few studies of harbor seals have used market fish contaminated “naturally” with mixtures of PCBs and multiple compounds of other classes). Nevertheless, a host of effects or responses have been reported in humans and other animals (sometimes requiring high doses to elicit). These effects have included lesions of the integument (chloracne, hyperkeratosis, nail deformities), reproductive disorders, immunotoxicity, teratogenicity, and likely carcinogenicity (Fuller and Hobson, 1986; Peakall, 1992; Burns et al., 1996; Pitot and Dragan, 1996). Since their discovery as persistent contaminants in the mid-1960s, PCBs have been reported in blubber of thousands of marine mammals of many species throughout the world (see O’Shea, 1999). Examples of maximum concentrations found in blubber of individual marine mammals (ppm total PCBs by wet weight unless otherwise indicated) include 145 to 450 ppm in small cetaceans and pinnipeds from southern California during the 1970s (DeLong et al., 1973; O’Shea et al., 1980), 240 to 800 ppm in belugas from the St. Lawrence River from the 1970s to the 1990s (Sergeant, 1980; Martineau et al., 1987; Muir et al., 1996a), 410 ppm in killer whales (Orcinus orca) from near Japan in 1986 (Kannan et al., 1989), as high as 3000 ppm (lipid weight) in diseased striped dolphins from the Mediterranean Sea in 1990 (Kannan et al., 1993b; Aguilar and Borrell, 1994a), and 770 to 5300 ppm (lipid weight) in pinnipeds from the Baltic and North Seas in the 1960s to the 1980s (Koeman, 1973; Blomkvist et al., 1992). Highest concentrations are typically found in males, piscivorous species from inshore locations, and in diseased individuals in poor body condition. Lower concentrations are found in female cetaceans and pinnipeds, and are usually orders of magnitude lower than in males. Marine mammals from more pelagic or pristine areas, baleen whales, and sirenians also tend to have lower levels (O’Shea et al., 1984; O’Shea and Brownell, 1994; Haynes et al., 1999; Tilbury et al., 1999; Vetter et al., 1999; O’Hara et al., 1999; Weisbrod et al., 2000a,b). PCBs are found in sea otters, particularly in the Aleutian Islands and Monterey Bay where up to 5.9 ppm (wet weight) have been reported in liver (Jarman et al., 1996b; Estes et al., 1997; Nakata et al., 1998). PCBs are the predominant organohalogens reported in polar bears (Norstrom et al., 1998). Prior to the late 1980s, most studies typically quantified PCB concentrations in marine mammals by comparison with a standard commercial mixture of congeners such as Aroclor 1254. Recent studies include congener-specific determinations. There is usually little variation
0839_frame_C22.fm Page 487 Tuesday, May 22, 2001 11:51 AM
Toxicology
487
in the proportions of various congeners relative to total PCBs among various tissues and organs within individuals, and these proportions often show low variation by age or sex within species in any particular region or study (Muir et al., 1988a; 1996b; Duinker et al., 1989; Salata et al., 1995; Gauthier et al., 1997). Lower chlorinated PCBs are seldom found; penta-, hexa-, and heptachlorobiphenyls predominate (Granby and Kinze, 1991; Wells and Echarri, 1992; Falandys et al., 1994) because of their greater resistance to metabolic degradation. In odontocetes, pinnipeds, and polar bears, the hexachlorinated di-ortho congeners 153, 138, and 180 often make up the highest proportion, whereas the non-ortho or coplanar PCBs 77, 126, and 169, which are considered the most toxic (Green et al., 1996), usually constitute a very small fraction of total PCBs (Norstrom et al., 1988; Kannan et al., 1993b; Falandysz et al., 1994; Borrell et al., 1996; van Scheppingen et al., 1996; Gauthier et al., 1997; Bernt et al., 1999; Vetter et al., 1999; Cleeman et al., 2000). Metabolism of PCBs can differ across groups and species of marine mammals (Boon et al., 1992; 1994; 1997; Wells et al., 1994). In addition to various PCB congeners, methyl sulfone metabolites of PCBs have been quantified in tissues of odontocetes, pinnipeds, and polar bears (Bergman et al., 1994; Letcher et al., 1995; 1998; Troisi et al., 1998).
Other Organohalogens The PCDFs, PCDDs, polychloroquaterphenyls (PCQs), and polychlorinated naphthalenes (PCNs) are by-products of combustion, PCB manufacture, and other industrial processes (Safe, 1986; 1991) that have been reported in tissues of marine mammals. Concentrations are typically low in comparison with PCBs and organochlorine pesticides and metabolites, and usually represent only a small fraction of TEQs (toxic equivalents) (Green et al., 1996). One of the PCDDs, 2,3,7,8-tetrachloro-dibenzo-p-dioxin (TCDD), is ranked as one of the most toxic compounds known and has been reported in low concentrations (ppt) in marine mammal tissues (Kannan et al., 1989; Buckland et al., 1990; Norstrom et al., 1990; Bergek et al., 1992; Oehme et al., 1995; Jarman et al., 1996a). PCDDs occur at greater concentrations in marine mammals of the high Arctic than in those of the sub-Arctic, probably in part because of atmospheric transport over the pole from urban sources in Europe and Asia (Norstrom et al., 1990) and possibly from North American sources (Muir, pers. comm.). Subcutaneous fat biopsies of South American sea lions (Otaria byronia) from coastal Argentina had low concentrations of PCDDs (averaging 6 to 17 ppt wet weight) and PCDFs (averaging 5 to 21 ppt), with greater numbers of congeners found in sea lions at a more industrialized area (Jimènez et al., 1999). PCDDs and PCDFs were reported at very low concentrations in livers of sea otters (Jarman et al., 1996b; Bacon et al., 1999), and are also found at low concentrations in polar bears (Norstrom et al., 1990; Letcher et al., 1996). PCDDs and PCDFs have been found in blubber of dugongs (Dugong dugon) from northeastern Australia at concentrations ranging up to 390 ppt (Haynes et al., 1999). The maximum concentration of 250 ppt PCDD in one of these dugongs exceeds maximum estimates for PCDDs in blubber of cetaceans and pinnipeds. Dugongs also have low PCB concentrations in blubber and thus, unlike other marine mammals, TEQs (see below) are attributed primarily to the PCDDs/PCDFs rather than PCBs. This unusual finding has not been fully explained, but may relate to feeding close to sediments, inputs from adjacent agricultural regions, biotransformation from precursors during hindgut digestion, or unique metabolic degradation capacities (Haynes et al., 1999). Tris-(4-chlorophenyl)methane (TCPMe) and tris-(4-chlorophenyl)methanol (TCPMeOH) have been detected in the tissues of pinnipeds, odontocetes, and polar bears from widely separated regions (Walker et al., 1989; Jarman et al., 1992; Zook et al., 1992; de Boer et al.,
0839_frame_C22.fm Page 488 Tuesday, May 22, 2001 11:51 AM
488
CRC Handbook of Marine Mammal Medicine
1996; Muir et al., 1996a; Watanabe et al., 1999). These compounds appear to accumulate in lipids and are usually highest in blubber (Watanabe et al., 1999). Concentrations (lipid weight) of TCPMe or TCPMeOH typically vary between 1 and 300 ppb in blubber, and 6.0 ppm in polar bear liver (Jarman et al., 1992; de Boer et al., 1996; Muir et al., 1996a; Falandysz et al., 1999; Minh et al., 1999; Watanabe et al., 1999). They have been found in marine mammals collected as early as 1952. The origins of these compounds are incompletely understood; they may stem from impurities in the manufacture of DDT or dicofol, or the synthesis of agrochemicals, polymers, and dyes (Jarman et al., 1992). The existence of toxic effects associated with concentrations observed in marine mammals is unknown. Other lipophilic organohalogens reported in marine mammal tissues include the polybrominated biphenyls (PBBs), octachlorostyrene (OCS), and polybrominated diphenyl ethers (PBDEs) (Kuehl et al., 1991; Kuehl and Haebler, 1995; Jarman et al., 1996a). These compounds occur at low concentrations, typically in low ppb quantities, with maxima of 1.3 ppm PBBs and 0.5 ppm OCS reported in blubber of bottlenose dolphins, and 3.0 to 16.3 ppm PBDEs in blubber of bottlenose dolphins and long-finned pilot whales (Kuehl and Haebler, 1995; Lindström et al., 1999). No links have been established between their presence in tissues and toxic effects in the cases studied. Extractable organobromines, organoiodines, and organochlorines have been measured in beluga tissues, but source compounds have not been fully determined (Kiceniuk et al., 1997). Polychlorinated terphenyls (PCTs) have been reported at low concentrations (about 40 ppb dry weight) in livers of sea otters from California and the Aleutian Islands (Jarman et al., 1996b) and at ≤1 ppm (lipid weight basis) in fat of gray seals from Sweden (Renberg et al., 1978). These compounds have been used as flame retardants in the past. Modern sources are unknown, but those detected in sea otters may be from atmospheric depositions or shipyard facilities (Jarman et al., 1996b).
Effects of Organochlorines on Metabolism Information on the metabolism and biochemical toxicity of organohalogens in cetaceans and pinnipeds has been reviewed by Boon et al. (1992; 1994), Reijnders (1994), Marsili and Focardi (1997), Brouwer (1999), and Busbee et al. (1999). Some organochlorines (especially PCBs) induce enzymes that could lead to endocrine imbalances. The mixed-function oxidase (MFO) system is the chief enzymatic pathway involved. Cytochrome P-450 enzymes of this system bind with oxygen and organochlorine substrates to convert the latter into more polar compounds that are more easily excreted (classical “Phase I” drug metabolism involves oxidation by these enzymes, whereas “Phase II” involves subsequent conjugation of a polar substrate by additional enzymes). The potential interactions between different compounds and enzymes present in marine mammal tissues are likely to be very complex; various organochlorine compounds can act as enzyme inhibitors, inducers, or substrates themselves, and their intermediate metabolites can be even more toxic. The MFO activity increases the capacity to detoxify multiple compounds, and this ability to metabolize organohalogens increases with increasing exposure. For example, elevated rates of metabolism of pentachlorobiphenyls by induced MFO activity in cetaceans heavily contaminated by PCBs has been proposed as an explanation for lower proportions of these congeners in tissues (Muir et al., 1990; Kannan et al., 1993b; Borrell et al., 1996). The MFO induction has been well studied in other mammals, and rates of induction and organohalogen metabolism can vary with species, genetic subdivisions, sex, age, reproductive state, diet, history of previous exposure, physical condition, and other factors (Nebert and Gonzalez, 1987; Rattner et al., 1989; Aguilar et al., 1999). Induction of MFO enzymes can result in altered rates of metabolism of endogenous compounds (including steroid hormones), and result in subsequent abnormalities in physiological and developmental processes.
0839_frame_C22.fm Page 489 Wednesday, May 23, 2001 10:49 AM
Toxicology
489
The toxic equivalents and toxic equivalency concepts have been applied to estimate potential toxic significance of PCBs, polychlorinated dibenzo-p-dioxins (PCDDs), and related compounds in marine mammals (Tanabe et al., 1987; Beckmen et al., 1999; Watanabe et al., 1999; Finklea et al., 2000). A toxic equivalency factor (TEF) can be calculated for a compound based on the level required to elicit a specific biological response, as compared with the level of tetrachlorodibenzo-p-dioxin (TCDD) to elicit the same response; the TCDD factor is set at 1.0. Biomarkers of MFO activity are often used as such responses. An overall estimate of the TCDD-like equivalents expressed as “TCDD” in micrograms of the full mixture of PCBs, PCDDs, and PCDFs present in marine mammal tissue samples constitutes the TEQ, calculated as the sum of the concentration of each PCB, PCDD, or PCDF, times its TEF. There are a number of caveats to the use of TEFs to estimate TEQs, in part because of different effects in different enzyme systems, in vitro simplicity vs. in vivo complexity, and species-specific variability in responses (Reijnders, 1994; Safe, 1994; Kamrin and Ringer, 1996). Toxic responses can be nonadditive as a result of antagonistic effects within a complex mixture. For example, immunotoxic responses can be much less than predicted on the basis of total TEQs (Safe, 1994). Bearing such cautions in mind, comparisons based on the TEF concept have focused on the coplanar PCBs, with non-ortho chlorine substitution in the biphenyl rings (especially those with four or more chlorine atoms in the para and meta positions) of greatest interest, because of structural similarity to TCDD. Responses to these congeners in rat liver systems include MFO activity induction, and are similar to TCDD but require higher doses. PCB 126 is the most potent congener, at 0.1 TEF. However, amounts of this and other coplanar PCBs (e.g., mono and di-ortho substitutions) occur in only trace amounts in marine mammals, typically three to five orders of magnitude lower than total PCBs, but in greater concentrations than PCDDs or PCDFs (Tanabe et al., 1987; 1988; Kannan et al., 1989). Some studies suggest that odontocetes seem to have a lower capacity than seals to metabolize congeners with neighboring H atoms in meta and para positions (Duinker et al., 1989; Boon et al., 1997). Highly chlorinated biphenyls in general are more slowly degraded or excreted than compounds with fewer chlorines. Polar bears, however, have a capacity to metabolize PCB congeners that are recalcitrant to metabolism in some seals and cetaceans (Norstrom et al., 1988; Letcher et al., 1996). Patterns of MFO activity in liver microsomal samples and skin biopsies have been determined for several marine mammals (Fossi and Marsili, 1997). These can be considered biomarkers of exposure. Hepatic cytochrome P-450, NADPH cytochrome c reductase, aniline hydroxylase (AH), arylhydrocarbon hydroxylase (AHH), aldrin epoxidase (ALDE), and ethoxyresorufin-odeethylase (EROD) have been measured in liver microsomal samples of short-finned pilot whales (Globicephala macrorhyncus), striped dolphins, and a killer whale (Watanabe et al., 1989). White et al. (1994) determined AHH, cytochrome P-450, cytochrome b5, EROD, estradiol- 2-hydroxylase (E2-2OH), and pentoxyresorufin-o-depentylase (PROD) from belugas of the Canadian Arctic, and verified that cytochrome P-4501A (CYP1A) activity was correlated with PCB concentrations in tissues. Benzo(a)pyrene monooxygenase (BPMO) activity was deter- mined in biopsy skin samples of striped dolphins and fin whales from the Mediterranean Sea, with associations found between BPMO activity and PCB concentrations in blubber (Fossi et al., 1992; Marsili et al., 1998). The hepatic EROD and cytochrome P-450 levels have been measured in gray seals, with the latter increasing with increasing PCB concentrations in blubber (Addison and Brodie, 1984; Addison et al., 1988). The hepatic EROD, cytochrome P-450 and cytochrome b5 have also been measured in harbor seals (Addison et al., 1986). Goksøyr et al. (1985; 1986; 1988; 1989) measured hepatic and renal cytochrome P-450, cytochrome b5, NADPH cytochrome P-450 reductase, AHH, aminopyrine N-demethylase (APDM), biphenyl 4-hydroxylase (biph4OH), E2-2OH, ethoxycoumarin-o-deethylase (ECOD), EROD, glutathione-S-transferase
0839_frame_C22.fm Page 490 Wednesday, May 23, 2001 10:49 AM
490
CRC Handbook of Marine Mammal Medicine
(GSH-T), PROD, and UDP-glucuronyl transferase (UDPGT) in minke whales, but did not compare these with organohalogen concentrations. Similarly, hepatic enzyme activities (EROD, PROD, ECOD, UDP-GT, NADPH, and others) have been measured in hooded (Cystophora cristata) and harp seals, but not in relation to organohalogens (Goksøyr et al., 1992). In polar bears, however, correlations were established for activity of EROD, PROD, CYP1A activity and various organochlorines, including TEQs (Letcher et al., 1996). Harbor seals also exhibit correlations among PCBs in blubber or liver and P-450, P-420, CYP1A, CYP2B, EROD and ECOD activity (Troisi and Mason, 1997). Although the amount of information on relationships between organochlorine concentrations in tissues or diets of marine mammals and MFO activity is growing, the lack of knowledge about the degree to which induced enzyme activity corresponds with meaningful interference of endocrine metabolism in wild marine mammals has thus far made toxicological interpretation of such biomarker studies difficult. The overall impact of “induction” to the individual or populations of marine mammals is unknown, and this reduces the value of this biomarker from a management perspective. However, proposed negative impacts have included interferences with endocrine processes and reproduction, and possible interference with the immune system (Colborn and Smolen, 1996; Busbee et al., 1999) and these should be pursued to determine whether induction could be applied to impact assessment. Among organochlorine pesticides, the relatively low percentage of oxychlordane in cetaceans in comparison with other mammals and marine birds suggests that cetaceans have relatively little capability to metabolize trans-nonachlor to oxychlordane, even in St. Lawrence River belugas with potentially elevated MFO capability (Muir et al., 1990). Narwhals (Monodon monocerus) have a low capacity to transform cis-nonachlor (Muir et al., 1990; 1992b). In vitro microsomal assays and congener analyses have been used to estimate the potential for metabolism of toxaphene components in white-beaked dolphins, harbor porpoises, and sperm whales. Sperm whale microsomal assays showed no capacity for chlorobornane (CHBs) metabolism, and only one CHB compound was metabolized by dolphins and porpoises (Boon et al., 1998).
Effects of Organochlorines on Reproduction and Endocrine Function Observations suggesting that organochlorines were linked to reproductive disorders in marine mammals were first made in the early 1970s. Ringed seals in the Baltic Sea with uterine stenosis and occlusions had elevated concentrations of sumDDT in blubber (averaging 130 ppm lipid weight) and PCBs (110 ppm) in comparison with pregnant females (88 ppm sumDDT, 73 ppm PCBs) (Helle et al. 1976a,b; Bergman and Olsson, 1985). California sea lions (Zalophus californianus) aborting fetuses and producing stillbirths also had higher concentrations of PCBs (112 ppm wet weight) and sumDDT (824 ppm) than sea lions with normal births (103 ppm sumDDT, 17 ppm PCBs) (DeLong et al., 1973). However, these observations were confounded, in that females with impaired reproduction for any reason lack avenues for excretion of organochlorines through lactation and, therefore, can be expected to have higher concentrations. Some Baltic ringed seals with normal uteri were also not pregnant and did not have significantly lower organochlorine concentrations than those with uterine pathology. California sea lions with abortions and stillbirths were also younger than females with normal births, and the abortion-inducing disease leptospirosis was present in the population (see reviews by Addison, 1989; O’Shea and Brownell, 1998). Later studies of gray seals in the Baltic failed to find an association between uterine pathology and organochlorine contamination (Blomkvist et al., 1992). The lack of a correlation between presence of lesions and contamination could be due to no effect, or to an inadequate statistical power of the study. Among other potential
0839_frame_C22.fm Page 491 Tuesday, May 22, 2001 11:51 AM
Toxicology
491
endocrine-related impacts, Bergman and Olsson (1985) advanced the hypothesis that organochlorine exposure causes adrenocortical dysfunction in marine mammals. Skull asymmetry and bone lesions that may be related to hyperadrenocortical effects of organochlorines have been associated with the onset of pollution in seals of the Baltic Sea (Zakharov and Yablokov, 1990; Bergman et al., 1992; Mortensen et al., 1992; Olsson et al., 1994). Kuiken et al. (1993), however, found no association between adrenal hyperplasia and concentrations of organochlorines in harbor porpoises. A landmark experiment carried out on captive harbor seals evaluated the effects of organochlorines on reproduction. Reijnders (1986) fed seals control and treatment diets (higher organochlorine levels) of market fish from two different regions. Seals fed the second diet had lower reproductive success, likely due to failures at implantation (Reijnders, 1986) rather than from late-term failure seen in California sea lions. There have been no experimental studies on reproductive effects of organochlorines in cetaceans. Indirect evidence for impaired reproduction in cetaceans is also limited. In Dall’s porpoises, Subramanian et al. (1987) obtained a weak negative correlation between testosterone concentrations in blood and DDE concentrations in blubber of 12 males collected in 1984 in the North Pacific, but no correlation between PCBs and testosterone. Martineau et al. (1987; 1994), Béland et al. (1993), and others suspect that elevated organochlorines (PCBs, in particular) have impacted reproduction in belugas in the St. Lawrence River. Béland et al. (1991; 1993) provided data on stranded adult female belugas from the St. Lawrence that included several observations of reproductive pathology, but no evidence of a cause-and-effect relationship. Recent reports indicate the St. Lawrence estuary beluga population is growing at approximately 10 to 20 belugas per year (Kingsley, 1998; 1999). Despite such limited data, it has been speculated that organochlorines may be widely affecting reproduction and health in cetaceans globally through endocrine disruption (Colborn and Smolen, 1996). However, the concept of endocrine disruption in mammals at typical environmental exposure levels is controversial (National Research Council, 1999). To test this hypothesis further in marine mammals, a recent workshop suggested that necropsy and health examinations routinely search for a number of developmental and reproductive end points of endocrine disruption, including several sets of morphometric data and inspections for malformations of genitalia (e.g., hypospadia in males) (see report of working group on endocrinology and reproduction in O’Shea et al., 1999).
Effects of Organochlorines on Immunocompetence and Epizootics Some organochlorine compounds have immunotoxic effects in laboratory animals; there is thus much interest in the possibility that organochlorines (particularly, PCBs) may render marine mammals more susceptible to disease through immunosuppression (Simmonds and Mayer, 1997; Busbee et al., 1999). Immunosuppression has been experimentally demonstrated in captive harbor seals fed market fish from two different regions contrasting in concentrations of a variety of organochlorines (Brouwer et al., 1989; de Swart et al., 1993; 1994; 1996; Ross et al., 1996). Cellular immunity was affected more than humoral immunity. Seals fed the diet with higher levels of organochlorines had lower serum vitamin A, and lower white blood cell and granulocyte counts (but not lymphocytes or monocytes). Natural killer cell activity and lymphocyte function assays after mitogen stimulation were also lower in seals exposed to higher levels of organochlorines. Field studies of potential relationships between organochlorine exposure and effects on the immune system or deaths due to disease have not shown consistent patterns. Immunocompetence and cumulative exposure to PCBs through milk were examined in wild, nursing gray seal
0839_frame_C22.fm Page 492 Tuesday, May 22, 2001 11:51 AM
492
CRC Handbook of Marine Mammal Medicine
pups. No significant relationships were found between PCB exposure and responses to challenges with morbillivirus vaccine, mitogen stimulation, or various other measures of health (Hall et al., 1997). Studies of harbor porpoises that died from infectious disease or parasitism in comparison with those that died from trauma (primarily in fisheries) around Great Britain from 1989 to 1992 did not show statistically significant differences in concentrations of organochlorines (25 PCB congeners and seven pesticides or metabolites) in blubber. However, sample sizes greater than the nearly 100 animals examined were probably required (only a limited geographic subset could be fully utilized) to partition various potential sources of variation (Kuiken et al., 1994). Also, animals that died of trauma were not necessarily free of infectious disease. In contrast, a second study of this same population based on samples from fresh carcasses collected from 1990 through 1996 (n = 67), showed significantly higher concentrations of sumPCBs and of 16 of 25 PCB congeners in harbor porpoises that died from infectious disease. No associations were found between sumPCBs and other variables, such as sex, age, region, year, season, and nutritional status, although the investigators suggested that additional data amenable to more powerful statistical analyses would improve conclusions (Jepson et al., 1999). Correlations were obtained between concentrations of certain organochlorines (DDT, DDE, o,p′-DDE, and higher-chlorinated PCBs) and reduced immune responses, as measured by in vitro mitogen-induced proliferation responses of lymphocyte cultures, from five freeranging male bottlenose dolphins from Florida (Lahvis et al., 1995). Effects of exposure to PCB 138, 153, 180, and 169 on immune functions of beluga peripheral blood leukocytes and splenocytes in vitro were reported by De Guise et al. (1998). Only PCB 138 reduced proliferative responses of beluga splenocytes, and none affected cell proliferation. A mixture of PCBs 138, 153, and 180 reduced splenocyte proliferation, but a mixture of these same congeners plus PCB 169 did not have this effect, illustrating complexities in predicting impacts of contaminant mixtures. Similarly, antagonistic properties of the mixture were among possible explanations when laboratory rats fed a diet with lipid containing organohalogen extracted from blubber of St. Lawrence River belugas showed no significant effects in multiple immune function assays (Lapierre et al., 1999). In vitro genotoxicity (micronuclei assay of DNA damage) to beluga skin fibroblast cultures treated with chlordane, DDT, or toxaphene has been reported, although exposures were generally higher than concentrations expected to occur in blood of St. Lawrence River belugas, and responses were greatly reduced in experiments that included the presence of an external metabolic factor (Gauthier et al., 1999a). Gauthier et al. (1999b) used similar assays of genetic damage (micronuclei, sister chromatid exchange, and/or chromosome aberration assays) to compare DNA damage in blood lymphocytes of Arctic belugas, bottlenose dolphins, and Atlantic gray and harp seals. This study established important baseline information and these assays should be applied when genotoxic mechanisms are implicated. In addition to linkages between immune suppression and infectious disease, possible relationships between cancers and organochlorine exposure have been hypothesized for belugas in the St. Lawrence River (Martineau et al., 1999) and California sea lions (Gulland et al., 1996). Attempts have been made to link recent mass die-offs of marine mammals caused by morbilliviruses with organochlorine exposure. However, morbilliviruses are extremely virulent in immunologically naive populations of mammals, directly damaging lymphoid tissue (thus causing immunosuppression and secondary infections in their own right) and have caused extremely high mortality of susceptible populations of terrestrial mammals prior to the synthesis of PCBs (Kennedy, 1999). Organochlorine concentrations in blubber of harbor seals that died in the 1988 phocine distemper virus (PDV) outbreak in Great Britain did not show significant differences from those that survived (Hall et al., 1992). Other studies have found
0839_frame_C22.fm Page 493 Tuesday, May 22, 2001 11:51 AM
Toxicology
493
no statistical associations between organochlorine concentrations in juvenile harbor seals collected before and during a PDV outbreak, and the addition of PCBs to the diet of captive harbor seals did not affect susceptibility to morbillivirus challenge (Blomkvist et al., 1992; Harder et al., 1992). Whereas thymulin production, important for development of T cells, was negatively correlated to morbillivirus titers in gray seals, organochlorines had no statistical effect on this relationship. No correlation was found between thymulin and organochlorine concentrations in blubber of harbor seals (Kendall et al., 1992). Concentrations of PCBs (especially coplanar forms) in blubber of striped dolphins from the Mediterranean Sea that died during a morbillivirus epidemic in the early 1990s were extremely high compared with other marine mammals, and with individuals sampled in the area by biopsy before and after the epidemic. This suggests that PCBs may have influenced susceptibility to morbillivirus, although other hypotheses have also been advanced to explain these differences (Kannan et al., 1993b; Aguilar and Borrell, 1994a).
Biotoxins The study of poisons from animals is “zootoxicology” and from living things in general is “biotoxicology” (Fowler, 1993); thus these complex chemical mixtures are called “biotoxins.” Many of these biotoxins play a yet-to-be-understood role in predator/prey relationships, food gathering, and defense mechanisms (Fowler, 1993). The naturally produced toxins originating from protozoans have been associated with seafoods for centuries (see review of marine toxins, Baden et al., 1995). These substances are often acutely toxic, but are currently poorly characterized from a toxicological perspective (Iverson and Truelove, 1994). Dinoflagellates and diatoms are important ecosystem components. Under certain conditions, however, “blooms” can occur so that the density of organisms is so great the color of the water changes, causing phenomena collectively referred to as “red tides,” although few are actually red. Such red tides can be associated with a number of different species of dinoflagellates and diatoms. Off North America, the responsible organisms commonly involved are Gonyaulax catenella (Pacific Coast), Gonyaulax tamarensis (Atlantic Coast), and Gymnodinium breve (previously referred to as Ptychodiscus brevis, in the Gulf of Mexico and Florida) (Fowler, 1993). Most biotoxins act acutely in the target species of concern following direct exposure or significant accumulation in prey (e.g., fish, clams). Exposure testing for toxins include in vivo assays (mouse bioassay), enzyme-linked immunosorbent assay (ELISA), and in vitro binding assays to biological receptors and high-performance liquid chromatography (HPLC) (Baden et al., 1995; Bossart et al., 1998; Costas and Lopez-Rodas, 1998; Hernandez et al., 1998; Scholin et al., 2000).
Brevetoxin Brevetoxins (brevetoxin A, B, etc., and hemolytic components) have both neurotoxic and hemolytic actions. Brevetoxin in general is a sodium channel activator that occurs in some dinoflagellates (e.g., G. breve) in the Gulf of Mexico, near Japan and New Zealand, and in other regions. Brevetoxins accumulate in shellfish, and can cause acute poisoning in mammals. Neurological and gastrointestinal signs predominate (Baden et al., 1995). Inhalation results in a unique clinical presentation: conjuctival irritation, rhinorrhea, nonproductive cough, and bronchoconstriction (Baden et al., 1995). Brevetoxin has high binding affinity in vitro to marine mammal brain tissue (Trainer and Baden, 1999). In 1996, 149 manatees (Trichecus manatus) died along the southwest coast of Florida concurrent with a bloom of dinoflagellates (G. breve) (Bossart et al., 1998). Levels of
0839_frame_C22.fm Page 494 Tuesday, May 22, 2001 11:51 AM
494
CRC Handbook of Marine Mammal Medicine
brevetoxin were considered elevated by 2- to 15-fold in manatee stomachs and tissues. Severe congestion of the nasopharynx, lungs, liver, kidney, and brain, and edema and hemorrhage of the nasopharynx and lung were present. The most common histological lesions noted were catarrhal rhinitis, pulmonary hemorrhage and edema, multiorgan hemosiderosis, and a nonsuppurative leptomeningitis (Bossart et al., 1998). Antibody staining of histological specimens revealed brevetoxin in lymphocytes and macrophages of the lung, liver, secondary lymphoid tissues, and inflammatory lesions of the nasal mucosa and meninges. These data indicate that the manatees very likely suffered from brevetoxicosis, that expression of such intoxications is not always acute, and that the route of exposure was likely inhalation (Bossart et al., 1998). Based on examination of live affected animals, a neurological syndrome (muscle fasciculations, incoordination, loss of righting reflex) may be present acutely followed by hemopathy (anemia, hemosiderosis) (Bossart et al., 1998). Brevetoxicoses have also been implicated in less well documented manatee mortality events in 1963 and 1982 (Layne, 1965; O’Shea et al., 1991), and as a possible factor in bottlenose dolphin mortality in 1946–1947 and 1987–1988 (Gunter et al., 1948; Geraci et al., 1989).
Paralytic Shellfish Poisoning The toxin most commonly associated with paralytic shellfish poisoning (PSP) is saxitoxin; however, chemically related toxins have been identified, and include neosaxiton and gonyautoxins I to IV. PSP results from the ingestion of shellfish that have accumulated these potent neurotoxins as a result of consuming dinoflagellates, and is a significant public health concern (Fowler, 1993). For example, saxitoxin levels as high as 3500 µg/100 g (pooled tissue) have caused significant illness, resulting in the hospitalization of 500 individuals, and at least 20 cases of human mortality in recent years along the Pacific Coast of Mexico (Ochoa et al., 1998). PSP was hypothesized to affect sea otter distribution in southeast Alaska, where a primary prey species, the butter clam (Saxidomus giganteus), accumulates PSP toxin as a chemical defense mechanism without showing toxic effects (Kvitek et al., 1991; 1993). Sea otters will avoid highly contaminated prey (Kvitek et al., 1991), although it is likely that mortality of sea otters due to PSP has occurred in Alaska (De Gange and Vacca, 1989). Ingestion of mackerel containing saxitoxins has been implicated as a cause of death in humpback whales (Megaptera novaeangliae) (Geraci et al., 1989). The Cap Blanc (Western Sahara coast) subpopulation of Mediterranean monk seals (Monachus monachus) was highly impacted by a mortality event in 1997 (Forcada et al., 1999). There is a hypothesis that these deaths were due to a phycotoxin related to saxitoxin (Costas and Lopez-Rodas, 1998; Hernandez et al., 1998). Marine dinoflagellates producing saxitoxin and other toxins (neosaxitoxin, gonyautoxins) were spatially and temporally associated with this mortality, and included Alexandrium minutum, G. catenatum, and Dinophysis acuta (Hernandez et al., 1998). These findings were based on standard chemical techniques, including a mouse bioassay, and dinoflagellate culture (Hernandez et al., 1998). Examination of tissues from six seals revealed significant levels of saxitoxin in their livers and brains. Signs of lethargy, motor incoordination, and paralysis (animals floated horizontally and appeared incapable of voluntary movement) were reported in a few terminally ill animals (Hernandez et al., 1998). However, the minimal observations of clinical signs, and a lack of histopathology (due to the inability to observe many live animals and to obtain fresh samples) make it impossible to completely rule out other potential causes for this event. Osterhaus et al. (1997), for example, championed a competing hypothesis that a morbillivirus was responsible for the deaths, based on the fact that 7 of 17 animals possessed morbillivirus antibodies and 3 of 14 yielded a morbillivirus virus isolate. The absence of more thorough examinations, lack of data on normal indicators
0839_frame_C22.fm Page 495 Tuesday, May 22, 2001 11:51 AM
Toxicology
495
of monk seal health, and no preexposure information on antibody and toxin levels in Mediterranean monk seals made definitive diagnoses impossible (Harwood, 1998).
Domoic Acid Domoic acid (DA) is an excitotoxin produced by a number of marine diatoms (e.g., Pseudonitzschia pungens forma multiseries, P. australis), that targets the circumventricular regions of the brain and hippocampus, mimicking glutamate at the kainate receptor. A regulatory level of 20 µg/g of DA in edible meat has been established (Iverson and Truelove, 1994). Severe cases of seizures and other central nervous system problems, as well as hippocampal and amygdalal degeneration in humans, have been documented as caused by DA (Teitelbaum et al., 1990; Cendes et al., 1995). Degeneration of neurons in the hippocampal CA1/CA3 subregions and gliosis were documented in laboratory rats injected intraperitoneally with DA at a dose of 1.32 mg/kg (Sobotka et al., 1996). In 1987, more than 100 humans became ill from consuming mussels harvested near Prince Edward Island (Canada); many were severely affected, and four died (Teitelbaum et al., 1990). Brown pelicans (Pelecanus occidentalis) and Brandt’s cormorants (Phalacrocorax penicillatus) died in September 1991 near Santa Cruz, California, reportedly from consumption of DA-contaminated anchovies (Work et al., 1993). However, diagnoses were quite difficult because the only histological or clinical evidence available were hemorrhage and necrosis of skeletal muscle, and increases in serum blood urea nitrogen and creatanine phosphokinase. Because of all the confounding variables and the unpredictable nature of DA “outbreaks,” diagnoses are typically very difficult to establish definitively and many other factors must be ruled out in the process (Iverson and Truelove, 1994). However, a mortality event in 1998 involving California sea lions in the area of Monterey Bay, California during an algal bloom (Pseudonitzschia australis) was well investigated. In this event, 70 sea lions were clinically affected. They displayed neurological dysfunction (seizures, head weaving, ataxia, depression, and abnormal scratching) but were in good body condition; hematological and serum chemistry parameters were within the normal range (Scholin et al., 2000). DA was detected in algal samples, anchovies (Engraulis mordax), and sea lions, but not blue mussels (Mytilus edulus) used as indicator species at the time. The success of this investigation stemmed from an intensive multidisciplinary approach (Scholin et al., 1997; 2000; Van Dolah et al., 1997; Miller and Scholin, 1998; Gulland, 2000). Techniques employed ranged from satellite imagery for confirmation of bloom locations and characteristics to molecular DNA probes for specific algal identification. The DA receptor assay and confirmatory analytical chemistry (HPLC) were carried out in many matrices, and were supported by detailed observations of clinical signs, sample collection from subject animals, and histopathology describing very specific brain lesions in the 70 sea lions temporally and spatially associated with the algal bloom. Brain lesions associated with DA exposure included zonal vacuolation of the hippocampal neuropile involving several architectural strata, which was most severe in the ventral hippocampus (Scholin et al., 2000). These were critical findings linking DA exposure (DA in tissues, stomach contents, prey) and clinical signs. These “DAtype” lesions are well described for other species in association with DA toxicoses (Strain and Tasker, 1991; Dakshinamurti et al., 1993). The authors emphasize that this process of linking exposure, tissue level effects, and clinical signs is essential either to “proving” the role of a toxin or to ruling it out. However, this is rarely attempted for most strandings, monitoring studies, or population declines, where a contaminant has been implicated. A retrospective analysis indicates that this type of event may have occurred previously for northern fur seals and sea lions in 1978, 1986, 1988, and 1992 in California (Scholin et al., 2000), and for dolphins and sea lions in Mexico (Ochoa et al., 1998).
0839_frame_C22.fm Page 496 Tuesday, May 22, 2001 11:51 AM
496
CRC Handbook of Marine Mammal Medicine
Ciguatera Certain reef and inshore fish (possibly 400 species) of the tropics accumulate ciguatoxins and/or maitotoxin produced by the dinoflagellate Gambierdiscus toxicus (Fowler, 1993). Ciguatoxins are suspected of being responsible for deaths of about 50 endangered Hawaiian monk seals (Monachus schauinslandi) on Laysan Island in 1978, possibly via toxin-induced anorexia in combination with parasitism (Gilmartin et al., 1980; Banish and Gilmartin, 1992).
Oil Marine mammals are exposed to oil pollution in the form of crude and weathered oils that can directly contact the skin, pelage, eyes, mouth (including baleen), and nares. They also can inhale volatile petroleum fractions at the water surface during spills, ingest oil directly, and consume petroleum components in food. Crude oils are complex mixtures of thousands of organic and inorganic compounds, and vary widely in composition. Oil pollution is widespread, and potential impacts of high profile spills on marine mammals receive considerable attention. The history of such incidents, in relation to impacts on marine mammals, has been documented by St. Aubin (1990a) and Geraci and St. Aubin (1990), whereas Neff (1990a,b) has summarized characteristics, sources, environmental fate, and potential impacts of oil on these species. Constituents of oil include aliphatic compounds, and the aromatic and polycyclic aromatic hydrocarbons (AHs and PAHs). Alkanes are the predominant compounds and range from normal to branched forms, with 1 to 60 or more carbon atoms. Branched alkanes include pristane and phytane. Cycloalkanes (naphthenes) can also be abundant. Ring-based compounds can constitute about 20% of the hydrocarbons in crude oil. The benzene ring is the basic “building block,” with PAHs consisting of two to nine linked benzene rings. Lower molecular weight forms with one (benzene), two (naphthalene), or three (phenanthrene) rings predominate over those with a greater number of rings. AHs are the most toxic fraction of oil, with the lower molecular weight compounds showing the greatest acute toxicity in in vitro systems (Neff, 1990a). Higher aromatic compounds, although they are at lower concentrations in most petroleum compounds and have lower solubilities, are much less volatile and can cause significant effects in experimental studies. Contact exposure to oil impacts sea otters, fur seals, and polar bears most severely. This is because fouling of the pelage destroys insulative qualities of their fur, and reduces the capacity to maintain body temperature and buoyancy. Experimental application of crude oil to 20% of the pelage of captive sea otters nearly doubled metabolic rates and heat loss, and the increased grooming response spread the oil and caused it to penetrate more deeply (Davis et al., 1988). Rehabilitated otters monitored after experimental and natural exposures to oil require a few days to 2 weeks to regain normal pelage insulative characteristics after cleaning with dish detergent (Dawn™, Procter & Gamble, Inc.). Although weathered oil may not penetrate as deeply into the fur as fresh crude oil, it will likely also be forced more deeply by grooming (see summaries by Riedman and Estes, 1990). Much was learned about the impacts of oil on sea otters and other marine mammals from the Exxon Valdez oil spill in Prince William Sound, Alaska. Histopathological lesions in sea otters that died after exposure to oil from this event included interstitial pulmonary emphysema, centrilobular hepatic necrosis, and hepatic and renal lipidosis. The pathogenesis of the emphysema and hepatic necrosis were unclear, whereas the lipidosis may have been the result of increased energy demands relative to food intake (Lipscomb et al., 1993; 1994). Hematology and blood chemistry in oil-contaminated sea otters that ultimately died included leukopenia,
0839_frame_C22.fm Page 497 Tuesday, May 22, 2001 11:51 AM
Toxicology
497
lymphopenia, azotemia, hyperkalemia, hypoproteinemia, hypoalbuminemia, hypoglycemia, and leakage of hepatic transaminases (Lipscomb et al., 1994). Clinical symptoms included shock and hypothermia. Gastric erosions and hemorrhage were also associated with exposure to crude oil. These lesions were common in carcasses of sea otters that were never captured, as well as in animals that died when held for rehabilitation, thus do not appear to be simple artifacts of the stress of captivity (Lipscomb et al., 1994). DNA damage (using flow cytometry) was greater in sea otters from eastern Prince William Sound (Alaska) that were involved in the Exxon Valdez oil spill, as compared to otters from the western end of Prince William Sound (Gauthier et al., 1999b). Total aliphatic hydrocarbon concentrations (6100 to 7700 ng/g dry weight) detected in liver, kidney, and muscle of oiled sea otters from the Exxon Valdez incident were double those of sea otters from an area in southeastern Alaska that had not experienced an oil spill, whereas total AH concentrations (600 to 1500 ng/g dry weight) were three to nine times higher (Mulcahy and Ballachey, 1994). Mammals metabolize petroleum hydrocarbons rapidly. However, an absence of elevated concentrations in tissues is difficult to interpret. Aromatic compounds that were the most useful indicators of exposure to oil in sea otters that died in the Exxon Valdez spill were naphthalene, fluorene, phenanthrene, dibenzothiophene, chrysene, and most alkylated derivatives, except C1-naphthalene. Among the aliphatic compounds detected, the C18 to C20 alkanes showed little variability among tissues from any individual sea otter; concentrations of higher-carbon alkanes were higher than shorter-chain compounds, with the C25 and C26 alkanes being the highest (Mulcahy and Ballachey, 1994). Harbor seals from Prince William Sound had significantly higher concentrations of phenanthrene equivalents and naphthalene equivalents in bile than seals from other areas, and the ratios of naphthalene to phenanthrene equivalents were lower in seals with greatest exposure (Frost et al., 1994). Harbor seals and other marine mammals probably metabolize hydrocarbons rapidly and efficiently, mediated through the induction of mixed-function oxidases. Concentrations of PAHs in most tissues (muscle, liver, brain, kidney, heart, and lung) of harbor seals exposed to the Exxon Valdez spill were near or below detection limits. However, oiled harbor seals had higher concentrations of PAHs in blubber than various reference samples, mostly compounds of low molecular weight (two- to three-ring aromatics, naphthalenes, and phenanthrenes). PAHs in milk and mammary tissue were variable, but the highest PAH concentration (1142 ng/g) detected in any sample of any harbor seal was in milk from the stomach of a pup with an oiled mother. Among aliphatic compounds, phytane was relatively high in brain of oiled seals (1228 to 7839 ppb wet weight) (Frost et al., 1994). However, it is thought that inhalation of volatile, short-chain AHs had the greatest impact on harbor seals, with levels immediately after the spill speculated to be sufficient to cause respiratory or cardiac arrest, or to interfere with breathing (Frost et al., 1994). Visibly oiled seals collected several weeks after the Exxon Valdez spill had mild acanthosis and orthokeratotic hyperkeratosis of the epidermis and mild, reversible hepatocellular necrosis and swelling with mild bile inspissation within canaliculi (Spraker et al., 1994). The most significant lesions, however, were intramyelinic edema of the large myelinated axons of the midbrain, neuronal swelling, neuronal necrosis, and axonal swelling and degeneration. These lesions were most severe in the thalamic nuclei, and were consistent with nervous system damage caused by highly volatile hydrocarbons (Spraker et al., 1994). Variability among concentrations of hydrocarbons in tissues and presence of nervous system lesions in these studies was probably related to variability in exposure histories of individual animals (Frost et al., 1994). No firm evidence of contamination of tissues or toxicological effects of the Exxon Valdez spill was obtained from examination of cetaceans (Loughlin, 1994).
0839_frame_C22.fm Page 498 Tuesday, May 22, 2001 11:51 AM
498
CRC Handbook of Marine Mammal Medicine
Limited laboratory and captive studies have been carried out on effects of oil on marine mammals (St. Aubin et al., 1985; Geraci and St. Aubin, 1990; St. Aubin, 1990a,b). Bottlenose dolphins’ tactile sense played a role in sensing and avoiding oil. Three captive polar bears were induced to swim through a pool covered with a slick of crude oil. Ingestion of oil during intensive grooming the first few days after fouling resulted in vomiting, diarrhea, and biochemical lesions indicative of liver and kidney damage. Three ringed seals died acutely after exposure to crude oil in a tank, but these deaths were also complicated by stress (Geraci and Smith, 1976). In other instances, ringed seals immersed in an experimental oil slick for 24 hours had transient eye irritation (profuse lacrimation, severe conjunctivitis, and corneal abrasions and ulcers that disappeared after being returned to clear water), detectable hydrocarbons in tissues, but few consistent patterns in hematology and biochemistry, other than elevated serum liver enzymes, and lesions in kidney and liver (Geraci and Smith, 1976; St. Aubin, 1990a). Epidermal acanthosis and hyperkeratosis, excessive lacrimation, conjunctivitis, and corneal abrasions and ulcers occur in seals after contact with oil (Geraci and Smith, 1976; St. Aubin, 1990a; Spraker et al., 1994). Harp seal pups experimentally coated with oil over the entire body on 2 consecutive days showed no pathological changes and maintained core body temperatures over the ensuing 4 days of the trials (Geraci and Smith, 1976). A small number of oral-dosing studies were carried out on marine mammals in the 1970s and 1980s. Ringed seals orally dosed with 5 ml crude oil daily for up to 5 days cleared residues from tissues and blood after 7 days through excretion in bile and urine, and no major histopathological or biochemical lesions were observed. Similar lack of effects was noted in harp seal pups with a single 75-ml dose of crude oil. Ringed seals were also dosed with crude oil with isotopically labeled AHs; the compounds were excreted in urine and declined within 2 weeks. Induction of aryl hydrocarbon hydroxylase was noted, but other enzyme activity did not indicate any biochemical lesions. No clinical, hematological, or biochemical effects were noted in a captive bottlenose dolphin dosed daily with 5 ml of machine oil for 99 days. These studies suggest that captive marine mammals can tolerate small amounts of ingested oil (Geraci and St. Aubin, 1990; St. Aubin, 1990b). However, results of these captive studies must be extrapolated to wild animals cautiously. Can these animals detect, avoid, and remove themselves from a spill area before achieving a significant exposure that results in adverse effects? In cetaceans, penetration of oil into the skin is impeded by tight intercellular bridges and the unusual thickness of the epidermis (which is 10 to 20 times thicker than that of humans). Direct application of various petroleum fractions to dolphin skin resulted only in subtle histological changes, which were reversed within a week after exposure (Geraci and St. Aubin, 1990). Oil fouling of baleen plates does not appear to have serious lasting structural or functional effects (Geraci and St. Aubin, 1990). Structural and chemical integrity of isolated baleen plates of seven species of whales were reported to remain intact when they were soaked in crude oil, gasoline, or tar over long periods. When baleen plates were exposed to oil in continuous-flow flumes, minor decreases in filtration rates due to fouling were observed, with variation in impairment based on the type of oil (Geraci and St. Aubin, 1990). There have been relatively few surveys for petroleum constituents in marine mammals. Low concentrations of PAHs (0.1 to 0.6 ppm wet weight chrysene equivalents) were detected in muscles of 26 harbor porpoises from the United Kingdom (Law and Whinnet, 1992). Low concentrations of two- to four-ring compounds, but not higher-weight PAHs, were detected in blubber of seven sperm whales stranded in the southern North Sea (Holsbeek et al., 1999). Low concentrations of PAHs (0.1 to 1.2 ppm dry weight chrysene equivalents) with a preponderance of low molecular weight compounds were also reported in small numbers of muscle samples from five species of cetaceans and four species of seals from the northwest Atlantic
0839_frame_C22.fm Page 499 Tuesday, May 22, 2001 11:51 AM
Toxicology
499
(Hellou et al., 1990). Skin and blubber biopsies of southern sea lions from the polluted Mar del Plata in Argentina had elevated concentrations (2785 ng/gm dry weight) of total PAHs in comparison with individuals from a less polluted area (578 ng/g). Predominant compounds in biopsies and blood were phenanthrene, naphthalene, and acenaphthene (Marsili and Focardi, 1997). Low levels (177 to 1821 ng/g wet weight) of total PAHs were also found in four South American fur seals (Arctocephalus australis) and a sub-Antarctic fur seal (Arctocephalus tropicalis) by Fossi and Marsili (1997). Varanasi et al. (1994) found large variation in PAH concentrations in stomach contents of gray whales (7 to 2100 ppb). It has been speculated that PAHs induced tumors in belugas of the St. Lawrence River estuary through the formation of DNA adducts (Martineau et al., 1988). However, DNA adducts occur at similar levels in livers of belugas from remote locations without significant PAH contamination (Ray et al., 1992). There are no published studies on PAHs or AHs in sirenians, and no reports conclusively linking harm to dugongs or manatees from oil exposure. Such exposure is feasible, however, in industrial areas. Dugongs were found dead in the Persian Gulf coincident with major oil spills from well platforms damaged during warfare, but carcasses were not examined (Preen, 1988).
Treatment and Diagnostic Procedures Dose Scaling The broad range of marine mammal body sizes (sea otters to blue whales) makes it important to consider toxin scaling (toxin exposure or level relative to response). For a discussion of drug dose scaling, see Chapter 31, Pharmaceuticals. Schmidt-Nielsen (1972) accounts for scaling of dosages and exposure and uses a tragic example in an elephant. Investigators hoped to induce “musth” behavior in an elephant by administering lysergic acid diethylamide (LSD) and simply scaled from the domestic cat dosage of 0.26 mg (0.1 mg/kg) by multiplying by the body weight factor (2970 kg), so used 297 mg in the elephant. The elephant died acutely. This points at an unresolved issue of interpretation of contaminant exposures in marine mammals: how should one scale exposures and toxin effects from such diverse mammals? For example, using some proposed techniques for calculating dosages, the range of LSD that might be administered to the elephant to achieve a desired behavioral effect varies from 0.4 to 297 mg (Schmidt-Nielsen, 1972), and the same level of uncertainty associated with toxicoses exists. How is one to interpret exposure or levels in tissues in the widely varying marine mammals? When assessing a toxin exposure in an animal without prior information on pharmacokinetics, dose–response relationship, and effects, it may be impossible to predict the outcome with any certainty. The differences in absorption, metabolism, nutritional condition, blood circulation, and body size will vary the response to an agent from no effect to acute toxicosis. Some of the peculiar aspects of marine mammals that need to be considered are diving, thermoregulation, hypoxia-reperfusion (oxidative stress-ameliorative mechanisms), a dynamic cardiovascular system, and associated physiological and metabolic mechanisms that will differ from better studied terrestrial mammals. It is important to reiterate that much of the toxin research has assessed solely chemical levels in tissues, and very little is known about response or adverse effects.
Treatment If treated early in the course of exposure/absorption, the removal of gastric and intestinal contents (induced emesis, lavage, activated charcoal) could help. However, caution is
0839_frame_C22.fm Page 500 Tuesday, May 22, 2001 11:51 AM
500
CRC Handbook of Marine Mammal Medicine
required because some of these techniques can result in pulmonary aspiration of gastric contents in weak or neurologically impaired animals, and the stress of treatment may aggravate clinical signs. Treatment of toxicoses can be fairly basic and requires only supportive care, or can be futile when irreversible lethal damage has occurred. During large mortality events involving toxic substances (e.g., algal blooms, oil spills), experienced marine mammal health personnel will need to triage and select those patients with the highest likelihood of recovery. This is the first step after securing the animal for examination. The assessment of the animal will involve identifying the most critical organ system(s) affected and possible decontamination. The animal must be able to tolerate the decontamination procedures and thus initial fluid therapy, respiratory support, and sedation may be required prior to the handling associated with decontamination. Immediately decontaminating a stressed and compromised animal may lead to rapid deterioration of its condition, and even death. For the critical care of the intoxicated animal, please refer to Chapter 30, Intensive Care, and Chapter 31, Pharmaceuticals, and Osweiller et al. (1985), Fowler (1993), Williams et al. (1994), Lorgue et al. (1996), or contact the groups listed in Table 2 (Plumb, 1999). Only rarely can an antidote that quickly and effectively reverses the toxicoses be given (e.g., atropine for organophosphorus compounds). Usually, only supportive care is indicated. Handling and treatment of contaminated animals can be harmful or nonproductive. For example, sea otters cleaned following crude oil contamination have difficulty thermoregulating (Davis et al., 1988; Williams et al., 1988). Contamination of sea otter fur with oil or dispersants reduces insulation. Washing with Dawn dishwashing detergent is the most effective treatment for removing crude oil, but also removes natural lipids (squalene) from the fur, although some insulative qualities are regained with washing (Williams et al., 1988). Other solvents, such as Shelsol, 70-viscosity mineral oil, allowed for further penetration of oil into the pelt and TABLE 2 Contact Information for the Intoxicated and Contaminated Animal Program Product Failure and Adverse Reaction Reporting
Targeted Concerns Drugs, devices, animal foods Biologics (including vaccines, bacterins, and diagnostic kits)
Food Animal Residue Avoidance Databank (FARAD)
National Animal Poison Information Center (NAPIC in U.S.) National Pesticide Telecommunications Network USP Veterinary Practitioners’ Reporting Program
Poisonings, toxicology, environmental chemistry, etc. Medical product adverse reactions, quality concerns, etc.
Source: Based on Plumb (1999).
Contact Information Food and Drug Administration (U.S.) (301) 443-4095 (collect) U.S. Department of Agriculture (515) 232-5789 Internet e-mail:
[email protected] or
[email protected] Web site: http://www.farad.org 1(900)680-0000 ($30 per case) or 1(800) 548-2423 or 1(888)426-4435 ($30 per case); credit cards only 1(800)858-7377 or 7378
1(800) 4-USP-PRN (1(800) 487-7776) http://www.usp.org
0839_frame_C22.fm Page 501 Tuesday, May 22, 2001 11:51 AM
Toxicology
501
prolonged cleansing (Williams et al., 1988). Rinsing, proper rehabilitation of the fur coat, and sufficient caloric intake for metabolic maintenance (rates double for animals with a 20% oiled surface) are critical for the recovery of fur insulative qualities and thermoregulation, and can take 3 to 6 days (Davis et al., 1988). Once the initial crisis is over and the animal has physiologically stabilized and/or been decontaminated, a more detailed assessment of the animal is needed (follow-up physical examination, serum chemistries, hematology). Monitoring for potential development of secondary problems such as infections or organ failure is important before recovery can be concluded. Treatment of toxicoses in wild animals can present significant challenges to personnel, housing and facilities, finances, and rehabilitation protocols, and can also create ethical dilemmas. Treatment of captive animals is more likely to be successful, as therapy can be instituted more immediately and intensively, and possibly with reduced stress. Treating animals for acute toxicoses is often complicated by the fact that a reliable diagnosis may take many days, if one is achieved at all, so that specific antidotal therapies cannot be implemented in a timely manner.
Diagnosis In some cases, response to therapy indicates the causative agent, as the effects of a toxin are reversed (e.g., atropine and 2-PAM for organophosphorus poisoning). In other cases, the toxin is excreted and signs abate. The collection of clinical data and archiving samples before the toxin is excreted may be critical for identifying a cause in legal cases involving accidents, liability, or malicious conduct. From a management perspective, a reliable diagnosis can drive the needed changes required if an anthropogenic role is involved. For chronic, low-level population changes tentatively implicating environmental contaminants, the investigation can be difficult and the diagnosis elusive. The investigation will likely become a research project, but the role of diagnostics cannot be diminished because controlled experiments are not feasible in most free-ranging marine animals. The effects may be subtle and may involve reproductive or immune status in ways that are currently unknown or unmeasureable. Effects that may be subtle in the individual animal may have significant implications for the population. During gross examinations and sampling, the diagnosis should not be presumed. If it is, the goal becomes to sample for the diagnosis, instead of the full assessment of the condition and health of the animal, and a reasonable consideration of all the possible etiologies. Gross examination of carcasses can be underemphasized in suspected toxic cases; emphasis can instead be inappropriately placed on examination for biomarkers of exposure. This is a very narrow vision. Toxinspecific biomarkers need to be evaluated in light of a general health assessment. Similarly, the population biologist may indicate a population is healthy based on census or count data, while underlying stresses may be operating that can result in future or region-specific declines that are not detected until they are well under way. The paucity of research efforts to describe the health of marine mammals has significantly handicapped diagnoses or investigations of potential toxicoses. Baseline health and toxicological data are essential for understanding the role of toxic substances during population declines and mortality events. Chemical residues, biochemical markers, and histological samples require appropriate collection and, once collected, no laboratory can improve the sample’s quality (see Chapter 21, Necropsy). Histopathology can indicate the response of the animal and pathogenesis of the observed effects. Without it, one will struggle to defend an association of high toxin levels and the presence of dead animals. Biochemical assays (enzyme inhibition, induction of some enzymes) can indicate the presence of biochemical lesions that can possibly link the presence of an agent with the
0839_frame_C22.fm Page 502 Tuesday, May 22, 2001 11:51 AM
502
CRC Handbook of Marine Mammal Medicine
observed effect and exposure, and increases diagnostic power when combined with histopathology, gross examination, and/or observations of clinical signs. Biochemical or in vitro assays suffer from an inability to indicate the severity of the effect and relate this to the impact on the individual animal, or the population. For some biochemical indicators, the response would be considered normal and, thus, is really not an adverse effect. Normal or expected response vs. adverse effect is commonly confused for some biomarkers. For example, the induction of the cytochrome P450 system would be considered normal in the presence of some organochlorines, but in itself is simply a response and may not indicate an adverse effect. Simple induction of this enzyme cannot be linked with whole-animal or population-level effects, only to an exposure to a particular toxin and induction of a metabolic process to clear the body of it. Analogies for biomarkers of the immune, endocrine, and other systems can be made as well. The authors encourage evaluation of the health of the species, with criteria for such assessments developed so that these biomarkers can be used to assess both individual and population adverse health effects. One must determine “Hysteria vs. Association vs. Cause-Effect” with careful and objective interpretation, and understand that the presence of contamination is not evidence of an effect. If understanding of these effects in the health assessments of marine mammal is not improved, hysterical associations will continue to be made.
Acknowledgments The authors thank Greg Bossart, Derek Muir, and Mike Ziccardi for reviewing this chapter, and Debbie Fauquier for editorial assistance.
References Addison, R.F., 1989, Organochlorines and marine mammal reproduction, Can. J. Fish. Aquat. Sci., 46: 360–368. Addison, R.F., and Brodie, P.F., 1984, Characterization of ethoxyresorufin-O-deethylase in grey seal Halichoerus grypus, Comp. Biochem. Physiol. C., 79: 261–263. Addison, R.F., Brodie, P.F., Edwards, A., and Sadler, M.C., 1986, Mixed-function oxidase activity in the harbor seal (Phoca vitulina) from Sable Is., N.S., Comp. Biochem. Physiol., 85, C: 121–124. Addison, R.F., Renton, K.W., Edwards, A.J., and Brodie, P.F., 1988, Polychlorinated biphenyls and mixedfunction oxidase in grey seals (Halichoerus grypus), Mar. Environ. Res., 24: 111–112. Aguilar, A., 1985, Compartmentation and reliability of sampling procedures in organochlorine pollution surveys of cetaceans, Residue Rev., 95: 91–114. Aguilar, A., 1987, Using organochlorine pollutants to discriminate marine mammal populations: A review and critique of the methods, Mar. Mammal Sci., 3: 242–262. Aguilar, A., and Borrell, A., 1994a, Abnormally high polychlorinated biphenyl levels in striped dolphins (Stenella coeruleoalba) affected by the 1990–1992 Mediterranean epizootic, Sci. Total Environ., 154: 237–247. Aguilar, A., and Borrell, A., 1994b, Reproductive transfer and variation of body load of organochlorine pollutants with age in fin whales (Balaenoptera physalus), Arch. Environ. Contam. Toxicol., 27: 546–554. Aguilar, A., and Borrell, A., 1994c, Assessment of organochlorine pollutants in cetaceans by means of skin and hypodermic biopsies, in Non-destructive Biomarkers in Vertebrates, Fossi, M.C., and Leonzio, C. (Eds.), Lewis Publishers, Boca Raton, FL, 245–267. Aguilar, A., Borrell, A., and Pastor, T., 1999, Biological factors affecting variability of persistent pollutant levels in cetaceans, J. Cetacean Res. Manage., 1: 83–116.
0839_frame_C22.fm Page 503 Tuesday, May 22, 2001 11:51 AM
Toxicology
503
Alexander, V., 1995, The influence of the structure and function of the marine food web on the dynamics of contaminants in Arctic Ocean ecosystems, Sci. Total Environ., 160/161: 593–603. Allen, J.R., and Barsotti, D.A., 1976, The effects of transplacental and mammary movement of PCBs on infant rhesus monkeys, Toxicology, 6: 331–340. Allen, J.R., Carstens, L.A., and Barsotti, D.A., 1974, Residual effects of short-term, low-level exposure of nonhuman primates to polychlorinated biphenyls, Toxicol. Appl. Pharmacol., 30: 440–451. Allen, J.R., Barsotti, D.A., and Carstens, L.A., 1980, Residual effects of polychlorinated biphenyls on adult nonhuman primates and their offspring, J. Toxicol. Environ. Health, 6: 55–66. Ames, A.L., and Van Vleet, E.S., 1996, Organochlorine residues in the Florida manatee, Trichechus manatus latirostris, Mar. Pollut. Bull., 32: 374–377. Anas, R.E., 1974, Heavy metals in the northern fur seal, Callorhinus ursinus, and harbor seals, Phoca vitulina, Fish. Bull., 72: 133–137. André, J.M., Boudo, A., and Ribeyre, F., 1991, Mercury accumulation in Delphinidae, Water Air Soil Pollut., 56: 187–201. Aono, S., Tanabe, S., Fujise, Y., Kato, H., and Tatsukawa, R., 1997, Persistent organochlorines in minke whale (Balaenoptera acutorostrata) and their prey species from the Antarctic and the North Pacific, Environ. Pollut., 98: 81–89. Armstrong, H.E.L., Corns, W.T., Stockwell, P.B., O’Conner, G., Ebdon, L., and Evans, E.H., 1999, Comparison of AFS and ICP-MS detection coupled with gas chromatography for the determination of methyl mercury in marine samples. Anal. Chim. Acta, 390: 245–253. Aulerich, R.J., and Ringer, R.K., 1977, Current status of PCB toxicity to mink, and effect on their reproduction, Arch. Environ. Contam. Toxicol., 6: 279–292. Aulerich, R.J., Bursian, S.J., Breslin, W.J., Olson, B.A., and Ringer, R.K., 1985, Toxicological manifestations of 2,4,5,2′,4′,5′-, 2,3,6,2′,3′,6′-, and 3,4,5,3′,4′,5′-hexachlorobiphenyl and Aroclor 1254 in mink, J. Toxicol. Environ. Health, 15: 63–79. Bacon, C.E., Jarman, W.M., Estes, J.A., Simon, M., and Norstrom, R.J., 1999, Comparison of organochlorine contaminants among sea otter (Enhydra lutris) populations in California and Alaska, Environ. Toxicol. Chem., 18: 452–458. Baden, D.G., Fleming, L.E., and Bean, J.A., 1995, Marine toxins, in Handbook of Clinical Neurology, Vol. 21 (65) Intoxications of the Nervous System Part II, de Wolff, F.A. (Ed.), Elsevier Sciences B.V., Amsterdam, the Netherlands, 141–175. Baird, R.W., Langelier, K.M., and Stacey, P.J., 1989, First records of false killer whales, Pseudorca crassidens, in Canada, Can. Field Nat., 103: 368–371. Baker, J.R., 1989, Pollution-associated uterine lesions in grey seals from the Liverpool Bay area of the Irish Sea, Vet. Rec., 125: 303. Banish, L.D., and Gilmartin, W.G., 1992, Pathological findings in the Hawaiian monk seal, J. Wild. Dis., 28: 428–434. Barsotti, D.A., Marlar, R.J., and Allen, J.R., 1976, Reproductive dysfunction in rhesus monkeys exposed to low levels of polychlorinated biphenyls (Aroclor 1248), Food Cosmet. Toxicol., 14: 99–103. Becker, P.R., Mackey, E.A., Demiralp, R., Suydam, R., Early, G., Koster, B., and Wise, S.A., 1995, Relationship of silver with selenium and mercury in the liver of two species of toothed whales (Odonotocetes), Mar. Pollut. Bull., 30: 262–271. Beckmen, K.B., 1999, Blood Organochlorine, Immune Function, and Health of Free Ranging Northern Fur Seal Pups (Callorhinus ursinus), Ph.D. thesis, Institute of Arctic Biology, University of Alaska, Fairbanks. Beckmen, K.B., Lowenstine, L.J., Newman, J., Hill, J., Hanni, K., and Gerber, J., 1997, Clinical and pathological characterization of northern elephant seal skin disease, J. Wild. Dis., 33: 438–449. Beckmen, K.B., Ylitalo, G.M., Towell, R.G., Krahn, M.M., O’Hara, T.M., and Blake, J.E., 1999, Factors affecting organochlorine contaminant concentrations in milk and blood of northern fur seal (Callorhinus ursinus) dams and pups from St. George Island, Alaska, Sci. Total Environ., 231: 183–200.
0839_frame_C22.fm Page 504 Tuesday, May 22, 2001 11:51 AM
504
CRC Handbook of Marine Mammal Medicine
Béland, P., De Guise, S., and Plante, R., 1991, Toxicology and Pathology of St. Lawrence Marine Mammals, Final Report, Wildlife Toxicology Fund, St. Lawrence National Institute of Ecotoxicology, World Wildlife Fund, Toronto, 95. Béland, P., De Guise, S., Girard, C., Lagacé, A., Martineau, D., Michaud, R., Muir, D.C.G., Norstrom, R.J., Pelletier, E., Ray, S., and Shugart, L.R., 1993, Toxic compounds and health and reproductive effects in St. Lawrence beluga whales, J. Great Lakes Res., 19: 766–775. Bergek, S., Bergqvist, P.-A., Hjelt, M., Olsson, M., Rappe, C., Roos, A., and Zook, D., 1992, Concentrations of PCDDs and PCDFs in seals from Swedish waters, Ambio, 21: 553–556. Bergman, Å., and Olsson, M., 1985, Pathology of Baltic grey seal and ringed seal females with special reference to adrenocortical hyperplasia: Is environmental pollution the cause of a widely distributed disease syndrome? Finn. Game Res., 44: 47–62. Bergman, Å., Olsson, M., and Reiland, S., 1992, Skull-bone lesions in the Baltic grey seal (Halichoerus grypus), Ambio, 21: 517–519. Bergman, Å., Norstrom, R.J., Haraguchi, K., Kuroki, H., and Béland, P., 1994, PCB and DDE methyl sulfones in mammals from Canada and Sweden, Environ. Toxicol. Chem., 13: 121–128. Bernhoft, A., Skaare, J.U., Wiig, O., Derocher, A.E., and Larsen, H.J., 2000, Possible immunotoxic effects of organochlorines in polar bears at Svalbard, J. Toxicol. Environ. Health A, 59: 101–114. Bernt, K.E., Hammill, M.O., Lebeuf, M., and Kovacs, K.M., 1999, Levels and patterns of PCBs and OC pesticides in harbour and grey seals from the St. Lawrence estuary, Canada, Sci. Total Environ., 243/244: 243–262. Betti, C., and Nigro, M., 1996, The Comet assay for the evaluation of the genetic hazard of pollutants in cetaceans: Preliminary results on the genotoxic effects of methylmercury on the bottle-nosed dolphin lymphocytes in vitro, Mar. Pollut. Bull., 32: 545–548. Bleavins, M.R., Aulerich, R.J., and Ringer, R.K., 1980, Polychlorinated biphenyls (Aroclors 1016 and 1242): Effects on survival and reproduction in mink and ferrets, Arch. Environ. Contam. Toxicol., 9: 627–635. Bleavins, M.R., Aulerich, R.J., Ringer, R.K., and Bell, T.G., 1982, Excessive nail growth in the European ferret induced by Aroclor 1242, Arch. Environ. Contam. Toxicol., 11: 305–312. Blomkvist, G., Roos, A., Jensen, S., Bignert, A., and Olsson, M., 1992, Concentrations of sDDT and PCB in seals from Swedish and Scottish waters, Ambio, 21: 539–545. Boon, J.P., Van Arnhem, E., Jansen, S., Kannan, N., Petrick, G., Schulz, D., Duinker, J.C., Reijnders, P.J.H., and Goksøyr, A., 1992, The toxicokinetics of PCBs in marine mammals with special reference to possible interactions of individual congeners with the cytochrome P450-dependent monooxygenase system: An overview, in Persistent Pollutants in Marine Ecosystems, Walker, C.H., and Livingstone, D.R. (Eds.), Pergamon Press, Oxford, U.K., 119–159. Boon, J.P., Oostingh, I., van der Meer, J., and Hillebrand, M.T.J., 1994, A model for the bioaccumulation of chlorobiphenyl congeners in marine mammals, Eur. J. Pharmacol. Environ. Toxicol. Pharmacol., 270(2–3): 237–251. Boon J.P., van der Meer, J., Allchin, C.R., Law, R.J., Klungsoyr, J., Leonards, P.E.G., Spliid, H., StorrHansen, E., McKenzie, C., and Wells, D.E., 1997, Concentration-dependent changes of PCB patterns in fish-eating mammals: Structural evidence for induction of cytochrome P450, Arch. Environ. Contam. Toxicol., 33: 298–311. Boon, J.P., Sleiderink, H.M., Helle, M.S., Dekker, M., van Schanke, A., Roex, E., Hillebrand, M.T.J., Klamer, H.J.C., Govers, B., Pastor, D., Morse, D., Wester, P.G., and de Boer, J., 1998, The use of a microsomal in vitro assay to study phase I biotransformation of chlorobornanes (toxaphene) in marine mammals and birds, Comp. Biochem. Physiol. C, 121: 385–403. Borrell, A., and Aguilar, A., 1990, Loss of organochlorine compounds in the tissues of a decomposing stranded dolphin, Bull. Environ. Contam. Toxicol., 45: L46–53. Borrell, A., and Reijnders, P.J.H., 1999, Summary of temporal trends in pollutant levels observed in marine mammals, J. Cetacean Res. Manage., 1: 149–155.
0839_frame_C22.fm Page 505 Tuesday, May 22, 2001 11:51 AM
Toxicology
505
Borrell, A., Bloch, D., and Desportes, G., 1995, Age trends and reproductive transfer of organochlorine compounds in long-finned pilot whales from the Faroe Islands, Environ. Pollut., 88: 283–292. Borrell, A., Aguilar, A., Corsolini, S., and Focardi, S., 1996, Evaluation of toxicity and sex-related variation of PCB levels in Mediterranean striped dolphins affected by an epizootic, Chemosphere, 32: 2359–2369. 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 immmunohistochemical features, Toxicol. Pathol., 26: 276–282. Bratton, G.R., Spainhour, C.B., Flory, W., Reed, M., and Jayko, K., 1993, Presence and potential effects of contaminants, in The Bowhead Whale, Special Publ. 2, Burns, J.J. Montague, J.J., and Cowles, C.J. (Eds.), Society for Marine Mammalogy, Lawrence, KS, 701–744. Bratton, G.R., Flory, W., Spainhour, C.B., and Haubold, E.M., 1997, Assessment of selected heavy metals in liver, kidney, muscle, blubber, and visceral fat of Eskimo harvested bowhead whales Balaena mysticetus from Alaska’s north coast, Final Report submitted to the North Slope Borough Department of Wildlife Management, Barrow, AK, 233 pp. Brouwer, A., 1999, Induction of biotransformation enzymes by polyhalogenated aromatic hydrocarbons (PHAHs): Potential impact on animal physiology and health, J. Cetacean Res. Manage., 1: 209–221. Brouwer, A., Reijnders, P.H.J., and Koeman, J.H., 1989, Polychlorinated biphenyl (PCB)-contaminated fish induces vitamin A and thyroid hormone deficiency in the common seal (Phoca vitulina), Aquat. Toxicol., 15: 99–106. Buckland, S.J., Hannah, D.J., Taucher, J.A., Slooten, E., and Dawson, S., 1990, Polychlorinated dibenzop-dioxins and dibenzofurans in New Zealand’s Hector’s dolphin, Chemosphere, 20: 1035–1042. Burns, L.A., Meade, B.J., and Munson, A.E., 1996, Toxic responses of the immune system, in Casarett and Doull’s Toxicology: The Basic Science of Poisons, 5th ed., Klaasen, C.D. (Ed.), McGraw-Hill, Health Professions Division, New York, 355–402. Busbee, D., Tizard, I., Stott, J., and Ferrick, D., 1999, Environmental pollutants and marine mammal health: The potential impact of hydrocarbons and halogenated hydrocarbons on immune system dysfunction, J. Cetacean Res. Manage., 1: 223–248. Casarett, L.J., and Bruce, M.C., 1980, Origin and scope of toxicology, in Casarett and Doull’s Toxicology; The Basic Science of Poisons, 2nd ed., Doull, J., Klaassen, C.D., and Amdur, M.O. (Eds.), Macmillan, New York, 12. Caurant, F., and Amard-Triquet, C., 1995, Cadmium contamination in pilot whales, Globicephala melas: Source and potential hazard to the species, Mar. Pollut. Bull., 30: 207–210. Cendes, F., Andermann, F., Carpenter, S., Zatorre, R.J., and Cashmann, N.R., 1995, Temporal lobe epilepsy caused by domoic acid intoxication: Evidence for glutamate receptor-mediated excitotoxicity in humans, Ann. Neurol., 37: 123–126. Cleeman, M., Riget, F., Paulsen, G.B., de Boer, J., and Dietz, R., 2000, Organochlorines in Greenland ringed seals (Phoca hispida), Sci. Total Environ., 245: 103–116. Cockcroft, V.G., De Kock, A.C., Lord, D.A., and Ross, G.J.B., 1989, Organochlorines in bottlenose dolphins, Tursiops truncatus, from the east coast of South Africa, S. Afr. J. Mar. Sci., 8: 207–217. Colborn, T., and Smolen, M.J., 1996, Epidemiological analysis of persistent organochlorine contaminants in cetaceans, Rev. Environ. Contam. Toxicol., 146: 92–172. Costas, E., and Lopez-Rodas, V., 1998, Paralytic phycotoxins in monk seal mass mortality, Vet. Rec., 142: 643–644. Cuvin-Aralar, M.L.A., and Furness, R.W., 1991, Mercury and selenium interaction: A review, Ecotoxicol. Environ. Saf., 21: 348–364. Dakshinamurti, K., Sharma, S.K., Sundaram, M., and Wantanabe, T., 1993, Hippocampal changes in developing postnatal mice following intrauterine exposure to domoic acid, J. Neurosci., 13: 4486–4495.
0839_frame_C22.fm Page 506 Tuesday, May 22, 2001 11:51 AM
506
CRC Handbook of Marine Mammal Medicine
Davis, R.W., Williams, T.M., Thomas, J.A., Kastelein, R.A., and Cornell, L.H., 1988, The effects of oil contamination and cleaning on sea otters II, metabolism, thermoregulation, and behavior, Can. J. Zool., 66: 2782–2790. Dearth, M.A., and Hites, R.A., 1991, Complete analysis of technical chlordane using negative ionization mass spectrometry, Environ. Sci. Technol., 25: 245–254. de Boer, J., and Wester, P.G., 1993, Determination of toxaphene in human milk from Nicaragua and in fish and marine mammals from the northeastern Atlantic and the North Sea, Chemosphere, 27: 1879–1890. de Boer, J., Wester, P.G., Evers, E.H.G., and Brinkman, U.A.T., 1996, Determination of tris(4-chlorophenyl) methanol and tris(4-chlorophenyl) methane in fish, marine mammals, and sediment, Environ. Pollut., 93: 29–47. De Gange, A.R., and Vacca, M.M., 1989, Sea otter mortality at Kodiak Island, Alaska, during summer 1987, J. Mammal., 70: 836–838. De Guise, S., Lagace A., and Beland, P., 1994, True hermaphroditism in a St. Lawrence beluga whale, J. Wildl. Dis., 30: 287–290. De Guise, S., Bernier, J., Martineau, D., Beland, P., and Fournier, M., 1996, Effects of in vitro exposure of beluga whale splenocytes and thymocytes to heavy metals, Environ. Toxicol. Chem., 15: 1357–1364. De Guise, S., Martineau, D., Béland, P., and Fournier, M., 1998, Effects of in vitro exposure of beluga whale leukocytes to selected organochlorines, J. Toxicol. Environ. Health A, 55: 479–493. de Kock, A.C., Best, P.B., Cockcroft, V., and Bosma, C., 1994, Persistent organochlorine residues in small cetaceans from the east and west coasts of southern Africa, Sci. Total Environ., 154: 153–162. DeLong, R.L., Gilmartin, W.G., and Simpson, J.G., 1973, Premature births in California sea lions: Association with high organochlorine pollutant residue levels, Science, 181: 1168–1170. Denton, G.R.W., Marsh, H., Heinsohn, G.E., and Burdon-Jones, C., 1980, The unusual metal status of the dugong Dugong dugon, Mar. Biol., 57: 201–219. de Swart, R.L., Kluten, R.M.G., Huizing, C.J., Vedder, E.J., Reijnders, P.J.H. Visser, I.K.G., Uyt de Haag, F.G.C.M., and Osterhaus, A.D.M.E., 1993, Mitogen and antigen induced B cell and T cell responses of peripheral blood mononuclear cells from the harbor seal (Phoca vitulina), Vet. Immunol. Immunopath., 37: 217–230. de Swart, R.L., Ross, P.S., Vedder, E.J., Timmerman, H.H., Heisterkamp, S.H., van Loveren, H., Vos, J.G., Reijnders, P.J.H., and Osterhaus, A.D.M.E., 1994, Impairment of immunological functions in harbor seals (Phoca vitulina) feeding on fish from polluted coastal 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 feeding study, Environ. Health Perspect., 104(Suppl. 4): 823–828. Dietz, R., Nielson, C.O., Hansen, M.M., and Hansen, C.T., 1990, Organic mercury in Greenland birds and mammals, Sci. Total Environ., 95: 41–51. Dietz, R., Nørgaard, J., and Hansen, J.C., 1998, Have Arctic marine mammals adapted to high cadmium levels? Mar. Pollut. Bull., 36: 490–492. Duinker, J.C., Hillebrand, M.T.J., Zeinstra T., and Boon, J.P., 1989, Individual chlorinated biphenyls and pesticides in tissues of some cetacean species from the North Sea and the Atlantic Ocean: Tissue distribution and biotransformation, Aquat. Mammals, 15: 95–124. Estes, J.A., Bacon, C.E., Jarman, W.M., Norstrom, R.J., Anthony, R.G., and Miles, A.K, 1997, Organochlorines in sea otters and bald eagles from the Aleutian Archipelago, Mar. Pollut. Bull., 34: 486–490. Falandysz, J., Yamashita, N., Tanabe, S., Tatsukawa, R., Ruciÿska, L., and Skÿra, K., 1994, Congenerspecific data on polychlorinated biphenyls in tissues of common porpoise from Puck Bay, Baltic Sea, Arch. Environ. Contam. Toxicol., 26: 267–272.
0839_frame_C22.fm Page 507 Tuesday, May 22, 2001 11:51 AM
Toxicology
507
Falandysz, J., Strandberg, B., Strandberg, L., and Rappe, C., 1999, Tris(4-chlorophenyl) methane and tris(4-chlorophenyl) methanol in sediment and food webs from the Baltic south coast, Environ. Sci. Technol., 33: 517–521. Finklea, B., Miller, G., and Busbee, D., 2000, Polychlorinated biphenyl residues in blubber of male Atlantic bottlenose dolphins (Tursiops truncatus) that stranded and died at Matagorda Bay, Bull. Environ. Contam. Toxicol., 64: 323–332. Forcada, J., Hammond, P. S., and Aguilar, A., 1999, Status of the Mediterranean monk seal Monachus monachus in the Western Sahara and the implications of a mass mortality event, Mar. Ecol. Prog. Ser., 188: 249–261. Fossi, M.C., and Marsili, L, 1997, The use of non-destructive biomarkers in the study of marine mammals, Biomarkers, 2: 205–216. Fossi, M.C., Marsili, L., Leonzio, C., Notarbartolo di Sciara, G., Zanardelli, M., and Focardi, S., 1992, The use of non-destructive biomarker in Mediterranean cetaceans: Preliminary data on MFO activity in skin biopsy, Mar. Pollut. Bull., 24: 459–461. Fowler, M.E. (Ed.), 1993, Veterinary Zootoxicology, CRC Press, Boca Raton, FL, 250 pp. Frank, A., Galgan, V., Roos, A., Olsson, M., Peterson, L.R., and Bignert, A., 1992, Metal concentrations in seals from Swedish waters, Ambio, 21: 529–538. Freeman, H.C., and Sangalang, G.B., 1977, A study of the effects of methylmercury, cadmium, arsenic, selenium, and a PCB (Aroclor 1254) on adrenal and testicular steroidogeneses in vitro, by the gray seal, Arch. Environ. Contam. Toxicol., 5: 369–383. Freeman, H.C., Sanglang G., Uthe, J.F., and Ronald, K., 1975, Steroidogensis in vitro in the harp seal without and with methylmercury treatment in vivo, Environ. Physiol. Biochem., 5: 428–439. Frost, K.J., Lowry, L.F., Sinclair, E.H., Hoef, J.V., and McAllister, D.C., 1994, Impacts on distribution, abundance, and productivity of harbor seals, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 395 pp. Fujise, Y., Honda, K., Tatsukawa, R., and Mishima, S., 1988, Tissue distribution of heavy metals in Dall’s porpoise in the northwestern Pacific, Mar. Pollut. Bull., 19: 226–230. Fujise, Y., Zenati, R., and Kato, H., 1998, An examination of W-stock hypothesis for North Pacific minke whales, with special reference to some biological parameters using data collected from JARPA surveys from 1994–1997, Paper SC/50/RMP12, IWC Scientific Committee, Oman, 14 pp. Fuller, G.B., and Hobson, W.C., 1986, Effect of PCBs on reproduction in mammals, in PCBs and the Environment, Vol. 2, Waid, J.S. (Ed.), CRC Press, Boca Raton, FL, 101–125. Gaskin, D.E., Holdrinet, M., and Frank, R., 1982, DDT residues in blubber of harbor porpoise, Phocoena (L.), from eastern Canadian waters during the five-year period 1969–1973, in Mammals in the Seas, FAO Fisheries Series, No. 5, 4: 135–143. Gaskin, D.E., Frank, R., and Holdrinet, M., 1983, Polychlorinated biphenyls in harbor porpoises Phocoena phocoena (L.) from the Bay of Fundy, Canada and adjacent waters, with some information on chlordane and hexachlorobenzene levels, Arch. Environ. Contam. Toxicol., 12: 211–219. Gauthier, J.M., Metcalfe, C.D., and Sears, R., 1997, Chlorinated organic contaminants in blubber biopsies from Northwestern Atlantic balaenopterid whales summering in the Gulf of St. Lawrence, Mar. Environ. Res., 44: 201–223. Gauthier, J.M., Dubeau, H., and Rassart, E., 1998, Mercury-induced micronuclei in skin fibroblasts of beluga whales, Environ. Toxicol. Chem., 17: 2487–2493. Gauthier, J.M., Dubeau, H., and Rassart, É, 1999a, Induction of micronuclei in vitro by organochlorine compounds in beluga whale skin fibroblasts, Mutat. Res., 43: 87–95. Gauthier, J.M., Dubeau, H., Rassart, E., Jarman, W.M., and Wells, R.S., 1999b, Biomarkers of DNA damage in marine mammals, Mutat. Res., 44: 427–439. 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. Geraci, J.R., and St. Aubin, D.J. (Eds.), 1990, Sea Mammals and Oil: Confronting the Risks, Academic Press, San Diego, CA, 282 pp.
0839_frame_C22.fm Page 508 Tuesday, May 22, 2001 11:51 AM
508
CRC Handbook of Marine Mammal Medicine
Geraci, J.R., and Smith, T.G., 1976, Direct and indirect effects of oil on ringed seals of the Beaufort Sea, J. Fish. Res. Board Can., 33: 1976–1984. 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. Gilmartin, W.G., DeLong, R.L., Smith, A.W., Griner, L.A., and Dailey, M.D., 1980, An investigation into unusual mortality in the Hawaiian monk seal, Monachus schauinslandi, in Proceedings on Status of Resource Investigation in the Northwestern Hawaiian Islands, Grigg, R.W., and Pfund, R.T. (Eds.), University of Hawaii, Honolulu, UNIHI-SEAGRANT-MR-80-04, 32–41. Goksøyr, A., Tarlebö, J., Solbakken, J.E., and Klungsöyr, J., 1985, Characteristics of the hepatic microsomal cytochrome P-450 system of the minke whale (Balaenoptera acutorostrata), Mar. Environ. Res., 17: 113–116. Goksøyr, A., Solbakken, J.E., Tarlebo, J., and Klungsoyr, J., 1986, Initial characterization of the hepatic microsomal cytochrome P-450-system of the piked whale (minke) Balaenoptera acutorostrata, Mar. Environ. Res., 19: 185–203. Goksøyr, A., Andersson, T., Förlin, L., Stenersen, J., Snowberger, E.A., Woodin, B.R., and Stegeman, J.J., 1988, Xenobiotic and steroid metabolism in adult and fetal piked (minke) whales, Balaenoptera acutorostrata, Mar. Environ. Res., 24: 9–13. Goksøyr, A., Andersson, T., Förlin, L., Snowberger, E.A., Woodin, B.R., and Stegeman, J.J., 1989, Cytochrome P-450 monooxygenase activity and immuno-chemical properties of adult and fetal piked (minke) whales, Balaenoptera acutorostrata, in Cytochrome P-450: Biochemistry and Biophysics, Schuster, I. (Ed.), Taylor & Francis, New York, 698–701. Goksøyr, A., Beyer, J., Larsen, H.E., Andersson, T., and Förlin, L., 1992, Cytochrome P-450 in seals: Monooxygenase activities, immunochemical cross-reactions and response to phenobarbital treatment, Mar. Environ. Res., 34: 113–116. Goldblatt, C.J., and Anthony, R.G., 1983, Heavy metals in northern fur seals (Callorhinus ursinus) from the Pribilof Islands, Alaska, J. Environ. Qual., 12: 478–482. Goodwin, T., Hammond, T., Li, L., Linnehan, R., Kaysen, J., Albert, T.F., and O’Hara, T.M., 1999, Preliminary report on the molecular analysis and culture of tissues from subsistence harvested bowhead whales, presented at IWC Meeting (51st), Grenada, May 1–15, SC/51/AS13. Granby, K., and Kinze, C.C., 1991, Organochlorines in Danish and west Greenland harbour porpoises, Mar. Pollut. Bull., 22: 458–462. Green, N.J.L., Jones, K.C., and Harwood, J., 1996, Contribution of coplanar and non-coplanar polychlorinated biphenyls to the toxic equivalence of grey seal (Halichoerus grypus) milk, Chemosphere, 33: 1273–1281. Gulland, F.M.D., 2000, Domoic acid toxicity in California sea lions stranded along the central California coast, May–October 1998, NOAA Technical Memo, National Marine Fisheries Service, U.S. Department of Commerce, NMFS-OPR-17, 45 pp. Gulland, F.M.D., Trupkiewicz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996, Metastatic carcinoma of probable transitional cell origin in 66 free-living California sea lions (Zalophus californianus), 1979 to 1994, J. Wildl. Dis., 32: 250–258. Gunter, G., Williams, R.H., Davis, C.C., and Walton Smith, F.G., 1948, Catastrophic mass mortality of marine mammals and coincident phytoplankton bloom on the west coast of Florida, November 1946 to August 1947, Ecol. Monogr., 18: 309–324. Hall, A.J., Law, R.J., Wells, D.E., Harwood, J., Ross, H.M., Kennedy, S., Allchin, C.R., Campbell, L.A., and Pomeroy, P.P., 1992, Organochlorine levels in common seals (Phoca vitulina) which were victims and survivors of the 1988 phocine distemper epizootic, Sci. Total Environ., 115: 145–162. Hall, A.J., Pomeroy, P., Green, N., Jones, K., and Harwood, J., 1997, Infection, hematology and biochemistry in grey seal pups exposed to chlorinated biphenyls, Mar. Environ. Res., 43: 81–98.
0839_frame_C22.fm Page 509 Tuesday, May 22, 2001 11:51 AM
Toxicology
509
Harder, T.C., Willhaus, T., Leibold, W., and Liess, B., 1992, Investigations on course and outcome of phocine distemper virus infection in harbor seals (Phoca vitulina) exposed to polychlorinated biphenyls, J. Vet. Med. B, 39: 19–31. Harwood, J., 1998, What killed the monk seals? Nature, 393: 17–18. Haynes, D., Müller, J., and McLachlan, M., 1999, Polychlorinated dibenzo-p-dioxins and dibenzofurans in Great Barrier Reef (Australia) dugongs (Dugong dugon), Chemosphere, 38: 255–262. Helle, E., Olsson, M., and Jensen, S., 1976a, DDT and PCB levels and reproduction in ringed seal from the Bothnian Bay, Ambio, 5: 188–189. Helle, E., Olsson, M., and Jensen, S., 1976b, PCB levels correlated with pathological changes in seal uteri, Ambio, 5: 261–263. Hellou, J., Stenson, G., Ni, I.-H., and Payne, J.F., 1990, Polycyclic aromatic hydrocarbons in muscle tissue of marine mammals from the Northwest Atlantic, Mar. Pollut. Bull., 21: 469–473. 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. Hickie, B.E., Mackay, D., and De Koning, J., 1999, Lifetime pharmacokinetic model for hydrophobic contaminants in marine mammals, Environ. Toxicol. Chem., 18: 2622–2633. Holden, A.V., 1978, Pollutants and seals: A review, Mammal Rev., 8: 53–66. Holsbeek, L., Joiris, C.R., Debacker, V., Ali, I. B., Roose, P., Nellissen, J.-P., Gobert, S., Bouquegneau, J.-M., and Bossicart, M., 1999, Heavy metals, organochlorines and polycyclic aromatic hydrocarbons in sperm whales stranded in the southern North Sea during the 1994/1995 winter, Mar. Pollut. Bull., 38: 304–313. Honda, K., Tatsukawa, R., Itano, K., Miyazaki, N., and Fujiyama, T., 1983, Heavy metal concentration in muscle, liver, and kidney tissue of striped dolphin, Stenella coeruleoalba, and their variations with body length, weight, age and sex, Agric. Biol. Chem., 47: 1219–1228. Honda, K., Fujise, Y., Tatsukawa, R., Itano, K., and Miyazaki, N., 1986a, Age-related accumulation of heavy metals in bone of the striped dolphin, Mar. Environ. Res., 20: 143–160. Honda, K., Yamamoto, Y., and Tatsukawa, R., 1986b, Heavy metal accumulation in the liver of Antarctic minke whale, Mem. Nat. Inst. Polar Res., 44: 182–184. Honda, K., Yamamoto, Y., Kato, H., and Tatsukawa, R., 1987, Heavy metal accumulations and their recent changes in southern minke whales, Arch. Environ. Contam. Toxicol., 16: 209–216. Iverson, F., and Truelove, J., 1994, Toxicology and seafood toxins: Domoic acid, Nat. Toxins, 2: 334–339. Iwata, H., Tanabe, S., Miyazaki, N., and Tatsukawa, R., 1994, Detection of butyltin compound residues in blubber of marine mammals, Mar. Pollut. Bull., 28: 607–612. Jarman, W.M., Simon, M., Norstrom, R.J., Bruns, S.A., Bacon, C.A., Simaret, B.R.T., and Risebrough, R.W., 1992, Global distribution of tris-(4-chlorophenyl) methanol in high trophic level birds and mammals, Environ. Sci. Technol., 26: 1770–1774. Jarman, W.M., Norstrom, R.J., Muir, D.C.G., Rosenberg, B., Simon, M., and Baird, R.W., 1996a, Levels of organochlorine compounds, including PCDDs and PCDFs, in the blubber of cetaceans from the west coast of North America, Mar. Pollut. Bull., 32: 426–436. Jarman, W.M., Bacon, C.E., Estes, J.A., Simon, M., and Norstrom, R.J., 1996b, Organochlorine contaminants in sea otters: The sea otter as a bioindicator, Endangered Species Update, 13: 20–22. Jepson, P.D., Bennett, P.M., Allchin, C.R., Law, R.J., Kuiken, T., Baker, J.R., Rogan, E., and Kirkwood, J.K., 1999, Investigating potential associations between chronic exposure to polychlorinated biphenyls and infectious disease mortality in harbour porpoises from England and Wales, Sci. Total Environ., 243/244: 339–348. Jiménez, B., González, M.J., Hernández, L.M., Eljarrat, E., Rivera, J., and Fossi, M.C., 1999, 2,3,7,8Substituted PCDDs and PCDFs in sea lion (Otaria flavescens) skin biopsies from two southwestern Atlantic populations, Chemosphere, 38: 507–515.
0839_frame_C22.fm Page 510 Tuesday, May 22, 2001 11:51 AM
510
CRC Handbook of Marine Mammal Medicine
Kamrin, M.A., and Ringer, R.K., 1996, Toxicological implications of PCB residues in mammals, 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, 153–163. Kannan, K., Falandysz, J., Tanabe, S., and Tatsukawa, R., 1993, Persistent organochlorines in harbour porpoises from Puck Bay, Poland, Mar. Pollut. Bull., 26: 162–165. Kannan, K.S., Corsolini, S., Focardi, S., Tanabe, S., and Tatsukawa, S., 1996, Accumulation pattern of butyltin compounds in dolphin, tuna, and shark collected from Italian coastal waters, Arch. Environ. Contam. Toxicol., 31: 19–23. Kannan, K., Senthilkumar, K., Loganathan, B.G., Takahashi, S., Odell, D.K., and Tanabe, S., 1997, Elevated accumulation of tributyltin and its breakdown products in bottlenose dolphins (Tursiops truncatus) found stranded along the U.S. Atlantic and Gulf Coasts, Environ. Sci. Technol., 31: 296–301. Kannan, K., Guruge, K.S., Thomas, N.J., Tanabe, S., and Giesy, J.P., 1998, Butyltin residues in southern sea otters (Enhydra lutris nereis) found dead along California coastal waters, Environ. Sci. Technol., 32: 1169–1175. Kannan, N., Tanabe, S., Ono, M., and Tatsukawa, R., 1989, Critical evaluation of polychlorinated biphenyl toxicity in terrestrial and marine mammals: Increasing impact of non-ortho and mono-ortho coplanar polychlorinated biphenyls from land to ocean, Arch. Environ. Contam. Toxicol., 18: 850–857. Kemper, C., Gibbs, P., Obendorf, P., Marvanek, S., and Lenghaus, C., 1994, A review of heavy metal and organochlorine levels in marine mammals in Australia, Sci. Total Environ., 154: 129–139. Kendall, M.D., Safieh, B., Harwood, J., and Pomeroy, P.P., 1992, Plasma thymulin concentrations, the thymus and organochlorine contaminant levels in seals infected with phocine distemper virus, Sci. Total Environ., 115: 133–144. Kennedy, S., 1999, Morbilliviral infections in marine mammals, J. Cetacean Res. Manage., 1: 267–273. Kiceniuk, J.W., Holzecher, J., and Chatt, A., 1997, Extractable organohalogens in tissues of beluga whales from the Canadian Arctic and the St. Lawrence estuary, Environ. Pollut., 97: 205–211. Kihlström, J.E., Olsson, M., Jensen, S., Johansson, A., Ahlborn, J., and Bergman, A., 1992, Effects of PCB and different fractions of PCB on the reproduction of the mink (Mustela vison), Ambio, 21: 563–569. Kim, G.B., Lee, J.S., Tanabe, S., Iwata, H., Tatsukawa, R., and Shimazaki, K., 1996, Specific accumulation and distribution of butyltin compounds in various organs and tissues of the Steller sea lion: Comparison with organochlorine accumulation pattern, Mar. Pollut. Bull., 32: 558–563. Kim, G.B., Nakata, H., and Tanabe, S., 1998, In vitro inhibition of hepatic cytochrome P450 and enzyme activity by butyltin compounds in marine mammals, Environ. Pollut., 99: 255–261. Kingsley, M.C.S., 1998, Population index estimates for the St. Lawrence belugas, 1973–1995, Mar. Mammal Sci., 14: 508–530. Kingsley, M.C.S., 1999, Population indices and estimates for the belugas of the St. Lawrence estuary, Canadian Technical Report of Fisheries and Aquatic Sciences, 2266, Fisheries and Oceans Canada, 27. Kleivane, L., Skaare, J.U., Bjørge, A., de Ruiter, E., and Reijnders, P.J.H., 1995, Organochlorine pesticide residue and PCBs in harbor porpoise (Phocoena phocoena) incidentally caught in Scandinavian waters, Environ. Pollut., 89: 137–146. Koeman, J.H., 1973, PCB in mammals and birds in the Netherlands, in PCB Conference II, Ahnland, E., and Akerblom, A. (Eds.), National Swedish Environmental Protection Board, Stockholm, publ. 1973:4E, 35–43. Koeman, J.H., and van de Ven, W.S.M., 1975, Mercury and selenium in marine mammals and birds, Sci. Total Environ., 3: 279–287. Krone, C.A., Robisch, P.A., Tilbury, K.L., Stein, J.E., Mackey, E.A., Becker, P.R., O’Hara, T.M., and Philo, L.M., 1999, Elements in liver tissues of bowhead whales (Balaena mysticetus), Mar. Mammal Sci., 15: 123–142.
0839_frame_C22.fm Page 511 Tuesday, May 22, 2001 11:51 AM
Toxicology
511
Kuehl, D., and Haebler, R., 1995, Organochlorine, organobromine, metal, and selenium residues in bottlenose dolphins (Tursiops truncatus) collected during an unusual mortality event in the Gulf of Mexico, 1990, Arch. Environ. Contam. Toxicol., 28: 494–499. Kuehl, D.W., Haebler, R., and Potter, C., 1991, Chemical residues in dolphins from the U.S. Atlantic Coast including Atlantic bottlenose obtained during the 1987/88 mass mortality, Chemosphere, 22: 1071–1084. Kuehl, D.W., Haebler, R., and Potter, C., 1994, Coplanar PCB and metal residues in dolphins from the U.S. Atlantic coast including Atlantic bottlenose obtained during the 1987/1988 mass mortality, Chemosphere, 23: 1245–1253. Kuiken, T., Höfle, U., Bennett, P.M., Allchin, C.R., Kirkwood, J.K., Baker, J.R., Appleby, E.C., Lockyer, C.H., Walton, M.J., and Sheldrick, M.C., 1993, Adrenocortical hyperplasia, disease and chlorinated hydrocarbons in the harbor porpoise (Phocoena phocoena), Mar. Pollut. Bull., 26: 440–446. Kuiken, T., Bennett, P.M., Allchin, C.R., Kirkwood, J.K., Baker, J.R., Lockyer, C.H., Walton, M.J., and Sheldrick, M.C., 1994, PCBs, cause of death and body condition in harbor porpoises (Phocoena phocoena) from British waters, Aquat. Toxicol., 28: 13–28. Kvitek, R.G., DeGange, A.R., and Beitler, M.K., 1991, Paralytic shellfish poisoning toxins mediate feeding behavior of sea otter, Limnol. Oceanogr., 36: 393–404. Kvitek, R.G., Bowlby, C.E., and Staedler, M., 1993, Diet and foraging behavior of sea otters in southeast Alaska, Mar. Mammal Sci., 9: 168–181. Lahvis, G.P., Wells, R.S., Kuehl, D.W., Stewart, J.L., Rhinehart, H.L., and Via, C.S., 1995, Decreased lymphocyte responses in free-ranging bottlenose dolphins (Tursiops truncatus) are associated with increased concentrations of PCBs and DDT in peripheral blood, Environ. Health Perspect., 103: 62–67. Lapierre, P., De Guise, S., Muir, D.C.G., Norstrom, R., Beland, P., and Fournier, M., 1999, Immune functions in the Fisher rat fed beluga whale (Delphinapterus leucas) blubber from the contaminated St. Lawrence estuary, Environ. Res., A80: S104–S112. Law, R.J., and Whinnet, J.A., 1992, Polycyclic aromatic hydrocarbons in muscle tissue of harbor porpoises from U.K. waters, Mar. Pollut. Bull., 24: 550–553. Law, R.J., Fileman, C.F., Hopkins, A.D., Baker, J.R., Harwood, J., Jackson, D.B., Kennedy, S., Martin, A.R., and Morris, R.J., 1991, Concentrations of trace metals in the livers of marine mammals (seals, porpoises, and dolphins) from waters around the British Isles, Mar. Pollut. Bull., 22: 183–191. Law, R.J., Stringer, R.L., Allchin, C.L., and Jones, B.R., 1996, Metals and organochlorines in sperm whales stranded around the North Sea during the 1994/95 winter, Mar. Pollut. Bull., 32: 72–77. Layne, J.N., 1965, Observations of marine mammals in Florida waters, Bull. Fl. State Mus., 9: 131–181. Lee, J.S., Tanabe, S., Umino, H., Tatsukawa, R., Loughlin, T.R., and Calkins, D.C., 1996, Persistent organochlorines in Steller sea lion (Eumetopias jubatus) from the bulk of Alaska and the Bering Sea, 1976–1981, Mar. Pollut. Bull., 32: 535–544. Lee, S.S., Mate, B.R., von der Trenck, K.T., Rimerman, R.A., and Buhler, D.R., 1977, Metallothionein and the subcellular localization of mercury and cadmium in the California sea lion, Comp. Biochem. Physiol., C57: 45–53. Leonzio, C., Focardi, S., and Fossi, C., 1992, Heavy metals and selenium in stranded dolphins of the northern Tyrrhenian (NW Mediterranean), Sci. Total Environ., 119: 77–84. Letcher, R.J., Norstrom, R.J., and Bergman, A., 1995, Geographical distribution and identification of methyl sulfone PCB and DDE metabolites in pooled polar bear (Ursus maritimus) adipose tissue from Western Hemisphere Arctic and subarctic regions, Sci. Total Environ., 160/161: 409– 420. Letcher, R.J. Norstrom, R.J., Lin, S., Ramsay, M.A., and Bandiera, S.M., 1996, Immunoquantitation and microsomal monoxygenase activities of hepatic cytochromes P4501A and P4502B and chlorinated hydrocarbon contaminant levels in polar bear (Ursus maritimus), Toxicol. Appl. Pharmacol., 137: 127–140.
0839_frame_C22.fm Page 512 Tuesday, May 22, 2001 11:51 AM
512
CRC Handbook of Marine Mammal Medicine
Letcher, R.J., Norstrom, R.J., and Muir, D.C.G., 1998, Biotransformation versus bioaccumulation: Sources of methyl sulfone PCB and 4,4′-DDE metabolites in the polar bear food chain, Environ. Sci. Technol., 32: 1656–1661. Lindström, G., Wingfors, H., Dam, M., and Bavel, B.V., 1999, Identification of 19 polybrominated diphenyl ethers (PBDEs) in long-finned pilot whale (Globicephala melas) from the Atlantic, Arch. Environ. Contam. Toxicol., 36: 355–363. Linzey, A.V., 1987, Effects of chronic polychlorinated biphenyls exposure on reproductive success of white-footed mice (Peromyscus leucopus), Arch. Environ. Contam. Toxicol., 16: 455–460. Linzey, A.V., 1988, Effects of chronic polychlorinated biphenyls exposure on growth and reproduction of second generation white-footed mice, Arch. Environ. Contam. Toxicol., 17: 39–45. Lipscomb, T.P., Harris, R.K., Moeller, R.B., Pletcher, J.M., Haebler, R.J., and Ballachey, B.E., 1993, Histopathologic lesions in sea otters exposed to crude oil, Vet. Pathol., 30: 1–11. Lipscomb, T.P., Harris, R.K., Rebar, A.H., Ballachey, B.E., and Haebler, R.J., 1994, Pathology of sea otters, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 265–280. Lockyer, C.H., McConnell, L.C., and Waters, T.D., 1984, The biochemical composition of fin whale blubber, Can. J. Zool., 62: 2553–2562. Lorgue, G., Lechenet, J., and Riviere, A., 1996, Clinical Veterinary Toxicology, Blackwell Sciences, Cambridge, MA, 210 pp. Loughlin, T.R. (Ed.), 1994, Marine Mammals and the Exxon Valdez, Academic Press, San Diego, CA, 395 pp. Louise Armstrong, H.E., Corns, W.T., Stockwell, P.B., O’Connor, G., Ebdon, L., and Evans, E.H., 1999, Comparison of AFS and ICP-MS detection coupled with gas chromatography for the determination of methylmercury in marine samples, Anal. Chim. Acta, 390: 245–253. Mackey, E.A., Demiralp, R., Becker, P.R., Greenberg, R.R., Koster, B.J., and Wise, S.A., 1995, Trace element concentrations in cetacean liver tissues archived in the national marine mammal tissue bank, Sci. Total Environ., 175: 25–41. Mackey, E.A., Becker, P.R., Demiralp, R., Greenberg, R.R., Koster, B.J., and Wise, S.A., 1996, Bioaccumulation of vanadium and other trace metals in livers of Alaskan cetaceans and pinnipeds., Arch. Environ. Contam. Toxicol., 30: 503–512. Malcolm, H.M., Boyd, I.L., Osborn, D., French, M.C., and Freestone, P., 1994, Trace metals in Antarctic fur seal livers from Bird Island, South Georgia, Mar. Pollut. Bull., 28: 375–380. Marchovecchio, J.E., Moreno, V.J., Bastida, R.O., Gerpe, M.S., and Rodriguez, D.H., 1990, Tissue heavy metal distribution in small cetaceans from the southwestern Atlantic Ocean, Mar. Pollut. Bull., 21: 299–304. Marsili, L., and Focardi, S., 1997, Chlorinated hydrocarbon (HCB, DDTs and PCBs) levels in cetaceans stranded along the Italian coasts: An overview, Environ. Monit. Assessments, 45: 129–180. Marsili, L., Fossi, M.C., Notarbartolo di Sciara, G., Zanardelli, M., Nani, B., Panigada, S., and Focardi, S., 1998, Relationship between organochlorine contaminants and mixed function oxidase activity in skin biopsy specimens of Mediterranean fin whales (Balaenoptera physalus), Chemosphere, 37: 1501–1510. Martin, J.H., Elliot, P.D., Anderlini, V.C., Girvin, D., Jacobs, S.A., Risebrough, R.W., Delong, R.L., and Gilmartin, W.G., 1976, Mercury-selenium-bromine imbalance in premature California sea lions, Mar. Biol., 35: 91–104. Martineau, D., Béland, P., Desjardins, C., and Lagacé, A., 1987, Levels of organochlorine chemicals in tissues of beluga whales (Delphinapterus leucas) from the St. Lawrence estuary, Quebec, Canada, Arch. Environ. Contam. Toxicol., 16: 137–147. Martineau, D., Lagace, A., Beland, P., Higgins, R., Armstrong, D., and Shugart, L.R., 1988, Pathology of stranded beluga whales from the St. Lawrence estuary, Quebec, Can. J. Comp. Pathol., 98: 287–311.
0839_frame_C22.fm Page 513 Tuesday, May 22, 2001 11:51 AM
Toxicology
513
Martineau, D., De Guise, S., Fournier, M., Shugart, L., Girard, C., Lagacé, A., and Béland, P., 1994, Pathology and toxicology of beluga whales from the St. Lawrence estuary, Quebec, Canada, past, present and future, Sci. Total Environ., 154: 201–215. Martineau, D., Lair, S., De Guise, S., Lipscomb, T.P., and Béland, P., 1999, Cancer in beluga whales from the St. Lawrence estuary, Quebec, Canada: A potential biomarker of environmental contamination, J. Cetacean Res. Manage., 1: 249–265. Martoja, R., and Viale, D., 1977, Accumulation de granules de séléniure mercurique dans le foie d’Odontocètes (Mammifères, Cétacés): Un mécanisme possible de détoxication du méthyl-mercure par le sélénium, C. R. Acad. Sci. Paris D, 285: 109–114. Meador, J.P., Varanasi, U., Robisch, P.A., and Chan, S.L., 1993, Toxic metals in pilot whales from strandings in 1986 and 1990 on Cape Cod, Massachusetts, Can. J. Fish. Aquat. Sci., 50: 2698–2706. Miller, P.E., and Scholin, C.A., 1998, Identification and enumeration of cultured and wild Pseudonitzschia (Bacillariophycae) using species-specific LSU rRNA-targeted fluorescent probes and filterbased whole cell hybridization, J. Phycol., 34: 371–382. Minh, T.B., Watanabe, M., Nakata, H., Tanabe, S., and Jefferson, T.A., 1999, Contamination by persistent organochlorines in small cetaceans from Hong Kong coastal waters, Mar. Pollut. Bull., 39: 383–392. Miyazaki, N., Itano, K., Fukushima, M., Kawai, S.-I., and Honda, K., 1979, Metals and organochlorine compounds in the muscle of dugong from Sulawesi Island, Sci. Rep. Whales Res. Inst. (Japan), 31: 125–128. Mochizuki, Y., Suzuke, K.T., Sunaga, H., Kobayashi, T., and Doi, R., 1985, Separation and characterization of metallothionein in two species of seals by high performance liquid chromatography–atomic absorption spectrophotometry, Comp. Biochem. Physiol. C, 82: 249–254. Mortensen, P., Bergman, A., Bignert, A., Hansen, H.-J., Harkonen, T., and Olsson, M., 1992, Prevalence of skull lesions in harbor seals (Phoca vitulina) in Swedish and Danish museum collections: 1835–1988, Ambio, 21: 520–524. Muir, D.C.G., Wagemann, R., Grift, N.P., Norstrom, R.J., Simon, M., and Lien, J., 1988a, Organochlorine chemical and heavy metal contaminants in white-beaked dolphins (Lagenorhynchus albirostris) and pilot whales (Globicephala melaena) from the coast of Newfoundland, Canada, Arch. Environ. Contam. Toxicol., 17: 613–629. Muir, D.C.G., Norstrom, R.J., and Simon, M., 1988b, Organochlorine contaminants in Arctic marine food chains: Accumulation of specific polychlorinated biphenyls and chlordane-related compounds, Environ. Sci. Technol., 22: 1071–1079. Muir, D.C.G., Ford, C.A., Stewart, R.E.A., Smith, T.G., Addison, R.F., Zinck, M.E., and Béland, P., 1990, Organochlorine contaminants in belugas, Delphinapterus leucas, from Canadian waters, in Advances in Research on the Beluga Whale, Delphinapterus leuca, Smith, T.G., St. Aubin, D.J., and Geraci, J.R. (Eds.), Can. Bull. Fish. Aquat. Sci., 224: 165–190. Muir, D.C.G., Wagemann, R., Hargrave, B.T., Thomas, D.J., Peakall, D.B., and Norstrom, R.J., 1992a, Arctic marine ecosystem contamination, Sci. Total Environ., 122: 75–134. Muir, D.C.G., Ford, C.A., Grift, N.P., and Stewart, R.E.A., 1992b, Organochlorine contaminants in narwhal (Monodon monoceros) from the Canadian Arctic, Environ. Pollut., 75: 307–316. Muir, D.C.G., Ford, C.A., Rosenberg, B., Norstrom, R.J., Simon, M., and Béland, P., 1996a, Persistent organochlorines in beluga whales (Delphinapterus leucas) from the St. Lawrence River estuary, I. Concentrations and patterns of specific PCBs, chlorinated pesticides and polychlorinated dibenzo-p-dioxins and dibenzofurans, Environ. Pollut., 93: 219–234. Muir, D.C.G., Koczanski, K., Rosenberg, B., and Béland, P., 1996b, Persistent organochlorines in beluga whales (Delphinaterus leucas) from the St. Lawrence River estuary. II. Temporal trends, 1982–1994, Environ. Pollut., 93: 235–245. Muir, D.C.G., Born, E.W., Koczansky, K., and Stern, G.A., 2000, Temporal and spatial trends of persistent organochlorines in Greenland walrus (Odobenus rosmarus rosmarus), Sci. Total Environ., 245: 73–86.
0839_frame_C22.fm Page 514 Tuesday, May 22, 2001 11:51 AM
514
CRC Handbook of Marine Mammal Medicine
Mulcahy, D.M., and Ballachey, B.E., 1994, Hydrocarbon residues in sea otter tissues, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 395 pp. Nakata, H., Kannan, K., Jing, L., Thomas, N., Tanabe, S., and Giesy, J. P., 1998, Accumulation pattern of organochlorine pesticides and polychlorinated biphenyls in southern sea otters (Enhydra lutris nereis) found stranded along coastal California, USA, Environ. Pollut., 103: 45–53. National Research Council, 1999, Hormonally Active Agents in the Environment, National Academy of Sciences Press, Washington, D.C., 430 pp. Nebert, D.W., and Gonzalez, F.J., 1987, P450 genes: Structure, evolution, and regulation, Annu. Rev. Biochem., 56: 945–993. Neff, J.M., 1990a, Composition and fate of petroleum and spill-treating agents in the marine environment, in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, New York, 1–33. Neff, J.M., 1990b, Effects of oil on marine mammal populations: Model simulations in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, New York, 35–54. Nendza, M., Herbst, T., Kussatz, C., and Gies, A., 1997, Potential for secondary poisoning and biomagnification in marine organisms, Chemosphere, 35: 1875–1885. Nigro, M., and Leonzio, C., 1996, Intracellular storage of mercury and selenium in different marine vertebrates, Mar. Ecol. Prog. Ser., 135: 137–143. Noda, K., Ichihashi, H., Loughlin, T.R., Baba, N., Kiyota, M., and Tatsukawa, R., 1995, Distribution of heavy metals in muscle, liver and kidney of northern fur seal caught off Sanriku, Japan and from the Pribilof Islands, Alaska, Environ. Pollut., 90: 51–59. Norheim, G., Skaare, J.U., and Wiig, O., 1992, Some heavy metals, essential elements, and chlorinated hydrocarbons in polar bear (Ursus maritimus) at Svalbard, Environ. Pollut., 77: 51–57. Norstrom, R.J., and Muir, D.C.G., 1994, Chlorinated hydrocarbon contaminants in Arctic marine mammals, Sci. Total Environ., 154: 107–128. Norstrom, R.J., Schweinsberg, R.E., and Collins, B.T., 1986, Heavy metals and essential elements in livers of the polar bear (Ursus maritimus) in the Canadian Arctic, Sci. Total Environ., 48: 195–212. Norstrom, R.J., Simon, M., Muir, D.C.G., and Schweinsburg, R.E., 1988, Organochlorine contaminants in Arctic marine food chains: Identification, geographical distribution and temporal trends in polar bears, Environ. Sci. Technol., 22: 1063–1071. Norstrom, R.J., Simon, M., and Muir, D.C.G., 1990, Polychlorinated dibenzo-p-dioxins and dibenzofurans in marine mammals in the Canadian north, Environ. Pollut., 66: 1–19. Norstrom, R.J., Belikov, S.E., Born, E.W., Garner, G.W., Malone, B., Olpinski, S., Ramsay, M.A., Schliebe, S., Stirling, I., Stishov, M.S., Taylor, M.K., and Wiig, Ø., 1998, Chlorinated hydrocarbon contaminants in polar bears from Eastern Russia, North America, Greenland, and Svalbard: Biomonitoring of Arctic pollution, Arch. Environ. Contam. Toxicol., 35: 354–367. Ochoa, J.I., Sierra-Beltran, A.P., Olaiz-Fernandez, G., and Del Villar Ponce, L.M., 1998, Should mollusk toxicity in Mexico be considered a public health issue? J. Shellfish Res., 17: 1671–1673. Oehme, M., Schabach, M., Hummert, K., Luckas, B., and Nordoy, E.S., 1995, Determination of levels of polychlorinated dibenzo-p-dioxins, dibenzofurans, biphenyls and pesticides in harp seals from the Greenland Sea, Sci. Total Environ., 162: 75–91. O’Hara, T.M., and Rice, C., 1996, Polychlorinated biphenyls, in Noninfectious Diseases of Wildlife, 2nd ed., Fairbrother, A., Locke, L., and Hoff, G. (Eds.), Iowa State University Press, Ames, 71–86. 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. Olafson, R.W., and Thompson, J.A.J., 1974, Isolation of heavy metal binding proteins from marine vertebrates, Mar. Biol., 28: 83–86. Olsson, M., Karlsson, B., and Ahnland, E., 1994, Diseases and environmental contaminants in seals from the Baltic and the Swedish west coast, Sci. Total Environ., 154: 217–227.
0839_frame_C22.fm Page 515 Tuesday, May 22, 2001 11:51 AM
Toxicology
515
O’Shea, T.J., 1999, Environmental contaminants and marine mammals, in Biology of Marine Mammals, Reynolds III, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 485–564. O’Shea, T.J., and Brownell, R.L., Jr., 1994, Organochlorine and metal contaminants in baleen whales: A review and evaluation of conservation implications, Sci. Total Environ., 154: 179–200. O’Shea, T.J., and Brownell, R.L., Jr., 1998, California sea lion (Zalophus californianus) populations and sDDT contamination, Mar. Pollut. Bull., 36: 159–164. O’Shea, T.J., and Tanabe, S., in press, Environmental toxicology and marine mammals: A retrospective overview, in Toxicology in Marine Mammals, Vos, J., Bossart, G., and Fournier, M. (Eds.), Taylor & Francis, London. O’Shea, T.J., Brownell, R.L., Jr., Clark, D.R., Jr., Walker, W.A., Gay, M.L., and Lamont, T.G., 1980, Organochlorine pollutants in small cetaceans from the Pacific and South Atlantic Oceans, November 1968–June 1976, Pest. Monit. J., 14: 35–46. O’Shea, T.J., Moore, J.F., and Kochman, H.I., 1984, Contaminant concentrations in manatees in Florida, J. Wildl. Manage., 48: 741–748. O’Shea, T.J., Rathbun, G.B., Bonde, R.K., Buergelt, C.D., and Odell, D.K., 1991, An epizootic of Florida manatees associated with a dinoflagellate bloom, Mar. Mammal Sci., 7: 165–179. O’Shea, T.J., Reeves, R.R., and Long, A.K. (Eds.), 1999, Marine mammals and persistent ocean contaminants, in Proceedings of a Workshop held in Keystone, Colorado, October 12–15, 1998, U.S. Marine Mammal Commission, Washington, D.C., 150. Osterhaus, A., Groen, I., Niesters, H., van de Bildt, M.B., Vedder, L., Vos, J., Van Egmond, H., Sibi Abou, B., and Ould Barham, M.H., 1997, [untitled], Nature, 388: 838. Osweiller, G.D., Carson, T.L., Buck, W.B., and Van Gelder, G.A., 1985, Clinical and Diagnostic Veterinary Toxicology, 3rd ed., Kendall/Hunt, Dubuque, IA, 494. Paludan-Müller, P., Agger, C.T., Dietz, R., and Kinze, C.C., 1993, Mercury, cadmium, zinc, copper and selenium in harbor porpoise (Phocoena phocoena) from west Greenland, Polar Biol., 13: 311–320. Parsons, E.C.M., 1998, Trace metal pollution in Hong Kong: Implications for the health of Hong Kong’s Indo-Pacific hump-backed dolphins (Sousa chinensis), Sci. Total Environ., 214: 175–184. Parsons, E.C.M., 1999, Trace element concentrations in the tissues of cetaceans from Hong Kong’s territorial waters, Environ. Conserv., 26: 30–40. Peakall, D., 1992, Animal Biomarkers as Pollution Indicators, Chapman & Hall, London, 291 pp. Peakall, D., 1999, Biomarkers as pollution indicators with special reference to cetaceans, J. Cetacean Res. Manage., 1: 117–124. Pitot III, H.C., and Dragan, Y.P., 1996, Chemical carcinogenesis, in Casarett and Doull’s Toxicology: The Basic Science of Poisons, 5th ed., Klaasen, C.D. (Ed.), McGraw-Hill, Health Professions Division, New York, 201–267. Plumb, D.C., 1999, Veterinary Drug Handbook, 3rd ed., Iowa State University Press, Ames, 711 pp. Preen, A., 1988, The status and conservation of the dugong in the Arabian region, final report, Meteorology and Environmental Protection Administration, Saudia Arabia, Vol. 2. Ramprashad, F., and Ronald, K.A., 1977, Surface preparation study on the effect of methylmercury on the sensory hair cell population in the cochlea of the harp seal, Pagophilus groenlandicus (Erxleben, 1777), Can. J. Zool., 55: 223–230. Rattner, B.A., Hoffman, D.J., and Marn, C.M., 1989, Use of mixed-function oxygenases to monitor contaminant exposure in wildlife, Environ. Toxicol. Chem., 8: 1093–1102. Rawson, A.J., Patton, G.W., Hofmann, S., Pietra, G.G., and Johns, L., 1993, Liver abnormalities associated with chronic mercury accumulation in stranded Atlantic bottlenose dolphins, Ecotoxicol. Environ., 25: 41–47. Rawson, A.J., Bradley, J.P., Teetsov, A., Rice, S.B., Haller, E.M., and Patton, G.W., 1995, A role for airborne particulates in high mercury levels of some cetaceans, Ecotoxicol. Environ. Saf., 30: 309–314.
0839_frame_C22.fm Page 516 Tuesday, May 22, 2001 11:51 AM
516
CRC Handbook of Marine Mammal Medicine
Ray, S., Dunn, B.P., Payne, J.F., Fancey, L., Helbig, R., and Beland, P., 1992, Aromatic DNA-carcinogen adducts in beluga whales from the Canadian Arctic and Gulf of St. Lawrence, Mar. Pollut. Bull., 22: 392–396. Reijnders, P.J.H., 1980, Organochlorine and heavy metal residues in harbor seals from the Wadden Sea and their possible effects on reproduction, Neth. J. Sea Res., 14: 30–65. 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., 1994, Toxicokinetics of chlorobiphenyls and associated physiological responses in marine mammals, with particular reference to their potential for ecotoxicological risk assessment, Sci. Total Environ., 154: 229–236. Reijnders, P.J.H., Aguilar, A., and Donovan, G.P., 1999, Chemical pollutants and cetaceans, J. Cetacean Res. Manage., 1: 273. Renberg, L., Sundström, G., and Reutergardh, L., 1978, Polychlorinated terphenyls (PCT) in Swedish white-tailed eagles and in grey seals: A preliminary study, Chemosphere, 6: 477–482. Ridlington, J.W., and Whanger, P.D., 1981, Interactions of selenium and antioxidants with mercury, cadmium and silver, Fundam. Appl. Toxicol., 1: 368–375. Riedman, M.L., and Estes, J.A., 1990, The sea otter (Enhydra lutris): Behavior, ecology, and natural history, U.S. Fish Wildl. Serv. Biol. Rep., 90: 1–126. Roberts, T.M., Heppleston, P.B., and Roberts, R.D., 1976, Distribution of heavy metals in tissues of the common seal, Mar. Pollut. Bull., 7: 194–196. Ronald, K., Tessaro, S.V., Uthe, J.F., Freeman, H.C., and Frank, R., 1977, Methylmercury poisoning in the harp seal (Pagophilus groenlandicus), Sci. Total Environ., 8: 1–11. Rosas, F.C., and Lehti, K.K., 1996, Nutritional and mercury content of milk of the Amazon river dolphin, Inia geoffrensis, Comp. Biochem. Physiol., 115A: 117–119. 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. Safe, S., 1986, Comparative toxicology and mechanism of action of polychlorinated dibenzo-p-dioxins and dibenzofurans, Ann. Rev. Pharmacol. Toxicol., 26: 371–399. Safe, S., 1991, Polychlorinated dibenzo-p-dioxins and related compounds: Sources, environmental distribution, and risk assessment, Environ. Carcinogenesis Ecotoxicol. Rev., C9: 261–302. Safe, S.H., 1994, Polychlorinated biphenyls (PCBs): Environmental impact, biochemical and toxic responses, and implications for risk assessment, Crit. Rev. Toxicol., 24: 87–149. St. Aubin, D.J., 1990a, Physiologic and toxic effects on pinnipeds, in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, New York, 103–127. St. Aubin, D.J., 1990b, Physiologic and toxic effects on polar bears, in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, New York, 235–240. St. Aubin, D.J., Geraci, J.R., Smith, T.G., and Friesen, T.G., 1985, How do bottlenose dolphins, Tursiops truncatas, react to oil films under different light conditions? Can. J. Fish. Aquat. Sci., 42: 430– 436. Salata, G.G., Wade, T.L., Sericano, J.L., Davis, J.W., and Brooks, J.M., 1995, Analysis of Gulf of Mexico bottlenose dolphins for organochlorine pesticides and PCBs, Environ. Pollut., 88: 167–175. Sanpera, C., Capelli, R., Minganti, V., and Jover, L., 1993, Total and organic mercury in North Atlantic fin whales: Distribution pattern and biological related changes, Mar. Pollut. Bull., 26: 135–139. Sanpera, C., Gonzalez, M., and Jover, L., 1996, Heavy metal levels in two populations of North Atlantic fin whales, Environ. Pollut., 91: 299–307. Schmidt-Nielsen, K., 1972, How Animals Work, Cambridge University Press, London, 114 pp. Scholin, C.A., Miller, P.E., Buck, K.R., and Chavez, F.P., 1997, Detection and quantification of Pseudonitzschia australis in cultured and natural populations using LSU rRNA-targeted probes, Limnol. Oceanogr., 42: 1265–1272.
0839_frame_C22.fm Page 517 Tuesday, May 22, 2001 11:51 AM
Toxicology
517
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. Schumacher, U., Zahler, S., Horney, H.-P., Hiedemann, G., Skirnisson, K., and Welsch, U., 1993, Histological investigations on the thyroid glands of marine mammals (Phoca vitulina, Phocoena phocoena) and the possible implications of marine pollution, J. Wildl. Dis., 29: 103–108. Senthilkumar, K., Kannan, K., Sinha, R., Tanabe, S., and Giesy, J.P., 1999, Bioaccumulation profiles of polychlorinated biphenyl congeners and organochlorine pesticides in Ganges River dolphins, Environ. Toxicol. Chem., 18: 1511–1520. Sergeant, D.E., 1980, Levels of Mercury and Organochlorine Residues in Tissues of Sea Mammals from the St. Lawrence Estuary, International Council for the Exploration of the Sea, Doc. CM/E:55, 10. Severinsen, T., Skaare, J.U., and Lydersen, C., 2000, Spatial distribution of persistent organochlorines in ringed seal (Phoca hispida) blubber, Mar. Environ. Res., 49: 291–302. Shlosberg, A., Bellaiche, M., Regev, S., Gal, R., Brizzi, M., Hanji, V., Zaidel, L., and Nyska, A., 1997, Lead toxicosis in a captive bottlenose dolphin (Tursiops truncatus) consequent to ingestion of air gun pellets, J. Wildl. Dis., 33: 135–139. Siebert, U., Joiris, C., Holsbeek, L., Benke, H., Failing, K., Frese, K., and Petzinger, E., 1999, Potential relation between mercury concentrations and necropsy findings in cetaceans from German waters of the North and Baltic Seas, Mar. Pollut. Bull., 38: 285–295. Simmonds, M.P., and Mayer, S.J., 1997, An evaluation of environmental and other factors in some recent marine mammal mortalities in Europe: Implications for conservation and management, Environ. Rev., 5: 89–98. Simmonds, M.P., Johnston, P.A., and French, M.C., 1993, Organochlorine and mercury contamination in United Kingdom seals, Vet. Rec., 132: 291–295. Simmonds, M.P., Johnston, P.A., French, M.C., Reeve, R., and Hutchinson, J.D., 1994, Organochlorines and mercury in pilot whale blubber consumed by Faroe Islanders, Sci. Total Environ., 149: 97–111. Skaare, J.U., 2000, Organochlorines in polar bear at Svalbard—Occurrence, levels, and effects, presented at Workshop on Persistent Organic Pollutants (POPs) in the Arctic: Human Health and Environmental Concerns, Rovaniemi, Finland, January 18–20, 2000, Abstr. 3. Smith, T.G., and Armstrong, F.A.J., 1975, Mercury in seals, terrestrial carnivores, and principal food items of the Inuit, from Holman, NWT, J. Fish. Res. Board Can., 32: 795–801. Sobotka, T.J., Brown, R., Quander, D.Y., Jackson, R., Smith, M., Long, S.A., Barton, C.N., Rountree, R.L., Hall, S., Eilers, P., Johannessen, J.N., and Scallet, A.C., 1996, Domoic acid: Neurobehavorial and neurohistological effects of low-dose exposure in adult rats, Neurotoxicol. Teratol., 18: 659–670. Spraker, T.R., Lowry, L.F., and Frost, K.J., 1994, Gross necropsy and histopathologic lesions found in harbor seals, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 281–312. Stern, G.A., Muir, D.C.G., Grift, N.P., Dewailly, E., Bidleman, T.F., and Wallace, M.D., 1992, Isolation and identification of two major recalcitrant toxaphene congeners, Environ. Sci. Technol., 26: 1838– 1840. Storelli, M.M., Zizzo, N., and Marcotrigiano, G. O., 1999, Heavy metals and methylmercury in tissues of Risso’s dolphin (Grampus griseus) and Cuvier’s beaked whale (Ziphius cavirostris) stranded in Italy (South Adriatic Sea), Bull. Environ. Contam. Toxicol., 63: 703–710. Strain, S.M., and Tasker, R.A.R., 1991, Hippocampal damage produced by systemic injections of domoic acid in mice, Neuroscience, 44: 343–352. Strandberg, B., Strandberg, L., Bergqvist, P.-A., Falandysz, J., and Rappe, C., 1998, Concentrations and biomagnification of 17 chlordane compounds and other organochlorines in harbour porpoise (Phocoena phocoena) and herring from the southern Baltic Sea, Chemosphere, 37: 2513–2523.
0839_frame_C22.fm Page 518 Tuesday, May 22, 2001 11:51 AM
518
CRC Handbook of Marine Mammal Medicine
Subramanian, A., Tanabe, S., Tatsukawa, R., Saito, S., and Miyazaki, N., 1987, Reduction in the testosterone levels by PCBs and DDE in Dall’s porpoises of northwestern North Pacific, Mar. Pollut. Bull., 18: 643–646. Szefer, P., Szefer, K., Pempkowiak, J., Skwarzec, B., Bojanowski, R., and Holm, E., 1994, Distribution and coassociations of select metals in seals of the Antarctic, Environ. Pollut., 83: 341–349. Tanabe, S., 1988, PCB problems in the future: Foresight from current knowledge, Environ. Pollut., 50: 5–28. Tanabe, S., 1999, Butyltin contamination in marine mammals — a review, Mar. Pollut. Bull., 39: 62–72. Tanabe, S., Tatsukawa, R., Tanaka, H., Maruyama, K., Miyazaki, N., and Fujiyama, T., 1981, Distribution and total burdens of chlorinated hydrocarbons in bodies of striped dolphins (Stenella coeruleoalba), Agric. Biol. Chem., 45: 2569–2578. Tanabe, S., Kannan, N., Subramanian, A., Watanabe, S., and Tatsukawa, R., 1987, Highly toxic coplanar PCBs: Occurrence, source, persistency and toxic implications to wildlife and humans, Environ. Pollut., 47: 147–163. Tanabe, S., Watanabe, S., Kan, H., and Tatsukawa, R., 1988, Capacity and mode of PCB metabolism in small cetaceans, Mar. Mammal Sci., 4: 103–124. Tanabe, S., Kumaran, P., Iwata, H., Tatsukawa, R., and Miyazaki, N., 1996, Enantiomeric ratios of x-hexachlorocyclohexane in blubber of small cetaceans, Mar. Pollut. Bull., 32: 27–31. Tanabe, S., Prudente, M., Mizuno, T., Hasegawa, J., Iwata, H., and Miyazaki, N., 1998, Butyltin contamination in marine mammals from North Pacific and Asian coastal waters, Environ. Sci. Technol., 32: 193–198. Teitelbaum, J.S., Zatorre, R.J., Carpenter, S., Gendrom, D., Evans, A.C., Gjedde, A., and Cashman, N.R., 1990, Neurologic sequelae of domoic acid intoxication due to the ingestion of contaminated mussels, N. Engl. J. Med., 322: 1781–1787. Tilbury, K.L., Adams, N.G., Krone, C.A., Meador, J.P., Early, G., and Varanasi, U., 1999, Organochlorines in stranded pilot whales (Globicephala melaena) from the coast of Massachusetts, Arch. Environ. Toxicol., 37: 125–134. Tillander, M., Miettinen, J.M., and Koivisto, I., 1972, Excretion rate of methylmercury in the seal, Marine Pollution and Sea Life, Ruivo, M. (Ed.), Fishing News (Books) Ltd., Surrey, U.K., 303–305. Tohyama, C., Himeno, S.I., Watanabe, C., Suzuki, T., and Morita, M., 1986, The relationship of the increased level of metallothionein with heavy metal levels in the tissue of the harbor seal (Phoca vitulina), Ecotoxicol. Environ. Saf., 12: 85–94. Tolley, K.A., and Heldal, H.E., in press, Inferring ecological stock structure from regional differences in radioactive caesium in harbour porpoises (Phocoena phocoena), Mar. Ecol. Trainer, V.L., and Baden, D.G., 1999, High affinity binding of red tide neurotoxins to marine mammal brain, Aquat. Toxicol., 46: 139–148. Troisi, G.M., and Mason, C.F., 1997, Cytochromes P450, P420 and mixed-function oxidases as biomarkers of polychlorinated biphenyl (PCB) exposure in harbour seals (Phoca vitulina), Chemosphere, 35: 1933–1946. Troisi, G.M., Haraguchi, K., and Simmonds, M.P., 1997, PCB-methyl sulphones in marine mammal tissues, with particular reference to epizootic victims, Eur. Res. Cetaceans, 11: 286–289. Troisi, G.M., Haraguchi, K., Simmonds, M.P., and Mason, C.F., 1998, Methyl sulphone metabolites of polychlorinated biphenyls (PCBs) in cetaceans from the Irish and Aegean Seas, Arch. Environ. Contam. Toxicol., 35: 121–128. van de Ven, W.S.M., Koeman, J.H., and Svenson, A., 1979, Mercury and selenium in wild and experimental seals, Chemosphere, 8: 539–555. Van Dolah, F.M., Leighfield, T.A., Haynes, B.L., Hampson, D.R., and Ramsdell, J.S., 1997, A microplate receptor assay for the amnesic shellfish poisoning toxin, domoic acid, utilizing a cloned glutamate receptor, Anal. Biochem., 245: 102–105.
0839_frame_C22.fm Page 519 Tuesday, May 22, 2001 11:51 AM
Toxicology
519
van Scheppingen, W.B., Verhoeven, A.J.I.M., Mulder, P., Addink, M.J., and Smeenk, C., 1996, Polychlorinated biphenyls, dibenzo-p-dioxins, and dibenzofurans in harbor porpoises (Phocoena phocoena) stranded on the Dutch coast between 1990 and 1993, Arch. Environ. Contam. Toxicol., 30: 492–502. Varanasi, U., Stein, J.E., Tilbury, K.L., Meador, J.P., Sloan, C.A., Clark, R.C., and Chan S.-L., 1994, Chemical contaminants in gray whales stranded along the west coast of North America, Sci. Total Environ., 145: 29–53. Vetter, W., Weichbrodt, M., Scholz, E., Luckas, B., and Oelschläger, H., 1999, Levels of organochlorines (DDT, PCBs, toxaphene, chlordane, dieldrin, and HCHs) in blubber of South African fur seals (Arctocephalus pusillus pusillus) from Cape Cross/Namibia, Mar. Pollut. Bull., 38: 830–836. Vos, J.G., Bossart, G., and Fournier, M. (Eds.), in press, Toxicology of Marine Mammals, Taylor & Francis, New York. Wade, T.L., Chambers, L., Gardinali, P.R., Sericano, J.L., Jackson, T.J., Tarpley, R.J., and Suydam, R., 1997, Toxaphene, PCB, DDT, and chlordane analyses of beluga whale blubber, Chemosphere, 34: 1351–1357. Wagemann, R., 1989, Comparison of heavy metals in two groups of ringed seals (Phoca hispida) from the Canadian Arctic, Can. J. Fish. Aquat. Sci., 46: 1558–1563. Wagemann, R., Snow, N.B., Lutz, A., and Scott, D.P., 1983, Heavy metals in tissues and organs of the narwhal (Monodon monoceros), Can. J. Fish. Aquat. Sci., 40: 206–214. Wagemann, R., Hunt, R., and Klaverkamp, J.F., 1984, Subcellular distribution of heavy metals in liver and kidney of a narwhal whale (Monodon monoceros): An evaluation for the presence of metallothionein, Comp. Biochem. Physiol. C, 78: 301–307. Wagemann, R., Stewart, R.E.A., Beland, P., and Desjardins, C., 1990, Heavy metals and selenium in tissues of beluga whales, Delphinapterus leucas, from the Canadian Arctic and the St. Lawrence estuary, Can. Bull. Fish. Aquat. Sci., 224: 191–206. Walker, W., Risebrough, R.W., Jarman, W.M., de Lappe, B.W., Tefft, J.A., and DeLong, R.L., 1989, Identification of tris(chlorophenyl)methanol in blubber of harbor seals from Puget Sound, Chemosphere, 18: 1799–1804. Warburton, J., and Seagers, D.J., 1993, Metal concentrations in liver and kidney tissues of Pacific walrus: Continuation of a baseline study, Technical Report R7/MMM 93-1, Marine Mammals Management, U.S. Fish and Wildlife Science, Anchorage, AK 99508, 23. Watanabe, S., Shimada, T., Nakamura, S., Nishiyama, N., Yamashita, N., Tanabe, S., and Tatsukawa, R., 1989, Specific profile of liver microsomal cytochrome P-450 in dolphin and whales, Mar. Environ. Res., 27: 51–65. Watanabe, M., Tanabe, S., Miyazaki, N., Petrov, E.A., and Jarman, W.M., 1999, Contamination of tris(4chlorophenyl) methane and tris(4-chlorophenyl) methanol in marine mammals from Russia and Japan: Body distribution, bioaccumulation and contamination status, Mar. Pollut. Bull., 39: 393–398. Weisbrod, A.V., Shea, D., Moore, M.J., and Stegeman, J.J., 2000a, Organochlorine exposure and bioaccumulation in the endangered Northwest Atlantic right whale (Eubalaena glacialis) population, Environ. Toxicol. Chem., 19: 654–666. Weisbrod, A.V., Shea, D., Moore, M.J., and Stegeman, J.J., 2000b, Bioaccumulation patterns of polychlorinated biphenyls and chlorinated pesticides in Northwest Atlantic pilot whales, Environ. Toxicol. Chem., 19: 667–677. Wells, D.E., and Echarri, I., 1992, Determination of individual chlorobiphenyls (CBs), including nonortho, and mono-ortho chloro substituted CBs in marine mammals from Scottish waters, Int. J. Environ. Anal. Chem., 47: 75–97. Wells, D., Campbell, L.A., Ross, H.M., Thompson, P.M., and Lockyer, C.H., 1994, Organochlorine residues in harbour porpoise and bottlenose dolphins stranded on the coast of Scotland, 1988–1991, Sci. Total Environ., 151: 77–99.
0839_frame_C22.fm Page 520 Tuesday, May 22, 2001 11:51 AM
520
CRC Handbook of Marine Mammal Medicine
White, R.D., Hahn, M.E., Lockhart, W.L., and Stegeman, J.J., 1994, Catalytic and immunochemical characterization of hepatic microsomal cytochromes P450 in beluga whale (Delphinapterus leucas), Toxicol. Appl. Pharmacol., 126: 45–57. Wiig, O., 1998, Survival and reproductive rates for polar bears at Svalbard, Ursus, 10: 25–32. Williams, T.M., Kastelein, R.A., Davis, R.W., and Thomas, J.A., 1988, The effects of oil contamination and cleaning on sea otters, I. Thermoregulatory implications based on pelt studies, Can. J. Zool., 66: 2776–2781. Williams, T.M., Antonelis, G.A., and Balke, J., 1994, Health evaluation, rehabilitation, and release of oiled harbor seal pups, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 227–242. Woodley, T., Brown, M., Kraus, S., and Gaskin, D., 1991, Organochlorine levels in North Atlantic right whale (Eubalaena glacialis) blubber, Arch. Environ. Contam. Toxicol., 21: 141–145. Work, T.M., Barr, B., Beale, A.M., Fritz, L., Quilliam, M.A., and Wright, J.L.C., 1993, Epidemiology of domoic acid poisoning in brown pelicans (Pelecanus occidentalis) and Brandt’s cormorants (Phalacrocorax penicillatus) in California, J. Zoo Wildl. Med., 24: 54–62. Woshner, V.M., 2000, Concentrations and Interactions of Selected Elements in Tissues of Four Marine Mammal Species Harvested by Inuit Hunters in Arctic Alaska, with an Intensive Histologic Assessment, Emphasizing the Beluga Whale, Ph.D. thesis, University of Illinois, Urbana, 264 pp. Yamamoto, J.T., Donohoe, R.M., Fry, D.M, Golub, M.S., and Donald, J.M., 1996, Environmental estrogens: Implications for reproduction in wildlife, in Noninfectious Diseases of Wildlife, 2nd ed., Fairbrother, A., Locke, L., and Hoff, G. (Eds.), Iowa State University Press, Ames, 61–70. Yang, F., Chau, Y.K., and Maguire, R.J., 1998, Occurrence of butyltin compounds in beluga whales, Appl. Organomet. Chem., 12: 651–656. Zakharov, V.M., and Yablokov, A.V., 1990, Skull asymmetry in the Baltic grey seal: Effects of environmental pollution, Ambio, 19: 266–269. Zhu, J., and Norstrom, R.J., 1993, Identification of polychlorocamphenes (PCCs) in the polar bear (Ursus maritimus) food chain, Chemosphere, 27: 1923–1936. Zook, D.R., Buser, H.-R., Bergqvist, P.-A., Rappe, C., and Olsson, M., 1992, Detection of tris(chlorophenyl) methane and tris(4-chlorophenyl) methanol in ringed seal (Phoca hispida) from the Baltic Sea, Ambio, 21: 557–560.
0839_frame_C23 Page 521 Tuesday, May 22, 2001 11:52 AM
23 Noninfectious Diseases Frances M. D. Gulland, Linda J. Lowenstine, and Terry R. Spraker
Introduction This chapter reviews a range of noninfectious diseases documented in marine mammals, excluding those known to be associated with toxicoses and nutritional deficiencies (see Chapter 22, Toxicology; Chapter 36, Nutrition). Most of these noninfectious conditions have been detected as a result of necropsies on individual stranded, harvested, or captive animals, rather than through systematic studies on wild populations. The literature consequently contains a multitude of scattered case reports with little information on the impact of these noninfectious conditions on wild marine mammal populations. The purpose of this chapter is to bring these individual reports together and provide an overview of the current literature available on noninfectious diseases in marine mammals. For ease of reference, separate sections of this chapter describe lesions identified as congenital defects, neoplastic lesions, and those associated with trauma, while other, miscellaneous lesions are described according to the organ system in which they were identified.
Congenital Defects Congenital defects are abnormalities of structure or function present at birth, although the defects may not be expressed or detected until later in life. The majority of congenital defects reported in cetaceans were actually detected in fetuses in utero during necropsies of pregnant animals. This is especially true of whales examined during the historical periods of commercial whaling. In contrast, most reports of congenital anomalies in pinnipeds are a result of necropsies performed on neonatal pups at rookeries and on stranded individuals postweaning. Most congenital anomalies are detected during post-mortem examination; therefore, most are descriptions of gross anatomical defects. Although a syndrome of “ill-thrift” is recognized in a number of species, the importance of microscopic or metabolic congenital defects that may cause such signs has yet to be determined. The anatomical development of marine mammals from fetus to mature adult is poorly documented compared with that of humans and domestic species. As a result, some structures may have been described as defects, even though they were part of the normal development of that species. For example, the presence of a patent ductus arteriosus in a young terrestrial animal after birth is considered a congenital defect compromising circulation, whereas it is common in marine mammals, and species differences in closure time still need to be established (Slijper, 1962; Leipold, 1980; Banish and Gilmartin, 1992). In addition, lack of history for an animal from a wild population can complicate interpretation
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
521
0839_frame_C23 Page 522 Tuesday, May 22, 2001 11:52 AM
522
CRC Handbook of Marine Mammal Medicine
of findings. Megaesophagus and hiatal hernias have been reported in harbor seal (Phoca vitulina) pups (Stroud, 1978; Harms et al., 1994) and a harbor porpoise (Phocoena phocoena) (Stephen, 1993) and deformity of the parietal bones in an elephant seal (Mirounga angustirostris) (Huey, 1925). However, it is unclear whether these lesions were congenital or acquired in early life. Similarly, lack of dentine has been observed in a bottlenose dolphin (Tursiops truncatus) (Brooks and Anderson, 1998), but whether this was due to the heritable defect amelogenesis imperfecta, to severe infection, or to nutritional deficiency in early life is unknown since there is no clinical history for this animal. Neither the prevalence nor the etiology of any congenital defects in marine mammals has been determined. The congenital defects reported in marine mammals are summarized in Table 1.
Neoplasia The number of reports of neoplasia in marine mammals has increased dramatically over the last 20 years. This may not be a real increase in incidence, however, but rather a reflection of the increased numbers of animals examined by pathologists. Geraci and co-workers critically and comprehensively reviewed the tumors reported as of 1987 (Geraci et al., 1987). Since then, a number of other tumors have been reported, especially in belugas (Delphinapterus leucas) from the St. Lawrence estuary and in California sea lions (Zalophus californianus) (Table 2). Although species differences in tumor prevalence may be due in part to the extent of the species interaction with humans (in zoological collections, likelihood of stranding and examination, hunting), they may also reflect different etiologies. Physical, chemical, and infectious agents have all been associated with neoplasms in other species. The effects of these carcinogens are further modulated by age, hormones, and genetics. As is the case for the majority of spontaneous tumors in other species, the etiology of most tumors in marine mammals is unknown. Certainly, marine mammals are exposed to many types of potential carcinogens, including potentially oncogenic viruses, as well as radioactive nucleotides and xenobiotic contaminants that accumulate in their tissues (Mossner and Ballschmiter, 1997; Watson et al., 1999). Although much attention has focused on the potential effects of toxins (see Chapter 22, Toxicology), infectious agents have been identified in papillomas in manatees (Trichechus manatus) (Bossart et al., 1998), gastric papillomas in belugas (De Guise et al., 1994b), and in squamous cell carcinomas of bottlenose dolphins (Bossart, in press). The epidemiology of lymphosarcoma in harbor seals suggests an infectious agent may be involved in pathogenesis, but none has yet been identified (Osborn et al., 1988). A herpesvirus has been observed in tumors in California sea lions, but its role is unknown (Gulland et al., 1996; Lipscomb et al., 2000). Tumors reported in marine mammals are listed in Table 2. In compiling this table, tumors in individual animals that were reported in more than one reference are only referred to once. As some of the lesions in the literature are questionable, the definitions of tumors used by Geraci et al. (1987) were used, and lesions they considered as probably not neoplastic have not been included.
Trauma Intraspecific Trauma Parallel superficial skin lesions (“rake marks”) due to intraspecific interactions are commonly observed in toothed cetaceans (Greenwood et al., 1974). More recently, severe traumatic lesions characterized by bilateral rib fractures and subcutaneous and pulmonary hemorrhages have been reported in bottlenose dolphin calves killed by adult conspecifics (Patterson et al., 1998).
0839_frame_C23 Page 523 Tuesday, May 22, 2001 11:52 AM
523
Noninfectious Diseases
TABLE 1 Congenital Defects Reported in Marine Mammals Species
Defect
Reference
Tursiops truncatus (bottlenose dolphin)
Ventricular septal defect Polydactyly Transposition of pulmonary artery and aorta Hermaphroditism Rudimentary hind limbs Conjoined twins
Gray and Conklin, 1974 Watson et al., 1994 Gray and Conklin, 1974
Stenella coeruleoalba (striped dolphin)
Delphinus delphis (common dolphin) Globicephala malaena (pilot whale) Megaptera novaeangliae (humpback whale) Delphinapterus leucas (beluga whale) Physeter macrocephalus (sperm whale) Balaena mysticetus (bowhead whale) Balaenoptera borealis (sei whale) Balaenoptera acutorostrata (minke whale) Balaenoptera physalus (fin whale) Fossil mysticete Trichechus manatus (manatee) Halichoerus grypus (gray seal) Phoca vitulina (harbor seal)
Mirounga leonina (southern elephant seal) Mirounga angustirostris (northern elephant seal)
Callorhinus ursinus (northern fur seal)
Polycystic kidney
Nishiwaki, 1953 Ohsumi, 1965 Kawamura and Kashita, 1971; Kamiya and Miyazaki, 1974 Howard, 1983
Block vertebrae
Cowan, 1966
Conjoined twins Rudimentary hind limbs Hermaphroditism
Kamiya et al., 1981 Andrews, 1921 De Guise et al., 1994c
Rudimentary hind limbs Pseudohermaphroditism
Ogawa and Kamiya, 1957; Nemoto, 1963 Tarpley et al., 1995
Conjoined twins
Kawamura, 1990
Conjoined twins
Zinchenko and Ivashin, 1987
Pseudohermaphroditism
Bannister, 1962
Spina bifida Ectrodactyly
Fordyce and Watson, 1998 Watson and Bonde, 1986
Flattened trachea
Baker, 1989a
Cleft palate Ectrodactyly Abnormal tooth number
Suzuki et al., 1992 Tarasoff and Pierard, 1970 Suzuki et al., 1990; Colyer, 1936 King, 1964 Spraker et al., 1994 Csordas, 1966 Laws, 1953 Griner, 1983; Trupkiewicz et al., 1997 Trupkiewicz et al., 1997
Alopecia, dental aplasia Penile deviation Penile malformation Conjoined twins Hydrocephalus Right ventricular hypoplasia and overriding aorta Ventricular septal defect Angiomatosis Pulmonary dysplasia Hydronephrosis Polydactyly Hypoplasia, skull Ocular and skull deformity Brachycephalia
Spraker, in press
(Continued)
0839_frame_C23 Page 524 Tuesday, May 22, 2001 11:52 AM
524
CRC Handbook of Marine Mammal Medicine
TABLE 1 Congenital Defects Reported in Marine Mammals (continued) Species
Monachus schauinslandi (Hawaiian monk seal)
Odobenus rosmarus (walrus) Zalophus californianus (California sea lion)
Ursus maritimus (polar bear)
Defect Cerebellar hypoplasia Hydrocephalus Agenesis, foreflipper Bilateral hypoplasia, forelimbs Persistent truncus arteriosus Agenesis, right ventricle Umbilical herniation with evisceration Hypoplasia, left atrium and ventricle, with ventricular septal defect Palatoschisis, hypoplasia of lung, chest, and limbs Hypoplasia, diaphragm Agenesis, tail Agenesis, right forelimb and right kidney Scoliosis Diverticulum, fundus Non-union pylorus to duodenum with stenosis of common bile duct Atresia, bile duct Atresia, anus and vulva Aplasia, segmental, small intestine Renal cysts Horseshoe kidney Hermaphroditism Partial albinism Ganglioneuroblastoma Microphthalmia; hypoplasia of phalanges; segmental aplasia of the ileum (all in single animal) Pseudopersistent urachus Polycystic kidneys Unilateral renal aplasia/ hypoplasia Fusion of splenic and hepatic capsule Pseudohermaphroditism Agenesis, radius
Reference
Spraker and Aguirre, pers. comm.
Cornell et al., 1975 Howard, 1983 Sweeney and Gilmartin, 1974; Howard, 1983 Sweeney and Gilmartin, 1974 Wiig et al., 1998 Lanthier et al., 1998
Fractured lower jaws have been seen in sperm whales (Physeter macrocephalus) and have been attributed variously to intraspecific aggression or collisions with boats (Slijper, 1962). Intraspecific aggression is more commonly observed in pinniped rookeries and may occur during territorial disputes and mating. Blunt traumatic lesions, characterized by bone fractures, hemorrhages, ruptured diaphragms, and hepatic fissures are commonly observed in pinniped pups on rookeries as a consequence of crushing by adults (Reiter et al., 1978; Banish and Gilmartin,
0839_frame_C23 Page 525 Tuesday, May 22, 2001 11:52 AM
525
Noninfectious Diseases
TABLE 2 Neoplastic Lesions Reported in Marine Mammals Species
Tumor Type
Organ Site
Reference
Balaena mysticetus (bowhead whale) Balaenoptera borealis (sei whale) Balaenoptera physalus (fin whale)
Lipoma
Liver
Migaki and Albert, 1982
Melanoma
Lip
Uys and Best, 1966
Lipoma Fibroma (papilloma?)
Dorsal muscle Skin Tongue Ovary Ovary
Cockrill, 1960 Stolk, 1952 Stolk, 1952 Rewell and Willis, 1949 Stolk, 1950
Brain Mediastinum Tongue Stomach Intestine Ovary Ovary Uterus Brain Tongue Urinary bladder Stomach Intestine
Pilleri, 1968 Rewell and Willis, 1950 Rewell and Willis, 1949 Cockrill, 1960 Cockrill, 1960 Rewell and Willis, 1949 Rewell and Willis, 1949 Stolk, 1950 Pilleri, 1966 Stolk, 1952 Martineau et al., 1985
Balaenoptera musculus (blue whale)
Megaptera novaeangliae (humpback whale) Delphinapterus leucas (beluga)
Granulosa cell tumor Carcinoma (? granulosa cell tumor) Neurofibroma Ganglioma Papilloma Lipoma Granulosa cell tumor Cystadenoma Fibromyoma Lipoma Papilloma Transitional cell carcinoma Adenocarcinoma
Carcinoma Papilloma Hemangioma
Monodon monoceros (narwhal) Mesoplodon densirostris (Blainville’s beaked whale) Globicephala macrorhynchus (short-finned pilot whale) Globicephala malaena (long-finned pilot whale)
Mammary gland Uterus Salivary gland Liver Stomach
De Guise et al., 1994a De Guise et al., 1994a; Martineau et al., 1995 De Guise et al., 1994a Lair et al., 1998 Girard et al., 1991 De Guise et al., 1994a Martineau et al., 1988; De Guise et al., 1994b De Guise et al., 1994a Martineau et al., 1988
Chondroma Pheochromocytoma Granulosa cell tumor
Penis Urinary bladder Lung Adrenal Ovary
Fibroma Lipoma Adenoma Papilloma
Spleen Lung Thyroid Skin
De Guise et al., 1994a De Guise et al., 1994a Martineau et al., 1988; De Guise et al., 1994a Martineau et al., 1985 Martineau et al., 1988 De Guise et al., 1994a Geraci et al., 1987
Fibroma
Vagina
Flom et al., 1980
Granulosa cell tumor Leiomyoma Leiomyoma
Ovary Uterus Uterus
Benirschke and Marsh, 1984 Bossart et al., 1991 Cowan, 1966 (Continued)
0839_frame_C23 Page 526 Tuesday, May 22, 2001 11:52 AM
526
CRC Handbook of Marine Mammal Medicine
TABLE 2 Neoplastic Lesions Reported in Marine Mammals (continued) Species
Tumor Type
Organ Site
Reference
Physeter macrocephalus (sperm whale)
Fibroma
Uterus Lower jaw Penis Liver
Uys and Best, 1966 Stolk, 1952; Clarke, 1956 Lambertsen et al., 1987 Stolk, 1953
Penis, skin
Taylor and Greenwood, 1974 Geraci et al., 1987; Bossart et al., 1996 Baker and Martin, 1992 Taylor and Greenwood, in Landy, 1980 Geraci et al., 1987 Geraci et al., 1987 Geraci et al., 1987 Geraci et al., 1987 Geraci et al., 1987 Howard et al., 1983
Orca orcinus (killer whale)
Papilloma Hemangioma (or sarcoma?) Papilloma
Skin Phocoena phocoena (harbor porpoise)
Adenocarcinoma Papilloma
Lagenorhyncus acutus (Atlantic white-sided dolphin)
Adenoma Leiomyoma Papilloma
Lagenorhyncus obliquidens (Pacific white-sided dolphin)
Fibroma
Squamous cell carcinoma Lymphosarcoma
Unknown Inia geoffrensis (boto) Trichechus manatus (manatee) Pagophilus groenlandicus (harp seal) Halichoerus grypus (gray seal) Mirounga leonina (southern elephant seal)
Skin Adrenal Intestine Penis Tongue Gingiva
Lymphoma Lymphoma Leiomyoma Leydig cell tumor Adenoma Reticuloendotheliosis
Skin Spleen, lymph nodes Liver Multiple Multiple Multiple Stomach Testis Kidney Lungs
Lymphosarcoma
Spleen
Lymphoma Squamous cell carcinoma Unknown Carcinoma
Multiple Oral mucosa Testis Pancreas
Fibroma Squamous cell carcinoma
Uterus Lung
Geraci et al., 1987 Howard et al., 1983 Bossart et al., 1997 Bossart et al., 1997 Cowan et al., 1986 Cowan et al., 1986 Migaki et al., 1978 Landy, 1980; Ridgway, in Geraci et al., 1987 Taylor and Greenwood, in Landy, 1980 Bossart et al., 1997 Bossart, in press Mawdesley-Thomas, 1974 Taylor and Greenwood, in Landy, 1980 Rewell and Willis, 1949 Geraci et al., 1987
Papilloma
Skin
Bossart et al., 1998
Lymphosarcoma
Lymph nodes
Migaki, in Landy, 1980
Leiomyoma
Uterus
Squamous cell carcinoma Granulosa cell tumor
Ovary
Mawdesley-Thomas and Bonner, 1971 Bergman, 1997 Mawdesley-Thomas, 1971
Leukemia Stenella frontalis (Atlantic spotted dolphin) Delphinus delphis (common dolphin) Tursiops truncatus (bottlenose dolphin)
Unknown Penis
Howard et al., 1983 Howard et al., 1983
0839_frame_C23 Page 527 Tuesday, May 22, 2001 11:52 AM
527
Noninfectious Diseases
TABLE 2 Neoplastic Lesions Reported in Marine Mammals (continued) Species
Tumor Type
Organ Site
Reference
Phoca vitulina (harbor seal)
Lymphosarcoma
Lymph nodes
Eumetopias jubatus (Steller sea lion)
Fibroleiomyoma Adenocarcinoma Skin tumors, probably fibromas Transitional cell carcinoma
Uterus Lung Eyelids
Larsen, 1962; Griner, 1971; Stroud and Stevens, 1980; Osborn et al., 1988 Morgan et al., 1996 Sato et al., 1998 Spraker, unpubl. data
Zalophus californianus (California sea lion)
Adenocarcinoma
Urinary bladder Kidney Multiple
Carcinoma
Multiple Liver Bile duct Lungs
Squamous cell carcinoma
Fibrosarcoma Lymphosarcoma Granulosa cell tumor Melanoma Adenoma
Bladder Pharynx Gingiva Skin Perineum Metastatic Mammary gland Lymph nodes Ovary Eye, brain Ovary Pancreas
Pituitary Adrenal Lipoma
Papilloma Fibroma
Esophagus Mammary gland Vagina Skin
Migaki, in Landy, 1980
Fox, 1941, in Howard et al., 1983; Griner, 1983; Simpson and Gardner, 1972; Stroud and Roffe, 1979; Brown et al., 1980; Simpson and Ridgway, in Landy, 1980; Howard et al., 1983 Griner, 1983; Gulland et al., 1996 Acevedo-Whitehouse et al., 1999 Schroeder et al., 1973 Landy, 1980; Howard et al., 1983 Sweeney, 1974 Griner, 1983 Bossart, 1990 Snyder, in Landy, 1980; Anderson et al., 1990 Griner, in Landy, 1980 Joseph et al., 1986 Snyder, in Landy, 1980 Taylor and Greenwood, in Landy, 1980 Howard et al., 1983 Griner, 1983 Howard et al., 1983 Griner, 1983; Landy 1980; Moore and Stackhouse, 1978 Landy, 1980 Griner, 1983; Sweeney, 1973; Landy, 1980 Griner, in Landy, 1980 Howard et al., 1983 Howard et al., 1983 Simpson and Gardner, 1972 (Continued)
0839_frame_C23 Page 528 Tuesday, May 22, 2001 11:52 AM
528
CRC Handbook of Marine Mammal Medicine
TABLE 2 Neoplastic Lesions Reported in Marine Mammals (continued) Species
Callorhinus ursinus (northern fur seal)
Odobenus rosmarus (walrus) Enhydra lutra (sea otter)
Tumor Type
Organ Site
Reference
Hemangioma Nephroblastoma Leiomyoma
Vagina Kidney Uterus
Ganglioneuroblastoma Fibroma Carcinoma Granulosa cell tumor Squamous cell carcinoma
Heart, stomach Skin Adrenal gland Ovary Vagina, cervix
Howard et al., 1983 Sweeney, 1974 Appleby, in Landy, 1980; Howard et al., 1983; Bossart, 1990 Spraker, in press
Lymphosarcoma Fibrosarcoma Osteosarcoma
Prepuce, penis Gingiva Lymph nodes Kidney Bone
Leukemia Leiomyoma
Multicentric Uterus
Carcinoma Pheochromocytoma Seminoma
Bile duct Adrenal Testis
Griner, 1983; Howard et al., 1983 Griner, 1983 Bossart, 1990 Stedham et al., 1977 Brown et al., 1975 Pierard et al., 1977 Larsen, 1962 Stetzer et al., 1981; Williams and Pullet, 1981 Stetzer et al., 1981 Stetzer et al., 1981 Reimer and Lipscomb, 1998
1992; Spraker, pers. obs.). Rare forms of severe blunt trauma to the chest result in rupture of the heart, aorta, stomach, and small intestine (Spraker, in press). Bites, resulting in puncture wounds associated with hemorrhage, are also common in pinnipeds of all ages (Greenwood et al., 1974; Reiter et al., 1978). These may become infected, resulting in abscesses, fascitis, and occasionally osteomyelitis, polyarthritis, and encephalitis. There is a common syndrome in northern fur seal (Callorhinus ursinus) pups that is characterized by cellulitis over the skull with occasionally suppurative meningitis. This occurs when pups are bitten on the head as they attempt to nurse a female other than their own mother (Spraker, in press). Displaced sexual behavior may also result in traumatic lesions to pinniped females and pups, including edema and hemorrhage to the dorsum, pulmonary hemorrhages, fractured or luxated vertebrae, and metritis associated with trauma (Reiter et al., 1978; Banish and Gilmartin, 1992). Sexual behavior in sea otters (Enhydra lutra) commonly results in injury to the nose of the female and fracture of the os penis in males (Morejohn et al., 1975; Tuomi, pers. comm.). Mating wounds in sea otters are often severe enough to cause death (see Chapter 44, Sea Otters). Cannibalism has been observed in gray seals (Halichoerus grypus) (Bedard et al., 1993), southern elephant seals (Mirounga leonina) (Campagna, in Wilkinson et al., 2000), and Hooker’s sea lions (Phocarctos hookeri) (Wilkinson et al., 2000).
Interspecific Trauma Acute traumatic lesions resulting from aggression between marine mammal species have been documented in a number of species. Parallel linear skin wounds commencing as three-cornered tears, subcutaneous bruising, shearing of the blubber from the subcutis, rib fractures occasionally associated with puncture of the underlying lung and pneumothorax, and dislocation of
0839_frame_C23 Page 529 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
529
thoracic vertebrae have been described in juvenile harbor porpoises as a result of violent interactions with bottlenose dolphins (Ross and Wilson, 1996; Jepson and Baker, 1998). Parallel scars on the skin of a captive killer whale (Orca orcinus) were attributed to teeth marks from dolphins with whom it was housed; similar scars were observed in a variety of odontocete cetaceans (Greenwood et al., 1974). Attacks by killer whales on various cetaceans result in lesions varying from parallel rake marks to death (George et al., 1994; George and Suydam, 1998). Skin wounds on sperm whales were attributed to killer whales, although they could have resulted from intraspecific aggression (Dufault and Whitehead, 1995). Healed pairs of puncture wounds have been observed in bowhead whales (Balaena mysticetus) that were attributed to attacks from walrus (Odobenus rosmarus). The reason for this assumption was that the spacing between the pair of lesions was equivalent to the distance between tusk points in mature walrus (Philo et al., 1993). The fractured tip of a narwhal (Monodon monoceros) tusk was once observed in the melon of a beluga that was foraging shortly before being harvested by Inuvialuit hunters (Orr and Harwood, 1998). Fractured skulls, proptosed eyeballs, subcutaneous hemorrhages, and puncture wounds in the soft tissue of the neck and head have been observed in female South American fur seals (Arctocephalus australis) and California sea lions dying as a result of interspecific sexual aggression by male southern (Otaria byroni) and Steller sea lions (Eumetopias jubatus), respectively (Miller, 1996). A number of sea lion species have been observed skinning and eating fur seal pups (Mattlin, 1978; Gentry and Johnson, 1981; Harcourt, 1993; Robinson et al., 1999). Wounds due to shark attacks are well described in a number of marine mammal species, with the species of shark and prey varying geographically (Kenyon, 1981; Alcorn and Kam, 1986; Corkeron et al., 1987a,b; Orams and Deakin, 1997). Cookie-cutter sharks (Isistius brasiliensis) use their lower teeth to remove plugs of flesh, leaving characteristic circular wounds that heal to form circular scars. These scars have been observed in cetaceans, elephant seals, and a Guadelupe fur seal (Arctocephalus townsendii) (Jones, 1971; LeBoeuf et al., 1987; Gallo-Reynoso and Figueroa-Carranza, 1992). Lesions due to other species of shark, such as the great white (Carcharodon carcharias), are typically an elliptical series or arc (of a radius varying with the size of the shark’s mouth) of deep puncture wounds or lacerations of varying lengths. Appendages may be amputated and there can be abrasions on bone and occasionally a tooth can be found in the lesion (Ames and Morejohn, 1980; Leboeuf et al., 1982). Vibrio carchariae has been isolated from shark bite wounds in humans, and is thought to result in cellulitis (Pavia et al., 1989; Klontz et al., 1993). A range of Vibrio spp. has been cultured from the mouths of sharks (Buck et al., 1984; Grimes et al., 1993), yet the significance of Vibrio spp. in shark bite wounds in marine mammals is unclear, as these organisms are often cultured from tissues of clinically normal animals. Although stingray spines have been reported embedded in the integument of a variety of species with no apparent ill effects (Castello, 1977), acute and chronic inflammatory lesions of internal organs have been observed in bottlenose dolphins as a result of penetration by stingray spines (Dasyatis spp.) (Jenkins and Cardeilhac, 1982; Walsh et al., 1988; McClellan et al., 1996; McFee et al., 1997). Pericarditis and pleuritis were reported in an Australian fur seal (Arctocephalus pusillus) following penetration of the esophagus and migration of the spine from a Urolophus paucimaculatus (Obendorf and Presidente, 1978). The death of a killer whale was attributed to penetration of the pharynx and cranial carotid rete by a stingray spine (Duignan et al., 2000). Healed lesions associated with embedded swordfish and marlin bills have been reported in bowhead whales (Philo et al., 1993). A Hawaiian monk seal (Monachus schauinslandi) pup died after swallowing an unidentified spiny-finned fish that lodged in the esophagus, causing esophageal necrosis and rupture, tracheal necrosis, and aspiration pneumonia (Banish and Gilmartin, 1992).
0839_frame_C23 Page 530 Tuesday, May 22, 2001 11:52 AM
530
CRC Handbook of Marine Mammal Medicine
Puncture wounds, hemorrhaging of tissues in the head and neck, and septicemia were described in harbor seal pups predated upon by coyotes (Steiger et al., 1989).
Anthropogenic Trauma Gross evidence of human-induced mortality in small cetaceans is reviewed by Read and Murray (2000). Entanglement of marine mammals in debris such as fishing nets and lines, packing bands, and rope is commonly observed in stranded dead animals (Heezen, 1957; Heyning and Lewis, 1990; Waring et al., 1990; Kuiken et al., 1994; Perrin et al., 1994). Apparently healthy freeliving animals with debris attached are also often observed (Fowler, 1986; Stewart and Yochem, 1987). Determining whether the death of a stranded animal resulted from entanglement and drowning or whether entanglement occurred after death can be difficult. Criteria for diagnosing death due to entanglement are reviewed by Kuiken (1994). Ante-mortem peripheral lesions are associated with varying degrees of hemorrhage, inflammation, and fibrosis. Lesions in the lung associated with drowning are inconsistent. When odontocetes drown, they rarely have water in their lungs, presumably because of their well-developed reflexes adapted for diving. Profound constriction of the bronchiolar smooth muscle resulting in obliteration of the lumen was observed in a bottlenose dolphin known to have drowned (Simpson and Gardner, 1972). However, this may have been due to rigor mortis, rather than to a specific response to drowning. The presence of diatoms in alveoli and femoral bone marrow is evidence of drowning in humans, and may be a useful sign in marine mammals that do not dry drown (Pollanen et al., 1997). Entanglement can also result in chronic injury rather than immediate death by drowning. Marine debris may surround parts of an animal’s body loosely at first, becoming tighter as the animal grows, or as drag on the gear causes it to tighten. Pinnipeds tend to retain gear around the head and neck (Fowler, 1986; Stewart and Yochem, 1987; Goldstein et al., 1999), whereas on cetaceans gear tends to move caudally, entangling flippers, flukes, or the peduncle. If the gear falls away, sigmoid or spiral scars may remain (Philo et al., 1992). Unusual cases have been observed in northern fur seals where monofilament nets have cut through the skull, humerus, and trachea (Spraker, in press). Boat collisions can also cause injury and mortality in marine mammals (Kraus, 1990; Philo et al., 1993), but distinguishing ante- and post-mortem collision can be difficult. Mandibular fractures observed in baleen whales have been attributed to boat strikes, but it is unclear how they affected health. A fracture of a bowhead whale’s mandible was not associated with weight loss (Philo et al., 1990), whereas a northern right whale (Eubalaena glacialis) with a similar fracture died after about 10 days (De Guise et al., 1999), and a gray whale (Eschrichtius robustus) with such a fracture was found dead, emaciated (Heyning, pers. comm.). Propeller injuries are most common in manatees (see Chapter 3, Manatee Case Study; Chapter 43, Manatees) although they are also observed in pinnipeds and cetaceans (Griner, 1983; Philo et al., 1993; Wells and Scott, 1997). Propeller-induced lesions are distinguished from shark bites as propeller wounds are usually situated on the dorsum (shark bites are usually ventral and caudal) and are parallel, sharp cuts of equal length that may slice cleanly through bone (shark bites usually have an elliptical pattern). Foreign body injuries associated with fishhooks, shotgun pellets, and bullets are common in pinnipeds on the West Coast of the United States (Stroud and Roffe, 1979; Griner, 1983; Goldstein et al., 1999). The extent of these lesions varies from little detectable impact on health (found as incidental lesions) to severe necrotizing lesions, gastrointestinal perforations resulting in peritonitis, skull fractures, and death. Gunshot injuries often do not have characteristic entry and exit wounds, as these may have healed by the time the animal is examined (Goldstein et al., 1999). Healed wounds associated with harpoon tips and exploding projectiles are well documented in bowhead whales (Philo et al., 1993).
0839_frame_C23 Page 531 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
531
Tags used for identifying marine mammals may also cause foreign body reactions (see Chapter 38, Tagging and Tracking). Pinnipeds tagged through the flippers may develop pressure necrosis beneath a tag if it is too tight, usually resulting in tag loss and subsequent healing. Cetacean tags can also elicit a foreign body reaction, although probably more due to the presence of foreign material and compromised blood supply; however, in a limited number of individuals examined closely, these wounds appear to heal well when the tag is removed (Orr et al., 1998). Explosives used for seismic exploration, underwater nuclear tests, and fishing have been reported to kill pinnipeds, dugongs (Dugong dugon), and river dolphins (reviewed in Richardson et al., 1995), although the lesions were not described. Sea otters killed near Amchitka Island, Alaska during nuclear testing had lesions in the lungs, heart, brain, and middle ear (R.L. Rausch, in Trasky, 1976). Two humpback whales (Megaptera novaeangliae) found dead near the site of repeated subbottom blasting had mechanical damage to the ears, including round window rupture, ossicular chain disruption, tissue dissection, bloody effusion of the peribullar spaces, and bilateral periotic fractures (Ketten et al., 1993). Inner ear damage was also reported in Weddell seals (Leptonychotes weddellii) that were exposed to blasts (Bohne et al., 1986).
Miscellaneous Integumentary System Little (other than above) is known about the etiology and pathogenesis of noninfectious skin lesions in marine mammals, although many of these lesions are observed and used to identify individuals (Kraus et al., 1986; Wilson et al., 1997). Sloughing of the epidermis from exposed areas of skin is common in stranded cetaceans, and is presumed to be a consequence of exposure to ultraviolet light (Chapter 6, Mass Strandings). However, it is rarely observed in healthy individuals maintained in climates with high ultraviolet exposure, and may thus be more of a response to drying (Ridgway, 1972; Greenwood et al., 1974; Geraci et al., 1986). Fissuring of the dermis around the blowhole is believed to be a consequence of drying (Simpson and Gardner, 1972). Ballooning degeneration of the epidermis has been observed by Maderson (in Simpson and Gardner, 1972) in a bottlenose dolphin maintained in fresh water. Patchy ulceration and necrosis of the epidermis are regularly observed in dolphins if the salt concentration of pool water falls below 1% (Simpson and Gardner, 1972; Greenwood et al., 1974). A number of traumatic skin lesions resulting from drying, abrasions, and pressure necrosis are described as consequences of capture and transport (Greenwood et al., 1974). The morphology of epidermal lesions classified as shallow lacerations, circular depressions, and epidermal sloughing in bowhead whales has been described in detail, although their etiology is unclear (Henk and Mullan, 1996). A characteristic ulcerative, hyperkeratotic skin disease has been described in northern elephant seals, but its etiology remains obscure (Beckmen et al., 1997) (see Chapter 41, Seals and Sea Lions). An alopecic skin lesion in a California sea lion with thinning of the epidermis, hyperkeratosis, and dilatation of cystic, keratin-filled hair follicles was reminiscent of hypothyroid skin conditions in other animal species (Howard, 1983). Circular areas of alopecia are commonly observed in Steller and California sea lions. The cause of these lesions is unknown, but occasionally mycotic organisms can be found within hair follicles on histopathology (Spraker, unpubl. data). Severe acanthosis and hyperkeratosis with secondary bacterial infections and septicemia were seen in two captive sea lions from one facility (Lowenstine, unpubl. obs.). The etiology was undetermined, but no further cases occurred after exhibit renovation. Extensive alopecia has been observed in gray seals around the United Kingdom, especially the Farne Isles (Hall, pers. comm.). Although some biopsies showed a mycotic dermatitis, the etiology remains obscure. Alopecia and acanthosis have also
0839_frame_C23 Page 532 Tuesday, May 22, 2001 11:52 AM
532
CRC Handbook of Marine Mammal Medicine
been described in polar bears (Ursus maritimus) (see Chapter 45, Polar Bears). Deposits of iron salts on the keratin of hairs of harbor seals resulting in a red pelage are common in seals from San Francisco Bay, California, and are rarely observed elsewhere (Allen et al., 1993). An unusual case of cutaneous gout, characterized by granulomatous dermatitis with intradermal uric acid deposits, which responded to treatment with allopurinol, was observed in an Amazon river dolphin (Inia geoffrensis) (Garman et al., 1983).
Musculoskeletal and Dental Systems Fractures are regularly observed in skeletons of wild marine mammals (Slijper, 1962; Ogden et al., 1981; Philo et al., 1990; De Guise et al., 1999). Although in recent times often attributed to boat strikes, Slijper (1962) states that fractures were as common in fossil whales, and believes that they were consequences of intraspecific aggression (see above). Rickets, characterized by multiple vertebral, rib, and limb bone fractures, and easily incised bones, has been diagnosed in two captive walrus calves fed an artificial formula (Griner, 1983). Osteomyelitis of the skull is well documented in pinnipeds (Cave and Bonner, 1987; Bergman et al., 1992; Junin and Castello, 1995). Cases are thought to result from abrasion or fracture of teeth, allowing entry of bacteria into the mandible (Stirling, 1969; Stroud and Roffe, 1979; Kenyon, 1981; Junin and Castello, 1995). Caries, characterized by destruction of the dental enamel, have been observed in bottlenose dolphins (Brooks and Anderson, 1998), sperm whales, killer whales (Slijper, 1962), and river dolphins (Ness, 1966). Gingivitis, recession of the gums, and periodontitis are regularly observed in California sea lions (Griner, 1983) and sea otters (see Chapter 44, Sea Otters). Hypomineralization of the teeth of a beluga was described by Overstrom et al. (1991). Dental anomalies such as malocclusion, accessory roots, and cystic retention of undeveloped teeth that have been documented in bottlenose dolphins (Brooks and Anderson, 1998) and harbor porpoises (Baker and Martin, 1992) may predispose animals to osteomyelitis of the mandible or skull. However, it has also been suggested that hyperadrenocorticism resulting from exposure to contaminants may enhance development of osteolytic lesions in the periodontal lamellae of gray and harbor seals (Bergman et al., 1992) (see Chapter 22, Toxicology). Peripheral limb and vertebral osteomyelitis and diskospondylitis are well documented in cetaceans and pinnipeds, and are often attributed to hematogenous spread of infection, although culture results are usually unavailable (Cowan, 1966; Lagier, 1977; Morton, 1978; Foley, 1979; Paterson, 1984; Thomas-Baker, 1986; Alexander et al., 1989). Pelvic neoplasms in sea lions often invade the pelvic bones and lumbosacral vertebral bodies, causing posterior paresis (Gulland et al., 1996). Degenerative and infectious arthritides have been described in the atlanto-occipital and/or humeroscapular joints in 22% of 59 bottlenose dolphins examined from the Gulf of Mexico (Turnbull and Cowan, 1999). Lesions ranged from mild roughening of the articular cartilage to complete erosion and ankylosis. Ankylosing spondylosis has also been observed in belugas (Martineau et al., 1988), a Bryde’s whale (Balaenoptera edeni) (Paterson, 1984), pilot whales (Globicephala malaena) (Cowan, 1966), harbor porpoises (Kinze, 1986; Baker and Martin, 1992), and California sea lions (Griner, 1983). Scoliosis and lordosis have been observed in cetaceans that survive stranding. However, freeliving bottlenose dolphins with lateral and sagittal deviation of the spine have been repeatedly observed swimming at sea, apparently able to survive (Pineau, pers. comm.; Tregenza, pers. comm.). Vertebral column malformations in ten wild delphinids sighted in New Zealand waters were recently reported by Berghan and Visser (2000). Vertebral abnormalities such as lateral curvature of the spinal processes and irregularities of the centra, as well as lordosis, have been
0839_frame_C23 Page 533 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
533
documented in dolphin and whale skeletons, but because the whole animals were not examined, the clinical significance of these lesions is unknown (Wells and Lawrence, 1976; Crovetto, 1982). Scoliosis has also been observed in a California sea otter (Rennie and Woodhouse, 1988). Severe necrotizing myopathies, with lesions consistent with capture myopathy of terrestrial animals have been described in California sea lions (Howard, 1983). A multifocal, necrotizing myopathy of skeletal muscle and myocardium was observed in northern fur seals on St. Paul Island, Alaska in 1990 and 1991, but the etiology remains obscure (Spraker, 1995). Emaciation is an extremely common finding in neonatal and juvenile marine mammals, especially pinnipeds. Emaciation of pups may be due to conditions affecting the mothers. Such conditions are numerous, and include failure of the mother to bond with her pup at birth; delay or failure of the mother to return from, or find her pup after, a foraging trip; trauma; agalactia; mastitis; or death. Factors affecting the pup that could result in emaciation are also numerous, and include dystocia, infection, hypothermia, and trauma. Emaciation of young pinnipeds is usually due to an inability to find food, especially during El Niño years (Trillmich et al., 1991); emaciation, however, may be secondary to a disease. Conversely, many emaciated animals have heavy parasite infections that may be secondary to emaciation. Gross lesions of emaciation are loss of adipose tissue and skeletal muscle, the distribution of which varies with species. In both pinnipeds and cetaceans, lipid is lost from the blubber layer resulting in a more fibrous appearance to the blubber. Loss of skeletal muscle mass and bone marrow fat may also occur in severely emaciated animals. The histological lesions observed in emaciated pinnipeds include serous atrophy of fat, atrophy of hepatocytes, myocytes, and myocardial cells, and loss of zymogen granules from the pancreatic acini.
Respiratory System Pneumonia and bronchopneumonia are frequently observed in many marine mammals in association with a variety of infectious agents and exposure to toxins (Chapter 15, Viral Diseases; Chapter 16, Bacterial Diseases; Chapter 18, Parasitic Diseases; Chapter 22, Toxicology), but are also occasionally observed in the absence of an obvious pathogen (Griner, 1983; Howard, 1983). Aspiration of milk following blunt trauma to the abdomen of recently nursed pups is occasionally seen in northern fur seals. Occasionally, pups playing in pools are trampled by older animals and aspirate contaminated rookery water, resulting in acute suppurative bronchopneumonia. Hemolytic Escherichia coli is commonly isolated from these pneumonic lungs (Spraker, in press). Intracellular hemosiderin has been regularly observed near blood vessels in areas of chronic inflammation (Simpson and Gardner, 1972). Anthracosis has been observed in lungs and mediastinal lymph nodes of bottlenose dolphins stranded off Florida (Rawson et al., 1991). Obstructive emphysema was described in a northern elephant seal (Saunders and Hubbard, 1966).
Digestive System Gastric and proximal intestinal erosions and ulcerations are common in pinnipeds, cetaceans, and sea otters. Although many are caused by parasites (Chapter 18, Parasitic Diseases), they may also be caused by bacterial infection (Chapter 16, Bacterial Diseases), stress (Chapter 13, Stress), or foreign bodies (Bossart et al., 1991). Occasionally, these eroded areas may perforate, resulting in peritonitis (Simpson and Gardner, 1972; Ridgway et al., 1975; Martineau et al., 1988; Fletcher et al., 1998). Gastric and intestinal obstructions resulting from ingestion of foreign bodies are widely reported in both captive (Appleby, 1962; Ridgway, 1965; Griner, 1983) and wild (Lambertsen and Kohn, 1987; Kastelein and Lavaleije, 1992; Tarpley and Marwitz, 1993; Baird and Hooker, 2000)
0839_frame_C23 Page 534 Tuesday, May 22, 2001 11:52 AM
534
CRC Handbook of Marine Mammal Medicine
marine mammals. Although stones are commonly observed in stomachs of healthy marine mammals (Taylor, 1993), they have also been associated with gastric ulceration and obstruction (Griner, 1983). Intestinal volvulus with necrosis has been observed in a pan-tropical spotted dolphin (Stenella attenuata) (Anderson and Rawson, 1997), bottlenose dolphins (Briggs and Murname, 1995; Higgins, in Anderson and Rawson, 1997), a bowhead whale (Heidal and Albert, 1994), a beluga (Martineau et al., 1988), a captive false killer whale (Pseudorca crassidens) (McBain, in Anderson and Rawson, 1997), two captive Australian fur seals (Reddacliff, 1988), a northern fur seal (Frasca et al., 1996), and sea otters (Williams and Pinard, 1983; Williams et al., 1987). Predisposing factors are unclear, although more active behavior has been suggested as important. Hepatic lipidosis can occur as a consequence of oil intoxication in sea otters (Lipscomb et al., 1994). Portal fibrosis and bile duct proliferation were observed in a captive California sea lion with periodic epilepsy (Simpson and Gardner, 1972). Cholelithiasis, with occlusion of the intrahepatic bile ducts by calculi, has been observed in California sea lions and northern elephant seals (Howard, 1983). Chronic periportal hepatitis is a common lesion in California sea lions and is thought to be secondary to biliary trematodiasis. Pancreatitis has been described in California sea lions and pilot whales, with leakage of pancreatic enzymes into the peripancreatic fat, causing fat necrosis (Howard, 1983; Bossart et al., 1991). Epithelial-lined cysts were observed in the pancreases of two pilot whales (Cowan, 1966).
Genitourinary System Renal calculi, both unidentified and triple phosphate, have been reported as incidental findings in harbor, ringed (Phoca hispida), northern elephant, and Weddell seals (Sweeney, 1974; Ridgway et al., 1975; Stroud, 1979; Griner, 1983), bottlenose dolphins (Simpson and Gardner, 1972), and in a beaked whale (Howard, 1983). In contrast, ammonium urate and uric acid calculi were associated with renal failure and tubular necrosis in harbor seals (Larsen, 1962; Stroud, 1979). Calculi causing obstruction of the ureter containing calcium, oxalate, and phosphate, have been observed in a stranded Pacific white-sided dolphin (Lagenorrhynchus obliquidens) (Cowan et al., 1986). Cystic kidneys are frequently observed in many species, and may be developmental, when not congenital or arising from obstructive disease (Howard, 1983). Tubular nephrosis has been observed in stranded cetaceans, and is associated with myoglobinuria, suggesting exertional myopathy (Cowan et al., 1986). Primary glomerular disease has rarely been reported, but Howard (1983) described a beaked whale with moderate membranoproliferative glomerulopathy, and a California sea lion with severe glomerulosclerosis. A pilot whale was observed with numerous sclerosed glomeruli, with small cortical arteries showing either recanalized occlusions or focal intimal thickening (Cowan, 1966). Renal and systemic amyloidosis have been diagnosed in older free-ranging sea lions, and are presumed to be of the secondary, AA, type (authors, unpubl. data). Vaginal calculi have been described in three species of delphinids, and were originally believed to be vaginal plugs formed from coagulated seminal fluid (Harrison, 1969; Sawyer and Walker, 1977; Benirschke et al., 1984; Cowan et al., 1986; Woodhouse and Rennie, 1991). However, further studies comparing composition and structure with those of delphinid fetuses suggest that calculi are fetal remains that can act as niduses for further calculus development (Benirschke et al., 1984; Perrin and Donovan, 1984; Woodhouse and Rennie, 1991). Allantoic calculi have been observed in the Ganges River dolphin (Platanista gangetica) (Pilleri, 1977). Uterine ruptures, prolapses, and torsions have been observed in California sea lions during late gestation after seizures due to domoic acid toxicity (Gulland, 2000). Reproductive abnormalities in northern fur seals on rookeries have been observed, including a ruptured
0839_frame_C23 Page 535 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
535
uterus with intra-abdominal delivery of the fetus, vaginal laceration with evisceration, and uterine prolapse (Spraker, unpubl. obs.). A uterine torsion was also observed in a California sea otter with scoliosis (Rennie and Woodhouse, 1988). Uterine horn stenosis and occlusion have been reported in gray and ringed seals (Helle et al., 1976; Bergman and Olsson, 1985; Baker, 1989b). These lesions have been attributed to contaminant exposure, although the etiology is unclear (Chapter 22, Toxicology). Luteinized ovarian cysts have been documented in striped dolphins (Munson et al., 1998). Interestingly, dystocia, resulting in death of the female with the calf lodged in the vagina, appears to be more common in harbor porpoises than in other marine mammal species, as several cases have been observed (Stroud and Roffe, 1979; Baker and Martin, 1992; Daoust and McBurney, 1997; Chivers, pers. com.; authors, unpubl. obs.).
Endocrine System Although the endocrine organs have important physiological effects that may become pathological, little is known about the effects of aging, reproductive cycle, season, starvation, and intercurrent disease on the histological appearance and function of these glands in marine mammals (Chapter 9, Anatomy). Interpretation of pathological significance of changes in histological appearance is thus still very subjective. Colloid goiter and granulomata have been reported in thyroids from pilot whales (Cowan, 1966), and colloid depletion and fibrosis in harbor seals and harbor porpoises (Schumacher et al., 1993). Chronic lymphocytic, interstitial thyroiditis and nodular hyperplasia of thyroid epithelial cells have been observed in northern right whale dolphins (Howard, 1983). Lipid depletion and vacuolar degeneration of the adrenals are frequently observed in stranded marine mammals, and are usually bilateral lesions presumed due to stress (Chapter 13, Stress) (Howard, 1983; Bossart et al., 1991). Adrenal cysts and nodules have been reported as incidental findings in belugas (De Guise et al., 1995). Adrenal hyperplasia and degeneration have also been documented, although their etiology remains unknown (Griner, 1983; Banish and Gilmartin, 1992; Lair et al., 1997). Hyperadrenocorticism associated with skull, uterine, and renal changes has been attributed to exposure to environmental contaminants (Bergman et al., 1992) (Chapter 22, Toxicology). An adrenal teratoma was observed in a Pacific white-sided dolphin as an incidental finding (Simpson and Gardner, 1972).
Cardiovascular System Degenerative disease of the aorta and coronary vessels, varying from small fibrous intimal plaques to larger fibrous plaques and medial necrosis, has been reported in a number of cetacean (Roberts et al., 1965; Cowan et al., 1986) and pinniped species (Prathap et al., 1966; Stout, 1969; Howard, 1983). Most commonly, fibrous intimal plaques were observed, with no complication by thrombi or hemorrhage (Truex et al., 1961; Roberts et al., 1965; Cowan, 1966; Stout, 1969). However, a killer whale with extensive atherosclerotic changes did develop ulceration and thrombi in the anterior descending coronary artery and aorta (Roberts et al., 1965). A 17-year-old Hawaiian monk seal with arteriosclerosis had cerebral infarcts and laminar necrosis of the cerebral cortex, and congestion of pulmonary and hepatic tissues suggestive of cardiac failure. Two other aged male seals had calcification of the internal elastic lamina in a number of vessels, suggesting age-related cardiovascular changes may occur in monk seals (Banish and Gilmartin, 1992). Etiology of these lesions is unclear, as a study of spontaneous lesions in Weddell seals showed that they do not contain significant amounts of lipid, and serum cholesterol levels did not correlate with presence of arterial lesions (Prathap et al., 1966). The similarity of these lesions to those seen in human syphilis cases
0839_frame_C23 Page 536 Tuesday, May 22, 2001 11:52 AM
536
CRC Handbook of Marine Mammal Medicine
has led to the suggestion that exposure to Treponema spp. may be important (Kelly and Jansen, 1960; Prathap et al., 1966). Disseminated intravascular coagulation has been documented in northern elephant seals as a consequence of vasculitis and septicemia associated with Otostrongylus circumlitus infestation (Gulland et al., 1997), and has been observed in other phocids associated with septicemia (authors, unpubl. obs.). Thrombosis of the pulmonary artery, with extensive lung necrosis, has been observed in elephant seals and northern fur seals (Griner, 1983). Myocardial contraction band necrosis, characterized by focal hypercontraction and lysis of contractile filaments in small groups of myocardial cells, has been well documented in a range of cetacean species following stranding (Bossart et al., 1985; 1991; Turnbull and Cowan, 1998). In terrestrial species, it is known to be a consequence of a catecholamine surge, and it is likely that this mechanism is important in stranded cetaceans, although it has yet to be investigated. Other myocardial lesions commonly observed include basophilic degeneration of myocardial fibers, and focal inflammation and scarring in a variety of species (Cowan, 1966; Griner, 1983). Subepicardial scarring was common in a set of stranded common dolphins (Delphinus delphis) and Pacific white-sided dolphins from Los Angeles County (Cowan et al., 1986) and was postulated to be a result of vascular spasm. Acute myocardial necrosis was seen in California sea lions dying acutely of domoic acid intoxication (Gulland et al., 2000). Vegetative valvular endocarditis has been described, although rarely, in pinnipeds, usually in association with other inflammatory lesions (Griner, 1983). Valvular endocarditis of the left atrioventricular valve was observed in a sea otter that died of thromboembolism, myocardial necrosis, and heart failure (Joseph et al., 1990). Valvular endocardiosis or valvular fibrosis was relatively common in neonatal manatees, and seemed to regress as the animals aged (Buergelt et al., 1990). Nodules have been observed on the mitral valve of a pilot whale (Cowan, 1966).
Lymphoid System Cowan (1966) reported fibrous and granulomatous nodules and scar tissue in the spleens of pilot whales killed for subsistence use. Hyaline sclerosis of the splenic capsule and residual subcapsular hemorrhages were observed in stranded common dolphins (Cowan et al., 1986), as well as in spinner dolphins caught as by-catch (Cowan and Walker, 1979). Siderotic plaques of the splenic capsule are common in northern fur seals and Steller sea lions on rookeries and haul-out areas, presumably due to blunt trauma to the abdomen as a result of hitting rocks, which causes splenic hemorrhage (Spraker, in press).
Nervous System and Special Senses Little is known about the effects of noise, other than blast injury, on marine mammal ears (see above, and Ketten, 1998). Hearing loss in an aged captive dolphin has been associated with cell loss and laminar demineralization of the inner ear, similar to that in humans with presbycusis (Ketten, 1998). Ocular lesions in pinnipeds are common (see Chapter 41, Seals and Sea Lions), but reports in other marine mammals are rare. Captive pinnipeds frequently develop corneal edema, ulcers, uveitis, cataracts, and panophthalmitis, but these lesions are rarely seen in wild pinnipeds, and their etiology and pathogenesis remain obscure (Griner, 1983). Panophthalmitis has been observed in wild northern fur seals following bite wounds to the head (Spraker, unpubl. obs.). Corneal opacity and blistering of unknown etiology have been observed in a bowhead whale (Zhu, 1997). Hemorrhagic encephalopathy suggestive of cerebrovascular occlusion and infarction has been described in two common dolphins and a California sea lion (Howard, 1983). In the
0839_frame_C23 Page 537 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
537
dolphins, cerebrovascular infarcts were located superficially in the cerebral hemisphere, with adjacent acute ischemic necrosis and early collapse of the neuropile. Encephalitis, characterized by lymphocytic cuffing, is occasionally observed as an apparently incidental finding in the brain of a number of species, including belugas (Martineau et al., 1988) and California sea lions (authors, unpubl. obs.). The etiology is currently unclear, and an infectious etiology is possible.
Acknowledgments The authors thank Todd O’Hara, Raymond Tarpley, Bill McClellan, and Pam Tuomi for helpful comments on the chapter; Rebecca Duerr and Kathy Zagzebski for editorial assistance; Ailsa Hall and Sylvain De Guise for critically reviewing the chapter; and Ailsa Hall, Susan Chivers, Nick Tregenza, John Heyning, and Pam Tuomi for personal communications.
References Acevedo-Whitehouse, K., Constantino-Casa, F., Aurioles-Gamboa, D., Rodriguez-Martinez, H.A., and Godinez-Reyes, C.R., 1999, Hepatic carcinoma with spleen metastasis in a California sea lion from the Gulf of California, J. Wildl. Dis., 35: 565–568. Albert, T.F., Migaki, G., Casey, H.W., and Philo, L.M., 1980, Healed penetrating injury of a bowhead whale, Mar. Fish. Rev., 42: 92–96. Alcorn, D.J. and Kam, A.K.H., 1986, Fatal shark attack on a Hawaiian monk seal (Monachus schauins landi), Mar. Mammal Sci., 2: 313–315. Alexander, J.W., Solangi, M.A., and Riegel, L.S., 1989, Vertebral osteomyelitis and suspected diskospondylitis in an Atlantic bottlenose dolphin (Tursiops truncatus), J. Wildl. Dis., 25: 118–121. Allen, S.G., Stephenson, M., Riseborough, R.W., Fancher, L., Shiller, A., and Smith, D., 1993, Redpelaged harbor seals of the San Francisco Bay region, J. Mammal., 74: 588–593. Ames, J.A., and Morejohn, C.V., 1980, Evidence of white shark, Carcharodon carcharias, attacks on sea otters, Enhydra lutris, Calif. Fish Game, 66: 196–209. Anderson, H.F., and Rawson, A.J., 1997, Volvulus with necrosis of intestine in Stenella attenuata, Mar. Mammal Sci., 13: 147–148. Anderson, W.I., Steinberg, H., Scott, D.W., and King, J.M., 1990, Cutaneous squamous cell carcinoma and multiple epidermoid cysts in a California sea lion, Aquat. Mammals, 16: 21–22. Andrews, R.C., 1921, A remarkable case of external hindlimbs in a humpback whale, Am. Mus. Novitates, 9: 1–6. Appleby, E.C., 1962, A case of gastric perforation by a foreign body in an elephant seal (Mirounga leonina), Nord. Veterinaermed., 14: 164–165. Baird, R.W., and Hooker, S.K., 2000, Ingestion of plastic and unusual prey by a juvenile harbour porpoise, Mar. Pollut. Bull., 40: 719–720. Baker, J.R., 1989a, Natural causes of death in non-suckling grey seals (Halichoerus grypus), Vet. Rec., 125: 500–503. Baker, J.R., 1989b, Pollution-associated uterine lesions in grey seals from the Liverpool Bay area of the Irish Sea, Vet. Rec., 125: 303. Baker, J.R., and Martin, A.R., 1992, Causes of mortality and parasites and incidental lesions in harbour porpoises (Phocoena phocoena) from British waters, Vet. Rec., 130: 554–558. Banish, L.D., and Gilmartin, W.G., 1992, Pathological findings in the Hawaiian monk seal, J. Wildl. Dis., 28: 428–434. Bannister, J.L., 1962, An intersexual fin whale Balaenoptera physalus (L.) from South Georgia, Proc. Zool. Soc. London, 141: 811–822.
0839_frame_C23 Page 538 Tuesday, May 22, 2001 11:52 AM
538
CRC Handbook of Marine Mammal Medicine
Beckmen, K.B., Lowenstine, L.J., Newman, J., Hill, J., Hanni, K., and Gerber, J., 1997, Clinical and pathological characterization of northern elephant seal skin disease, J. Wildl. Dis., 33: 438–449. Bedard, C., Kovacs, K., and Hammill, M., 1993, Cannibalism by grey seals, Halichoerus grypus, on Amet Island, Nova Scotia, Mar. Mammal Sci., 9: 421–424. Benirschke, K., and Marsh, H., 1984, Anatomic and pathologic observations of female reproductive organs in the short-finned pilot whale, Globicephala macrorhynchus, Rep. Int. Whaling Comm., 6: 451–454. Benirschke, K., Henderson, J.R., and Sweeney, J.C., 1984, A vaginal mass, containing fetal bones, in a common dolphin, Delphinus delphis, in Reproduction of Whales, Dolphins and Porpoise, Perrin, W.F., Browning, R.L., Jr., and DeMaster, D.P. (Eds.), Report of the International Whaling Commission, Special Issue 6, Cambridge, U.K., 457–458. Berghan, J., and Visser, I.N., 2000, Vertebral column malformations in New Zealand delphinids with a review of cases worldwide, Aquat. Mammals, 26: 17–25. Bergman, Å., 1997, Trends of disease complex in Baltic grey seals from 1977 to 1996: Improved gynecological health but still high prevalence of fatal intestinal wounds, Working Paper 19, presented at the Working Group on Seals and Small Cetaceans in European Seas, Stockholm, Sweden. Bergman, Å., and Olsson, M., 1985, Pathology of Baltic grey seal and ringed seal females with special reference to adrenocortical hyperplasia: Is environmental pollution the cause of a widely distributed disease syndrome? Finn. Game Res., 44: 47–62. Bergman, Å., Olsson, M., and Reiland, S., 1992, Skull bone lesions in the Baltic grey seal, Ambio, 21: 517–517. Bohne, B.A., Bozzay, D.G., and Thomas, J.A., 1986, Evaluation of inner ear pathology in Weddell seals, Antarct. J. U.S., 21: 208. Bossart, G.D., 1990, Invasive gingival squamous cell carcinoma in a California sealion (Zalophus californianus), J. Zoo Wildl. Med., 21: 92–94. Bossart, G.D., in press, Oral squamous cell carcinoma (suspected papilloma virus associated) in bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med. Bossart, G.D., Odell, D.K., and Altman, N.H., 1985, Cardiomyopathy in stranded pygmy and dwarf sperm whales, J. Am. Vet. Med. Assoc., 187: 1137–1140. Bossart, G.D., Walsh, M.T., Odell, D.K., Lynch, J.D., Beusse, D.O., Friday, R., and Young, W.G., 1991, Histopathologic findings of a mass stranding of pilot whales (Globicephala macrorhynchus), in Marine Mammal Strandings in the United States, Reynolds, J.E., and Odell, D.K. (Eds.), NOAA Technical Report, NMFS 98: 85–90. Bossart, G.D., Cray, C., Solorzano, J.L., Decker, S.J., Cornell, L.H., and Altman, N.H., 1996, Papillomavirallike papillomatosis in a killer whale, 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., Ewing, R., Lowe, M., Murphy, D., and Sweat, M., 1998, Cutaneous viral papillomatosis in manatees, in Proceedings, Captive Manatee Reintroduction/Release Workshop, Florida Marine Research Institute, St. Petersburg, FL, 21–22. Briggs, M.B., and Murname, R., 1995, Jejunal herniation and volvulus in an adult bottlenose dolphin, in Proceedings of the International Association for Aquatic Animal Medicine, 110–111. Brooks, L., and Anderson, H.F., 1998, Dental anomalies in bottlenose dolphins, Tursiops truncatus, from the west coast of Florida, Mar. Mammal Sci., 14: 849–853. Brown, R.J., Smith, A.W., and Keyes, M.C., 1975, Renal fibrosarcoma in the northern fur seal, J. Wildl. Dis., 11: 23–25. Brown, R.J., Smith A.W., Morejohn, G.V., and De Long, R.L., 1980, Metastatic adenocarcinoma in two California sea lions (Zalophus californianus), J. Wildl. Dis., 16: 261–266. Buck, J.D., Spotte, S., and Gadbaw, J.J., Jr., 1984, Bacteriology of the teeth from a great white shark: Potential medical implications for shark bite victims, J. Clin. Microbiol., 20: 849–851.
0839_frame_C23 Page 539 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
539
Buergelt, C.D., Bonde, R.K., Beck, C.A., and O’Shea, T.J., 1990, Myxomatous transformation of heart valves in Florida manatees, J. Zoo Wildl. Med., 21: 220–227. Castello, H.P., 1977, Food of a killer whale: Eagle sting-ray, Myliobatis, found in the stomach of a stranded Orcinus orca, Sci. Rep. Whales Res. Inst., 29: 107–111. Cave, A.J.E., and Bonner, W.N., 1987, Facial asymmetry in a leopard seal, Hydrurga leptonyx, Br. Antarct. Surv. Bull., 75: 67–71. Clarke, R., 1956, Sperm whales of the Azores, Discovery Rep., 28: 237–298. Cockrill, W.R., 1960, Pathology of the cetacea: A veterinary study on whales, Br. Vet. J., 116: 1–30. Colyer, F., 1936, Variations and Disease of the Teeth of Animals, John Bale, Sons & Danielsson Limited, London, 115–118. Corkeron, P.J., Morris, R.J., and Bryden, M.M., 1987a, Interactions between bottlenose dolphins and sharks in Moreton Bay, Queensland, Aquat. Mammals, 13: 111–114. Corkeron, P.J., Morris, R.J., and Bryden, M.M., 1987b, A note on the healing of large wounds in bottlenose dolphins, Tursiops truncatus, Aquat. Mammals, 13: 96–98. Cornell, L.H., Golden, B.J., and Osborn, K.G., 1975, Pseudopersistent urachus in a baby walrus, J. Am. Vet. Med. Assoc., 167: 548–549. Cowan, D.F., 1966, Pathology of pilot whale (Globicephala melaena), a comparative study, Arch. Pathol., 82: 178–189. Cowan, D.F., and Walker, W.A., 1979, Disease factors in Stenella attenuata and Stenella longirostris taken in the eastern tropical Pacific yellowfin tuna purse seine fishery, NOAA National Marine Fisheries Service, Southwest Fisheries Center, La Jolla, CA, Administrative Report, LJ-79-32C, 21 pp. Cowan, D.F., Walker, W.A., and Brownell, R.L., Jr., 1986, Pathology of small cetaceans stranded along the southern California beaches, in Research on Dolphins, Bryden, M.M., and Harrison, R. (Eds.), Oxford University Press, Oxford, U.K., 323–367. Crovetto, A., 1982, A propos d’une malformation laterale da la colonne vertebrale d’une baleine bleue, Balaenoptera musculus L. (Cetacea, Balaenopteridae) et de sa signification fonctionnelle, in 107 Congres National des Societes Savantes, Brest, Sciences, 2: 89–98. Csordas, S.E., 1966, Congenital penis malformation in southern elephant seals (Mirounga leonina), J. Mammal., 47: 731. Daoust, P., and McBurney, M., 1997, Wildlife Health Centre Newsletter, Canadian Cooperative Wildlife Health Centre, 4: 5. De Guise, S., Lagace, A., and Beland, P., 1994a, Tumors in St. Lawrence beluga whales (Delphinapterus leucas), Vet. Pathol., 31: 444–449. De Guise, S., Lagace, A., and Beland, P., 1994b, Gastric papillomas in eight St. Lawrence beluga whales (Delphinapterus leucas), J. Vet. Diagn. Invest., 6: 385–388. De Guise, S., Lagace, A., and Beland, P., 1994c, True hermaphroditism in a St. Lawrence beluga whale (Delphinapterus leucas), J. Wildl. Dis., 30: 287–290. De Guise, S., Lagace, A., Beland, P., Girard, C., and Higgins, R., 1995, Non-neoplastic lesions in beluga whales (Delphinapterus leucas) and other marine mammals from the St. Lawrence estuary, J. Comp. Pathol., 112: 257–271. De Guise, S., St. Aubin, D.J., Morre, M., and Knowlton, A., 1999, A non-union chronic fracture in a north Atlantic right whale: The importance of histopathology, in Proceedings of the 13th Biennial Conference on the Biology of Marine Mammals, Wailea, HI, November 28–December 3, 45. Dufault, S., and Whitehead, H., 1995, An encounter with recently wounded sperm whales (Physeter macrocephalus), Mar. Mammal Sci., 11: 560–563. Duignan, P.J., Hunter, J.E.B., Visser, I.N., Jones, G.W., and Nutman, A., 2000, Stingray spines: A potential cause of killer whale mortality in New Zealand, Aquat. Mammals, 17: 143–147. Fletcher, D., Gulland, F.M.D., Haulena, M., Lowenstine, L.J., and Dailey, M., 1998, Nematode-associated gastrointestinal perforations in stranded California sea lions (Zalophus californianus), in International Association for Aquatic Animal Medicine 29th Annual Conference Proceedings, San Diego, CA, 29: 59.
0839_frame_C23 Page 540 Tuesday, May 22, 2001 11:52 AM
540
CRC Handbook of Marine Mammal Medicine
Flom, J.O., Brown, R.J., Jones, R.E., and Schonewald, J., 1980, Vaginal fibromas in a beaked whale, Mesoplodon densirostris, J. Wildl. Dis., 16: 99–102. Foley, R.H., 1979, Osteomyelitis of the flipper of a bottlenose dolphin, J. Am. Vet. Med. Assoc., 175: 999. Fordyce, R.E., and Watson, A.G., 1998, Vertebral pathology in an early Oligocene whale (Cetacea, ?Mysticeti) from Wharekuri, North Otago, New Zealand, Mainz. Naturwiss. Arch., Beiheft: Festschrift zum 70 Geburtstag von Karlheinz Rothausen, Grimm, K.I., Grimm, M.C., and Morlo, M. (Eds.), 21: 161–176. Forrester, D.J., 1992, Manatees, in Parasites and Diseases of Wild Mammals in Florida, University Press of Florida, Gainseville, 255–274. Fowler, C.W., 1986, Marine debris and northern fur seals: A case study, Mar. Pollut. Bull., 18: 326–335. Frasca, S., Dunn, J.L., and Van Kruningen, H.J., 1996, Acute gastric dilatation with volvulus in a northern fur seal (Callorhinus ursinus), J. Wildl. Dis., 32: 548–551. Gallo-Reynoso, J., and Figueroa-Carranza, A.L., 1992, A cookiecutter shark wound on a Guadalupe fur seal male, Mar. Mammal Sci., 8: 428–429. Garman, R.H., Nuzzi, M.J., and Geraci, J.N., 1983, Cutaneous gout in an Amazon dolphin, J. Am. Vet. Med. Assoc., 183: 1292–1294. Gentry, R.L., and Johnson, J.H., 1981, Predation by sea lions on northern fur seal neonates, Mammalia, 45: 423–430. George, J.C., and Suydam, R., 1998, Observations of killer whale (Orcinus orca) predation in the northeastern Chukchi and western Beaufort Seas, Mar. Mammal Sci., 14: 330–332. George, J.C., Philo, L.M., Hazard, K., Withrow, D., Carroll, G.M., and Suydum, R., 1994, Frequency of killer whale (Orcinus orca) attacks and ship collisions based on scarring on bowhead whales (Balaena mysticetus) of the Bering-Chukchi-Beaufort Seas Stock, Arctic, 47: 247–255. Geraci, J.R., St. Aubin, D.J., and Hicks, B.D., 1986, The epidermis of odontocetes: A view from within, in Research on Dolphins, Bryden, M.M., and Harrison, R. (Eds.), Clarendon Press, Oxford, U.K., 3–21. Geraci, J.R., Palmer, C., and St. Aubin, D.J., 1987, Tumors in cetaceans: Analysis and new findings, Can. J. Fish. Aquat. Sci., 44: 1289–1300. Girard, C., Lagace, R., Higgins, R., and Beland, P., 1991, Adenocarcinoma of the salivary gland in a beluga whale (Delphinapterus leucas), J. Vet. Diagn. Invest., 3: 264–265. 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. Gray, K.N., and Conklin, R.H., 1974, Multiple births and cardiac anomalies in the bottle-nosed dolphin, J. Wildl. Dis., 10: 155. Greenwood, A.G., Harrison, R.J., and Whitting, H.W., 1974, Functional and pathological aspects of the skin of marine mammals, in Functional Anatomy of Marine Mammals, Harrison, R.J. (Ed.), Academic Press, New York, 73–110. Grimes, D.J., Jacobs, D., Swartz, D.G., Brayton, P.R., and Colwell, R.R., 1993, Numerical taxonomy of Gram-negative oxidase-positive rods from carcharhinid sharks, Int. J. Syst. Bacteriol., 43: 88–89. Griner, L.A., 1971, Malignant leukemic lymphoma in two harbor seals (Phoca vitulina geronimensis), Am. J. Vet. Res., 32: 827–830. Griner, L.A., 1983, Pathology of Zoo Animals, Zoological Society of San Diego, San Diego, CA, 465 pp. Gulland, F.M.D., 2000, Domoic acid toxicity in California sea lions stranded along the central California coast, May–October 1998, NOAA Technical Memo, NMFS-OPR-17, National Marine Fisheries Service, U.S. Department of Commerce, 45 pp. 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.
0839_frame_C23 Page 541 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
541
Gulland, F.M.D., Beckmen, K., Burek, K., Lowenstine, L., Werner, L., Spraker, T., and Harris, E., 1997, Otostrongylus circumlitus infestation of northern elephant seals (Mirounga angustirostris) stranded in central California, Mar. Mammal Sci., 13: 446–459. Harcourt, R., 1993, Individual variation in predation on fur seals by southern sea lions (Otaria byronia) in Peru, Can. J. Zool., 71: 1908–1911. Harms, C.A., Gage, L.J., Smith, D.M., and Beckmen, K.B., 1994, Clinical challenge, J. Zoo Wildl. Med., 25: 301–303. Harrison, R.J., 1969, Reproduction and reproductive organs, in The Biology of Marine Mammals, Anderson, H.T. (Ed.), Academic Press, New York, 253–348. Heezen, B.C., 1957, Whales entangled in deep-sea cables, Deep Sea Res., 4: 105–115. Heidel, J.R., and Albert, T.F., 1994, Intestinal volvulus in a bowhead whale Balaena mysticetus, J. Wildl. Dis., 30: 126–128. Helle, E., Olsson, M., and Jensen, S., 1976, PCB levels correlated with pathological changes in seal uteri, Ambio, 5: 261–263. Henk, W.G., and Mullan, D.L., 1996, Common epidermal lesions in the bowhead whale, Balaena mysticetus, Scanning Microsc., 10: 905–916. Heyning, J.E., and Lewis, T.D., 1990, Entanglement of baleen whales in fishing gear off southern California, Rep. Int. Whaling Comm., 40: 427–431. Howard, E.B., 1983, Miscellaneous diseases, in Pathobiology of Marine Mammal Diseases, Vol. 2, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 164–225. Howard, E.B., Britt, J.O., and Simpson, J.G., 1983, Neoplasms in marine mammals, in Pathobiology of Marine Mammal Diseases, Vol. 2, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 95–162. Huey, L.M., 1925, Recent observations on the elephant seal, J. Mammal., 5: 237–242. Jenkins, R.L., and Cardeilhac, P.T., 1982, Intestinal perforation by a stingray spine in an Atlantic bottlenose dolphin resulting in fatal peritonitis, Abstr., in Proceedings of the 13th International Association for Aquatic Animal Medicine Association, May 9–13, Baltimore, MD, 12. Jepson, P.D., and Baker, J.D., 1998, Bottlenose dolphins (Tursiops truncatus) as a possible cause of acute traumatic injuries in porpoises (Phocoena phocoena), Vet. Rec., 143: 614–615. Jones, E.C., 1971, Isistius brasiliensis, a squaloid shark, the probable cause of crater wounds on fishes and cetaceans, Fish. Bull. U.S., 69: 791–798. Joseph, B.E., Cornell, L.H., and Migaki, G., 1986, Metastatic squamous cell carcinoma in a beached California sea lion (Zalophus californianus), J. Wildl. Dis., 22: 281–283. Joseph, B., Spraker, T.R., and Migaki, G., 1990, Valvular endocarditis in a northern sea otter (Enhydra lutris), J. Zoo Wildl. Med., 21: 88–91. Junin, M., and Castello, H.P., 1995, Osteomyelitis of the skull in a leopard seal, Hydrurga leptonyx, Mar. Mammal Sci., 11: 403–406. Kamiya, T., and Miyazaki, N., 1974, A malformed embryo of Stenella coeruleoalba, Sci. Rep. Whales Res. Inst., 26: 259–263. Kamiya, T., Miyazaki, N., and Shiraga, S., 1981, First case of dicephaly in Odontoceti, Sci. Rep. Whales Res. Inst., 33: 127–129. Kastelein, R.A., and Lavaleije, M.S.S., 1992, Foreign bodies in the stomach of a female harbour porpoise (Phocoena phocoena) from the North Sea, Aquat. Mammals, 18: 40–46. Kawamura, A., 1990, A malformed fetus of a southern Sei whale, Sci. Rep. Cetacean Res., 1: 93–95. Kawamura, A., and Kashita, K., 1971, A rare double monster of dolphin Stenella coeruleoalba, Sci. Rep. Whales Res. Inst., 23: 139–140. Kelly, A.L., and Jansen, D., 1960, Chronic aortitis in the California sea lion, Zalophus californianus, Nature, 186: 731. Kenyon, K.W., 1981, Monk seals, in Handbook of Marine Mammals, Vol. 2: Seals, Ridgway, S.H., and Harrison, R.J. (Eds.), Academic Press, London, 195–220.
0839_frame_C23 Page 542 Tuesday, May 22, 2001 11:52 AM
542
CRC Handbook of Marine Mammal Medicine
Ketten, D.R., 1998, Marine mammal auditory systems: A summary of audiometric data and anatomical data and its implications for underwater acoustic impacts, NOAA Technical Memo, NMFSSWFSC-256, 74 pp. 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. King, J.E., 1964, Seals of the World, Trustees of the British Museum, London, 51–56. Kinze, C.C., 1986, Note on the occurrence of spondylitis deformans in a sample of harbour porpoises (Phocoena phocoena (L.)) taken in Danish waters, Aquat. Mammals, 12: 25–27. Klontz, K.C., Mullen, R.C., Corbyons, T.M., and Barnard, W.P., 1993, Vibrio wound infections in humans following shark attack, J. Wilderness Med., 4: 68–72. Kraus, S.D., 1990, Rates and potential causes of mortality in North Atlantic right whales (Eubalaena glacialis), Mar. Mammal Sci., 6: 278–291. Kraus, S.D., Moore, K.E., Price, C.A., Crone, M.J., Watkins, W.A., Winn, H.E., and Prescott, J.H., 1986, The use of photographs to identify individual North Atlantic right whales (Eubalaena glacialis), in Right Whale: Past and Present Status, Brownell, R.L., Jr., Best, P.B., and Prescott, J.H. (Eds.), Report of the International Whaling Commission, Special Issue, 10: 145–151. Kuiken, T., 1994, Review of the criteria for the diagnosis of by-catch in cetaceans, in Proceedings of the 2nd European Cetacean Society Workshop on Cetacean Pathology, Montpellier, France, 2 March. Kuiken, T., Simpson, V.R., Allchin, C.R., Bennett, P.M., Codd, G.A., Harris, E.A., Howes, G.J., Kennedy, S., Kirkwood, J.K., Law, R.J., Merrett, N.R., and Phillips, S., 1994, Mass mortality of common dolphins (Delphinus delphis) in south-west England due to incidental capture in fishing gear, Vet. Rec., 134: 81–89. Lagier, R., 1977, Spondylosis in the whale—an anatomicroradiologic study of an osteophyte, Scand. J. Rheumatol., 6: 245–249. Lair, S., Beland, S., De Guise, S., and Martineau, D., 1977, Adrenal hyperplastic and degenerative changes in beluga whales, J. Wildl. Dis., 33: 430–437. Lair, S., De Guise, S., and Martineau, D., 1998, Uterine adenocarcinoma with abdominal carcinomatosis in a beluga whale, J. Wildl. Dis., 34: 373–376. Lambertsen, R.H., and Kohn, B.A., 1987, Unusual multisystemic pathology in a sperm whale bull, J. Wildl. Dis., 23: 510–514. Lambertsen, R.H., Kohn, B.A., Sundberg, J.P., and Buergelt, C.D., 1987, Genital papillomatosis in sperm whale bulls, J. Wildl. Dis., 23: 361–367. Landy, R.B., 1980, A review of neoplasia in marine mammals, in The Comparative Pathology of Zoo Animals, Montali, R., and Migaki, G. (Eds.), Smithsonian Institution Press, Washington, D.C., 579–563. Lanthier, C., Dupuis, J., and Pare, J., 1998, Agenesis of a radius in a polar bear cub (Ursus maritimus), J. Zoo Wild Anim. Med., 29: 65–67. Larsen, S., 1962, A survey of the post mortem findings in pinnipedia autopsied during a ten-year period, 1952–1961, Nord. Veterinaermed., 14: 150–160. Laws, R.M., 1953, The elephant seal (Mirounga leonina Linn.) I. Growth and age, Falkland Islands Dependency Survey Scientific Report 8. LeBoeuf, B.J., Riedman, M., and Keyes, R.S., 1982, White shark predation on pinnipeds in California coastal waters, Fish. Bull. U.S., 80: 891–895. LeBoeuf, B.J., Cosker, J.E., and Hewitt, J., 1987, Crater wounds in northern elephant seals: The cookiecutter shark strikes again, Fish. Bull. U.S., 85: 387–392. Leipold, H.W., 1980, Congenital defects in zoo and wild animals: A review, in The Comparative Pathology of Zoo Animals, Montali, R.J., and Migaki, G. (Eds.), Smithsonian Institution Press, Washington, D.C., 457–470. Lipscomb, T.P., Harris, R.K., Rebar, A.H., Ballachey, B.E., and Haebler, R., 1994, Pathology of sea otters, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 265–281.
0839_frame_C23 Page 543 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
543
Lipscomb, T., Scott, D.P., Garber, R.L., Krafft, A.E., Tsai, M.M., Lichy, J.H., Taubenberger, J.K., Schulman, F.Y., and Gulland, F.M.D., 2000, Common carcinoma of California sea lions: Evidence for genital origin and association with a novel herpesvirus, Vet. Pathol., 37: 609–617. Martineau, D., Lagace, A., Masse, R., Morin, M., and Beland, P., 1985, Transitional cell carcinoma of the urinary bladder in a beluga whale (Delphinapterus leucas), Can. Vet. J., 26: 297–302. 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., and Beland, P., 1995, Intestinal adenocarcinomas in two beluga whales (Delphinapterus leucas) from the estuary of the St. Lawrence River, Can. Vet. J., 36: 563–565. Mattlin, R.H., 1978, Pup mortality of the New Zealand fur seal (Arctocephalus forster Lesson), N.Z. J. Ecol., 1: 138–144. Mawdesley-Thomas, L.E., 1971, An ovarian tumour in a southern elephant seal (Mirounga angustirostris), Vet. Pathol., 8: 9–15. Mawdesley-Thomas, L.E., 1974, Some aspects of neoplasia in marine mammals, Adv. Mar. Biol., 12: 151–231. Mawdesley-Thomas, L.E., and Bonner, W.N., 1971, Uterine tumours in a grey seal (Halichoerus grypus), J. Pathol., 103: 205–208. McClellan, W.A., Thayer, V.G., and Pabst, D.A., 1996, Stingray spine mortality in a bottlenose dolphin, Tursiops truncatus, from North Carolina waters, J. Elisha Mitchell Sci. Soc., 112: 98–101. McFee, W., Root, H., Friedman, R., and Kolman E., 1997, A stingray spine in the scapula of a bottlenose dolphin, J. Wildl. Dis., 33: 921–924. Migaki, G., and Albert, T.F., 1982, Lipoma of the liver in a bowhead whale (Balaena mysticetus), Vet. Pathol., 19: 329–331. Migaki, G., Woodard, J.C., and Goldston, R.T., 1978, Renal adenoma in an Atlantic bottlenosed dolphin, Am. J. Vet. Res., 39: 1920–1921. Miller, E.H., 1996, Violent interspecific sexual behavior by male sea lions (Otariidae): Evolutionary and phylogenetic implications, Mar. Mammal Sci., 12: 468–476. Moore, J.J., and Stackhouse, L.L., 1978, Pancreatic duct adenoma and strangulation of the small intestine in a California sea lion (Zalophus californianus), J. Wildl. Dis., 14: 445–446. Morejohn, G.V., Ames, J.A., and Lewis, D.B., 1975, Post mortem studies of sea otters, Enhydra lutris L., in California, Marine Resources Technical Report 30, California Department of Fish and Game. Morgan, L., Hanni, K., and Lowenstine, L., 1996, Age and pathological findings for two female Steller sea lions stranded on the northern California coast, Calif. Fish Game, 82: 81–86. Morton, B., 1978, Osteomyelitis of the spine of a dolphin, J. Am. Vet. Med. Assoc., 173: 1109–1110. Mossner, S., and Ballschmiter, K., 1997, Marine mammals as global pollution indicators for organochlorines, Chemosphere, 34: 1285–1296. Munson, L., Calzada, N., Kennedy, S., and Sorensen, T.B., 1998, Luteinized ovarian cysts in Mediterranean striped dolphins, J. Wildl. Dis., 34: 656–660. Ness, A.R., 1966, Dental caries in the plantanistid whale, Inia geoffrensis, J. Comp. Pathol., 76: 271–272. Nishiwaki, M., 1953, Hermaphroditism in a dolphin (Prodelphinus caeruleo-albus), Sci. Rep. Whales Inst., (Tokyo), 8: 215–218. Obendorf, D.L., and Presidente, P.J.A., 1978, Foreign body perforation of the esophagus initiating traumatic pericarditis in an Australian fur seal, J. Wildl. Dis., 14: 451–454. Ogawa, T. and Kamiya, T., 1957, A case of the cachelot with protruded rudimentary hindlimbs, Sci. Rep. Whales Inst., 12: 197–208. Ogden, J.A., Conlogue, G.J., Light, T.R., and Sloan, T.R., 1981, Fractures of the radius and ulna in a skeletally immature fin whale, J. Wildl. Dis., 17: 111–116. Orams, M.B., and Deakin, R.B., 1997, Report on the healing of a large wound in a bottlenose dolphin (Tursiops truncatus), in Marine Mammal Research in the Southern Hemisphere, Vol. 1, Status, Ecology and Medicine, Hindell, M., and Kemper, C. (Eds.), Surrey, Beatty and Sons, Chipping, Norton, New South Wales, Australia, 170–173.
0839_frame_C23 Page 544 Tuesday, May 22, 2001 11:52 AM
544
CRC Handbook of Marine Mammal Medicine
Orr, J.R., and Harwood, L.A., 1998, Possible aggressive behavior between narwhal (Monodon monoceros) and a beluga (Delphinapterus leucas), Mar. Mammal Sci., 14: 182–185. Orr, J.R., St. Aubin, D.J., Richard, P.R., and Heide-Jorgensen, M.P., 1998, Recapture of belugas, Delphinapterus leucas, tagged in the Canadian Arctic, Mar. Mammal Sci., 14: 829–834. Osborn, K.G., Joseph, B.E., Cornell, L.H., Duffield, D.A., Chamberlin-Lea, J., Wilson, B.J., Munn, R.J., Kluge, J.D., and Marx, P.E., 1988, Leukemic lymphoma in harbor seals (Phoca vitulina): Report of three cases occurring over two years and discussion of pathogenesis, in Proceedings of the American Association of Zoo Veterinarians, Nov. 6–10, Toronto, Canada, 96. Oshuni, S., 1965, A dolphin (Stenella caeruleoalba) with protruded rudimentary hindlimbs, Sci. Rep. Whales Inst., 19: 135–136. Overstrom, N.A., Spotte, S., Dunn, L., Goren, A.D., and Kaufman, H.W., 1991, A resident belukha whale (Delphinapterus leucas) in Long Island Sound, in Marine Mammal Strandings in the United States, Reynolds, J.E., and Odell, D.K. (Eds.), NOAA Technical Report, NMFS 98: 143–149. Nemoto, T., 1963, New records of sperm whales with protruded rudimentary hindlimbs, Sci. Rep. Whales Inst., 8: 127–132. Paterson, R.A., 1984, Spondylitis deformans in a Bryde’s whale (Balaenoptera edeni Anderson) stranded on the southern coast of Queensland, J. Wildl. Dis., 20: 250–252. Patterson, I.A.P., Reid, R.J., Wilson, B., Grellier, K., Ross, H.M., and Thompson, P.M., 1998, Evidence for infanticide in bottlenose dolphins: An explanation for violent interactions with harbour porpoises? Proc. R. Soc. London Biol. Sci., 265: 1167–1170. Pavia, A.T., Bryan, J.A., Maher, K.L., Hester, T.R., and Farmer III, J.J., 1989, Vibrio carchariae infection after shark bite, Ann. Intern. Med., 111: 85–86. Perrin, W.F., and Donovan, G.P., 1984, Report of the workshop, in Reproduction of Whales, Dolphins and Porpoises, Perrin, W.F., Brownell, R.L., Jr., and DeMaster, D.P. (Eds.), Report of the International Whaling Commission, Special Issue 6, Cambridge, U.K., 2–24. Perrin, W.F., Brownell, R.L., Jr., and DeMaster, D.P. (Eds.), 1984, Reproduction of Whales, Dolphins and Porpoises, Report of the International Whaling Commission, Special Issue 6, Cambridge, U.K., 457–458. Perrin, W.F., Donovan, G.P., and Barlow, J., 1994, Gillnets and Cetaceans, Report of the International Whaling Commission, Special Issue 15, Cambridge, U.K., 629 pp. Philo, L.M., Hanns, C., and George, J.C., 1990, Fractured mandible and associated oral lesions in a subsistence-harvested bowhead whale (Balaena mysticetus), J. Wildl. Dis., 26: 125–128. Philo, L.M., George, J.C., and Albert, T.F., 1992, Rope entanglement of bowhead whales (Balaena mysticetus), Mar. Mammal Sci., 8: 306–311. Philo, L.M., Shotts, E.B., and George, J.C., 1993, Morbidity and mortality, in The Bowhead Whale, Burns, J., Hanns, C., Montague, J., and Cowles, C. (Eds.), The Society for Marine Mammalogy, Lawrence, Kansas, 275–312. Piérard, J., Bisaillon, A., and Larivière, N., 1977, Hypertrophic bone lesion in an Atlantic walrus (Odobenus odobenus), Vet. Pathol., 14: 291–293. Pilleri, G., 1966, Hirnlipom beim Buckelwal, Megaptera novaeangliae, Pathol. Vet., 5: 35–40. Pilleri, G., 1968, Cerebrale Neurofibrome beim Finnwal, Balaenoptera physalus, Pathol. Vet., 5: 35–40. Pilleri, G., 1977, Hippomanes (allantoic “calculi”) in the Ganges dolphin, Platanista gangetica, Invest. Cetacea, 8: 121–122. Pollanen, M.S., Cheung, C., and Chiasson, D.A., 1997, The diagnostic value of the diatom test for drowning, I. Utility: A retrospective analysis of 771 cases of drowning in Ontario, Canada, J. Forens. Sci., 42: 281–285. Prathap, K., Ardlie, N.G., Paterson, J.C., and Schwartz, C.J., 1966, Spontaneous arterial lesions in the Antarctic seal, Arch. Pathol., 82: 287–296. Rawson, A.J., Anderson, H.F., Patton, G.W., and Beecher, T., 1991, Anthracosis in the Atlantic bottlenose dolphin (Tursiops truncatus), Mar. Mammal Sci., 7: 413–416.
0839_frame_C23 Page 545 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
545
Read, A.J., and Murray, K.T., 2000, Gross evidence of human-induced mortality in small cetaceans, U.S. Department of Commerce, NOAA Technical Memorandum, NMFS-OPR-15, 21 pp. Reddacliffe, G., 1988, Fatal intestinal torsion in two captive fur seals, in Marine Mammals of Australia, Augee, M.L. (Ed.), Royal Zoological Society of New South Wales, Sydney, Australia, 135–136. Reimer, D.C., and Lipscomb, T.P., 1998, Malignant seminoma with metastasis and herpesvirus infection in a free-living sea otter (Enhydra lutris), J. Zoo Wild Anim. Med., 29: 35–39. Reiter, J., Stinson, N.L., and LeBoeuf, B.J., 1978, Northern elephant seal development: The transition from weaning to nutritional independence, Behav. Ecol. Sociobiol., 3: 337–367. Rennie, C.J., and Woodhouse, C.D., 1988, Scoliosis and uterine torsion in a pregnant sea otter (Enhydra lutris) from California, J. Wildl. Dis., 24: 582–584. Rewell, R.E., and Willis, R.A., 1949, Some tumors of wild animals, J. Pathol. Bacteriol., 61: 454–456. Rewell, R.E., and Willis, R.A., 1950, Some tumors found in whales, J. Pathol. Bacteriol., 62: 450–452. Richardson, W.J., Greene, C.R., Malme, C.I., and Thomson, D.H. (Eds.), 1995, Marine Mammals and Noise, Academic Press, San Diego, CA, 576 pp. Ridgway, S.H., 1965, Medical care of marine mammals, J. Am. Vet. Med. Assoc., 71: 237–242. Ridgway, S.H., 1972, Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 590–747. Ridgway, S.H., Geraci, J.R., and Medway, W., 1975, Diseases of pinnipeds, Rapp. P. V. Reun. Cons. Int. Explor. Mer, 169: 327–337. Roberts, J.C., Boice, R.C., Brownell, R.L., Jr., and Brown, D.H., 1965, Spontaneous atherosclerosis in Pacific toothed and baleen whales, in Comparative Atherosclerosis, Roberts, J.C., and Strauss, R. (Eds.), Harper & Row, New York, 151. Robinson, S., Wynen, L., and Goldsworthy, S., 1999, Predation by a Hooker’s sea lion (Phocarctos hookeri) on a small population of fur seals (Arctocephalus spp.) at Macquarie Island, Mar. Mammal Sci., 15: 888–893. Ross, H.M., and Wilson, B., 1996, Violent interactions between bottlenose dolphins and harbour porpoises, Proc. R. Soc. London B, 263: 283–286. Sato, S., Kitamura, H., Mori, M., Fukazawa, M., Takeda, M., and Kadota, K., 1998, Adenocarcinoma of the lung in a Steller sea lion (Eumetopias jubatus), J. Vet. Med. Sci., 60: 1349–1351. Saunders, A.M., and Hubbard, R.C., 1966, Obstructive emphysema in an elephant seal, Mirounga angustirostris, Lab. Anim. Care, 16: 217–223. Sawyer, J.E., and Walker, W.A., 1977, Vaginal calculi in the dolphin, J. Wildl. Dis., 13: 346–348. Schroeder, R.J., DelliQuadri, C.A., McIntyre, R.W., and Walker, W.A., 1973, Marine mammal disease surveillance program in Los Angeles County, J. Am. Vet. Med. Assoc., 163: 580. Schumacher, U., Zahler, S., Horney, H.-P., Hiedemann, G., Skirnisson, K., and Welsch, U., 1993, Histological investigations on the thyroid glands of marine mammals (Phoca vitulina, Phocoena phocoena) and the possible implications of marine pollution, J. Wildl. Dis., 29: 103–108. Simpson, J.G., and Gardner, M.B., 1972, Comparative microscopic anatomy of selected marine mammals, in Mammals of the Sea, Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 298–418. Slijper, E.J., 1962, Whales, Cornell University Press, Ithaca, NY, 511 pp. Spraker, T.R., 1995, Multifocal necrotizing myopathy of northern fur seals (Callorhinus ursinus) from St. Paul Island, Alaska, in Proceedings of the American Association of Zoo Veterinarians/American Association of Wildlife Veterinarians, Aug. 12–17, East Lansing, MI, 505. Spraker, T.R., in press, Pathological findings in northern fur seals from the Pribilof Islands, 1986–2000, NOAA Technical Memo, National Marine Mammal Laboratory, Seattle, WA. Spraker, T.R., Lowry, L.F., and Frost, K.J., 1994, Gross necropsy and histopathological lesions found in harbor seals, in Marine Mammals and the Exxon Valdez, Academic Press, San Diego, CA, 281–311. Stedham, M.A., Casey, H.W., and Keyes, M., 1977, Lymphosarcoma in an infant northern fur seal, J. Wildl. Dis., 13: 176–179.
0839_frame_C23 Page 546 Tuesday, May 22, 2001 11:52 AM
546
CRC Handbook of Marine Mammal Medicine
Steiger, G.H., Calambokidis, J., Cubbage, J.C., Skilling, D.E., Smith, A.W., and Gribble, D.H., 1989, Mortality of harbor seal pups at different sites in the inland waters of Washington, J. Wildl. Dis., 25: 319–328. Stephen, C., 1993, Hiatal hernia in a harbor porpoise (Phocoena phocoena), J. Wildl. Dis., 29: 364–366. Stetzer, E., Williams, T.D., and Nightingale, J.W., 1981, Cholangiocellular adenocarcinoma, leiomyoma and pheochromocytoma in a sea otter, J. Am. Vet. Med. Assoc., 179: 1283. Stewart, B.S., and Yochem, P.K., 1987, Entanglement of pinnipeds in synthetic debris and fishing net and line fragments at San Nicolas and San Miguel Islands, California, 1978–1986, Mar. Pollut. Bull., 18: 336–339. Stirling, I., 1969, Tooth wear as a mortality factor in the Weddell seal Leptonychotes weddellii, J. Mammal., 50: 559–565. Stolk, A., 1950, Tumors in whales, Amsterdam Nat., 1: 28–33. Stolk, A., 1952, Some tumors in whales, Proc. K. Ned. Akad. Wet., 55: 275–278. Stolk, A., 1953, Some tumors in whales II, Proc. K. Ned. Akad. Wet., 56: 369–374. Stout, C., 1969, Atherosclerosis in exotic carnivora and pinnipedia, Am. J. Pathol., 57: 673–687. Stroud, R.K., 1978, Esophageal dilation in a harbor seal, Phoca vitulina, J. Zoo Anim. Med., 9: 20–22. Stroud, R.K., 1979, Nephrolithiasis in a harbor seal, J. Am. Vet. Med. Assoc., 175: 924–925. Stroud, R.K., and Roffe, T.J., 1979, Causes of death in marine mammals stranded along the Oregon coast, J. Wildl. Dis., 15: 91–97. Stroud, R.K., and Stevens, D.B., 1980, Lymphosarcoma in a harbor seal (Phoca vitulina richardsii), J. Wildl. Dis., 16: 267–270. Suzuki, M., Ohtaishi, N., and Nakane, F., 1990, Supernumary post canine teeth in the Kuril seal (Phoca vitulina stejnegeri), the larga seal (Phoca largha), and the ribbon seal (Phoca fasciata), Jpn. J. Oral Biol., 32: 323–329. Suzuki, M., Kishimoto, M., Hayama, S., Ohtaishi, N., and Nakane F., 1992, A case of cleft palate in a Kuril seal (Phoca vitulina stejnegeri) from Hokkaido, Japan, J. Wildl. Dis., 28: 490–493. Sweeney, J.C., 1973, Management of pinniped diseases, in Proceedings of the American Association of Zoo Veterinarians, 141–171. Sweeney, J.C., 1974, Common diseases of pinnipeds, J. Am. Vet. Med. Assoc., 165: 805–810. Sweeney, J.C., and Gilmartin, W.G., 1974, Survey of diseases in free-living California sea lions, J. Wildl. Dis., 10: 370–376. Tarasoff, F.J., and Pierard, J., 1970, Ectrodactylism in the harbor seal, Phoca vitulina L. (Mammalia: Phocidae), Can. J. Zool., 48: 1381. Tarpley, R.J., and Marwitz, S., 1993, Plastic debris ingestion by cetaceans along the Texas coast: Two case reports, Aquat. Mammals, 19: 93–98. Tarpley, R.J., Jarrell, G.H., George, J.C., Cubbage, J., and Stott, G.G., 1995, Male pseudohermaphroditism in the bowhead whale, Balaena mysticetus, J. Mammal., 76: 1267–1275. Taylor, D.C., and Greenwood, A.G., 1974, Functional and pathological aspects of the skin of marine mammals, in Functional Anatomy of Marine Mammals, Harrison, R.J. (Ed.), Academic Press, New York, 73–110. Taylor, M.A., 1993, Stomach stones for feeding or buoyancy? The occurrence and function of gastroliths in marine tetrapod, Philos. Trans. R. Soc. London Ser. B, 341: 163–175. Thomas, N.J., and Cole, R.A., 1996, The risk of disease and threats to the wild population. Special Issue: Conservation and management of the southern sea otter, Endangered Species Update, 13: 23–27. Thomas-Baker, B., 1986, Diskospondylitis in a California sea lion, J. Am. Vet. Med. Assoc., 189: 1151. Trasky, L.L., 1976, Environmental impact of seismic exploration and blasting in the aquatic environment, Report for Alaska Department of Fish and Game, Anchorage, 23 pp. Trillmich, F., Ono, K.A., Costa, D.P., DeLong, R.L., Feldkamp, S.D., Francis, J.M., Gentry, R.L., Heath, C.B., LeBoeuf, B.J., Majluf, P., and York, A.E., 1991, The effects of El Niño on pinniped populations in the eastern Pacific, in Pinnipeds and El Niño, Responses to Environmental Stress, Trillmich, F., and Ono, K.A. (Eds.), Springer-Verlag, New York, 247–287.
0839_frame_C23 Page 547 Tuesday, May 22, 2001 11:52 AM
Noninfectious Diseases
547
Truex, R.C., Nolan, F.G., Truex, R.C., Schneider, H.P., and Perlmutter, H.I., 1961, Anatomy and pathology of the whale heart with special reference to the coronary circulation, Anat. Rec., 141: 325–354. Trupkiewicz, J.G., Gulland, F.M.D., and Lowenstine, L.J., 1997, Congenital defects in northern elephant seals stranded along the central California coast, J. Wildl. Dis., 33: 220–225. Turnbull, B.S., and Cowan, D.F., 1998, Myocardial contraction band necrosis in stranded cetaceans, J. Comp. Pathol., 118: 317–327. Turnbull, B.S., and Cowan, D.F., 1999, Synovial joint disease in wild cetaceans, J. Wildl. Dis., 35: 511–518. Uys, C.J., and Best, P.B., 1966, Pathology and lesions observed in whales flensed at Saldanha Bay, South Africa, J. Comp. Pathol., 76: 407–412. Walsh, M.T., Beusse, D., Bossart, G.D., Young, W.G., O’Dell, D.K., and Patton, G.W., 1988, Ray encounters as a mortality factor in Atlantic bottlenose dolphins (Tursiops truncatus), Mar. Mammal Sci., 4: 154–162. Waring, G.T., Gerrior, P., Payne, P.M., Parry, B.L., and Nicolas, J.R., 1990, Incidental take of marine mammals in foreign fishery activities off the northeast United States, Fish. Bull. U.S., 88: 347–360. Watson, A.G., and Bonde, R.K., 1986, Congenital malformations of the flipper in three West Indian manatees, Trichechus manatus, and a proposed mechanism for development of ectrodactyly and cleft hand in mammals, Clin. Orthop. Relat. Res., 202: 294–301. Watson, A.G., Stein, L.E., Marshall, C., and Henry, G.A., 1994, Polydactyly in a bottlenose dolphin, Tursiops truncatus, Mar. Mammal Sci., 10: 93–100. Watson, W.S., Sumner, D.J., Baker, J.R., Kennedy, S., Reid, R., and Robinson, I., 1999, Radionucleotides in seals and porpoises in coastal waters around the U.K., Sci. Total Environ., 234: 1–13. Wells, C., and Lawrence, P., 1976, A pathological dolphin, Med. Biol. Illus., 26: 35–37. Wells, R., and Scott, M., 1997, Seasonal incidence of boat strikes on bottlenose dolphins near Sarasota, Florida, Mar. Mammal Sci., 13: 475–480. Wiig, O., Derocher, R.E., Cronin, N.M., and Skaare, J.U., 1998, Female pseudohermaphrodite polar bears at Svalbard, J. Wildl. Dis., 34: 792–796. Wilkinson, I.S., Childerhouse, S.J., Duignan, P.J., and Gulland, F.M.D., 2000, Infanticide and cannibalism in the New Zealand sea lion, Phocarctos hookeri, Mar. Mammal Sci., 16: 494–500. Williams, T.D., and Pinard, W., 1983, Pneumoperitoneum associated with intestinal volvulus in a sea otter, J. Am. Vet. Med. Assoc., 183: 1288–1289. Williams, T.D., and Pullet, L.T., 1981, Leiomyomas in two sea otters, Enhyda lutris, J. Wildl. Dis., 17: 401. Williams, T.D., Farwell, C.J., Soule, J.D., and Ticer, J., 1987, Volvulus in a sea otter, Calif. Vet., 5: 97. Wilson, B., Thompson, P.M., and Hammond, P.S., 1997, Skin lesions and physical deformities in bottlenose dolphins in the Moray Firth: Population prevalence and age-sex differences, Ambio, 26: 243–247. Wilson, T.M., 1972, Diffuse muscular degeneration in captive harbor seals, J. Am. Vet. Med. Assoc., 161: 608. Woodhouse, C.D., and Rennie, C.J., 1991, Observations of vaginal calculi in dolphins, J. Wildl. Dis., 27: 421–427. Zhu, Q., 1997, First record of an eye disease in a bowhead whale (Balaena mysticetus), Chin. J. Oceanol. Limnol., 15: 192–193. Zinchenko, V.L., and Ivashin, M.V., 1987, Siamese twins of minke whales of the Southern Hemisphere, Sci. Rep. Whales Res. Inst., 38: 165–169.
0839_frame_C23 Page 548 Tuesday, May 22, 2001 11:52 AM
0839_frame_C24 Page 549 Tuesday, May 22, 2001 11:53 AM
V
Diagnostic Imaging in Marine Mammals
0839_frame_C24 Page 550 Tuesday, May 22, 2001 11:53 AM
0839_frame_C24 Page 551 Tuesday, May 22, 2001 11:53 AM
24 Overview of Diagnostic Imaging William Van Bonn and Fiona Brook
Introduction The problem-oriented approach to clinical practice is largely a process of information gathering. The astute practitioner will progress from patient signalment (i.e., species, age, gender, reproductive status) to current and past medical histories, through complete physical examination and minimum data collection, to special studies. Along the way, the practitioner gathers enough information to reach an accurate diagnosis and formulate an effective treatment plan. Following is a series of eloquent quotes on the value and contribution of diagnostic imaging (taken from the Mayo Biomedical Imaging Resource Analyze® software Web page and printed here with permission) by Dr. R. A. Robb. Human vision provides an extraordinarily powerful and effective means for acquiring information. Much of what we know about ourselves and our environment has been derived from images produced by various instruments, ranging from microscopes to telescopes which extend the range of human vision into realms beyond that which is naturally accessible. In addition to art and aesthetics, images have profound scientific significance and value. More than “pretty” pictures, science needs “useful” pictures. Recent development of advanced quantitative methods to fully analyze the intrinsic information contained in images has begun to unearth the rich treasures they contain and to exploit their full scientific, educational and/or biomedical value.
Imaging Science The process of forming an image involves the mapping of an object or set of objects, and/or some properties of the object(s), into or onto what is called image space. This space is used to visualize an object and its properties, and may be used to quantitatively characterize its structure and/or its function. Imaging science may be defined as the study of these mappings and development of ways to better understand them, to improve them and to productively use them. Generally, the steps involved in imaging procedures include image-data acquisition, image display and analysis (including evaluation of image quality) and image observation (i.e., perception, cognition, and interpretation). Most modern imaging devices are digital computer-based, and produce an image in the form of an array of picture elements (pixels). Emerging imaging modalities produce three-dimensional images of volume elements (voxels). The numbers associated with these pixels and voxels represent the mappings of object properties that can be detected and localized spatially, and which quantitatively characterize these properties. 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
551
0839_frame_C24 Page 552 Tuesday, May 22, 2001 11:53 AM
552
CRC Handbook of Marine Mammal Medicine
The challenge of imaging science is to define and facilitate development of advanced capabilities for acquisition, processing, visualization and quantitative analysis of images in order to significantly increase the faithful extraction of the scientific, educational, and/or clinical information which they contain. This is a formidable task, one which consistently suggests that continued advances are required to address it effectively. The need for new approaches to image analysis have become increasingly important and pressing as advances in imaging technology enable more complex objects and processes to be imaged and simulated. The imaging modalities used in biology and medicine are based on a variety of energy sources, including light, electrons, lasers, x-rays, radionuclides, ultrasound and nuclear magnetic resonance. The images produced span orders of magnitude in scale, ranging from molecules and cells to organ systems and the full body. The advantages and limitations of each modality are primarily governed by the basic physical and biological principles, which influence the way each energy form interacts with tissues, and by the specific engineering implementation for a particular medical or biological application. The variety of disease processes and abnormalities affecting all regions of the human body are so numerous and different that each imaging modality possesses attributes that make it uniquely helpful in providing the desired understanding and/or discrimination of the disease or abnormality, and therefore no single method has prevailed to the complete exclusion of others. In general, the methodologies are complementary, together providing a powerful armamentarium of clinical diagnostic, therapeutic and biomedical research capabilities which has potential to significantly advance the practice of medicine and the frontiers of biological understanding. The imaging scientist resonates with the motivation of Matisse, but with a shift from the art to the science: “I want to reach a point of condensation of measurements to make a useful picture.” *
From Human to Marine Mammal Diagnostic Imaging Veterinary practitioners today are able to enjoy the fruits of this continuing development of the same advanced imaging technologies. Never before has it been easier to obtain critical information about the internal structure and function of patients, using such relatively noninvasive methods as radiography, computed tomography (CT), magnetic resonance imaging (MRI), ultrasonography, endoscopy, and thermography. Undoubtedly, the new millennium will bring with it many significant advances in imaging technology. Those entrusted with the care of marine mammals must stay current, and strive to apply these advances to their endeavors. They owe it to the animals. There are several limitations to the application of diagnostic imaging techniques in the practice of marine mammal care. Most of the equipment in use today for image acquisition requires an alternating current (AC) power source and has been designed for use in sheltered diagnostic suites—not out-of-doors, in ambient light, near saline environments, or sometimes obstreperous patients. The use of delicate, expensive, and often bulky electronic equipment in varied exhibit or field environments is not without significant risk to patient, personnel, and equipment alike. Transportation, assembly, and use of such equipment often require careful advanced planning to minimize this risk. Often in the unique environs and situations encountered working with marine mammals, creative solutions are part of the problem solving. The authors of the chapters in this diagnostic imaging section (Chapters 24 through 28) have included suggestions and solutions to these limitations and challenges, based on experience in the use of each of the respective techniques with marine mammals. * Quotations from R. A. Robb, Mayo Biomedical Imaging Resource Analyze® software Web page. With permission.
0839_frame_C24 Page 553 Tuesday, May 22, 2001 11:53 AM
Overview of Diagnostic Imaging
FIGURE 1 Montage of diagnostic imaging modalities. (Image credits: U.S. Navy, S. Dover, and M. Walsh.)
553
0839_frame_C24 Page 554 Tuesday, May 22, 2001 11:53 AM
554
CRC Handbook of Marine Mammal Medicine
Application of Diagnostic Imaging Techniques There are several important concepts that apply to the application of any diagnostic imaging technique. Standardization, documentation, and archiving are central to the maximum use of image data. In an ideal situation, all imaging studies with a given technique would be performed with the same equipment and settings, and with the patient in the same orientation with respect to the energy source and any receiving devices. Such standardization optimizes the precision (repeatability) of any study. Expected “normal” images can be produced more quickly, and unexpected “abnormal” images are more easily recognizable. Atlases of anatomy by species and technique can be readily compiled, and provide invaluable references for practitioners unfamiliar with an imaging technique or the anatomy of interest (see Chapter 9, Anatomy). This ideal, of course, is not always possible; yet, it is a standard for which all those performing diagnostic imaging must aim. Standardization of patient positioning and image orientation is very important. At the very least, this can be practiced at the individual level, so that one always produces and reviews an image in the same way. This allows for accurate comparison of images over time in an individual patient, and facilitates interpatient comparisons. Such comparisons are often extremely valuable in the interpretation of equivocal images, and are particularly important where there is a paucity of data or reference images. If unsure about whether a finding in an image is normal or not, a duplicate image using a control (a clinically normal animal or matched body part) can be produced and compared. Standardization at the facility level ensures that interpretation of images produced by colleagues at that facility is more consistent and reliable. At the “community” level, standardization allows for greater sharing of image data and interpretation by consult. Everyone benefits. All medical images should include more image data than just a map of the tissues of interest. The patient/subject signalment, date, time, and location of image capture, equipment and technique used, and pertinent transducer orientations should be permanently affixed to the image display. These data are critical for interpretation, accurate archiving, and retrieval of image data, and allow for sound retrospective investigations. Most contemporary diagnostic imaging systems have means that allow for the inclusion of these data in the product image. Often it may only be necessary to enter the patient signalment manually, while other data are recorded in default settings customizable by the operator. Even consumer-oriented video recording devices such as “camcorders” are valuable for real-time capture of dynamic imaging studies such as endoscopy or sonography. These devices generally have time/date stamp capabilities and character generators. The use of these features to include pertinent study information on the permanent image product is prudent practice. Regardless of the means, analog or digital images should be produced during all diagnostic procedures. If the image is not recorded, it might as well not have been collected. Images captured during survey examination of clinically normal animals are especially valuable resources for normal anatomy references. Individual animal medical records should include all images collected, static or dynamic. Contemporary image management software and devices allow for easy transmittal of image data from one locale to another. A picture can truly be “worth a thousand words,” and in this age of e-mail, scanners, and the World Wide Web, image data can be transmitted as easily as text or verbal descriptors. The days of descriptions being limited to “raised, red, with irregular margins” are gone—now supplanted with the more valuable “see attached TIFF file.” An efficient method of image data archiving and retrieval is essential to optimal patient care and facilitates continued improvement in the knowledge, methods, and skills required to master each modality. Review of image data often reveals information overlooked at initial interpretation. In field situations, ambient light and glare often mask data. Reviewed in a dimly lit room, these
0839_frame_C24 Page 555 Tuesday, May 22, 2001 11:53 AM
Overview of Diagnostic Imaging
555
data may become more apparent. Immediate review during the imaging procedure is a valuable learning tool when practicing a new technique. Subtle effects of transducer, scope, or patient movement, for example, are never easier to interpret than immediately after the procedure of collection, when these actions are fresh in the memory of the operator. However, complete interpretation of images should not be rendered until thorough review under optimal conditions. It is especially useful to include audio tracks on dynamic image archives, such as videotapes. Complete patient records, easily retrievable and searchable, aid the rapid assessment of changes in patient condition over time, as well as comparative studies of other animals with similar presentations. Inclusion of image data in patient records greatly increases the amount of information that can be used in formulation of diagnostic and treatment plans. Contemporary computer-based record-keeping systems allow for powerful queries of this information at a moment’s notice. The ability quickly to retrieve matched, high-quality normal images to compare with a presenting patient cannot be overemphasized. A thorough understanding of the physics of the energy source and its interaction with tissues and the receptor, along with image display, and capture methods used for any modality, is critical to optimum performance of the studies and maximal information acquisition. All the hardware and software elements of imaging equipment and systems available today are too numerous to detail here. Readers are encouraged to consult the available literature and documentation specific to the systems at their disposal to maximize efficiency of use. For example, it makes little sense to publish technique charts for one particular X-ray tube-head, screen, and film combination when such a variety of devices are available and in use. Every chart needs to be customized for a particular facility, user, and application, but standardized, documented, and archived. The chapters that follow in this section assume a good working knowledge of these basics. Therefore, the focus is on the applications of specific technologies to marine mammal patients, along with the unique challenges that each may bring. Finally, the imaging of necropsy or other cadaver specimens is encouraged. Post-mortem examination of bodies, organs, and tissues is safely and easily carried out in most clinical facilities. One can easily compare modalities and techniques using the same specimen to gain a more complete working understanding of the pertinent clinical anatomy. Radiographs of necropsy-harvested tissues, or selected regions of whole animals, can be produced using high-contrast techniques requiring long exposure times without being hampered by patient motion or regard for patient safety. Sonograms produced by waterbath submersion of specimens can be produced with higher-resolution (higher-frequency) transducers and without the limitations of in situ acoustic windows. Fresh post-mortem specimens provide excellent opportunities to learn endoscopic and laparoscopic approaches and anatomy. Even archived fixed or frozen organs and tissues are valuable resources for imaging experimentation. For obvious reasons, it is much easier to obtain post-mortem CT and MRI images of marine mammal patients than it is to obtain these images with a live patient. Images produced by such studies are of high value as reference material and should be produced whenever possible.
Conclusion Diagnostic imaging remains one of the most powerful techniques available to the clinician. Each of the tools available has specific indications and limitations. They are directly complementary to each other, and to other sources of information in patient management. Not all pretty pictures are useful, but all useful pictures are beautiful.
0839_frame_C24 Page 556 Tuesday, May 22, 2001 11:53 AM
556
CRC Handbook of Marine Mammal Medicine
Acknowledgments The authors thank Patrick Moore and Sam Ridgway for review of the text, Mark Todd for image-processing expertise, and the many veterinary students they have hosted who have prompted the authors to stay current. The authors also wholeheartedly thank the animals that have cooperated with them in development of contemporary imaging methods.
0839_frame_C25 Page 557 Tuesday, May 22, 2001 11:53 AM
25 Radiology, Computed Tomography, and Magnetic Resonance Imaging William Van Bonn, Eric D. Jensen, and Fiona Brook
Introduction The use of X-rays in the clinical setting is certainly not new to marine mammal practice. There are several published descriptions of techniques and images that focus on clinical care of pinniped and cetacean species, including the first edition of this book (Morris, 1990; Sweeney, 1990). Nonetheless, most published accounts of radiology in marine mammal species are descriptions of research applications; clinical methods have not been standardized to the extent they have in domestic animal species (Felts and Spurrell, 1966; Sommer et al., 1968; Viamonte et al., 1968; Kooyman et al., 1970; Sumner-Smith et al., 1972; Bryden and Felts, 1974; Sweeney and Ridgway, 1975; Ogden et al., 1981; Ponganis et al., 1992). This chapter describes methods that have proved effective in producing diagnostic radiographs of dolphins, sea lions, and seals. The establishment of standard methods of image capture, display, and interpretation is encouraged. Standard radiographs today can be digitized by photographing them with a digital camera. Films are easily digitized by photographing them while viewed on a standard light box. The use of a light stand, tripod, and delayed shutter activation will avoid motion artifact. This method is valuable to the practitioner and useful in applying postprocessing techniques to standard films. Contemporary imaging software packages such as Scion Image for Windows® (Scion Corporation, Frederick, MD), Adobe PhotoShop® (Adobe Systems, Inc., Seattle, WA), Corel Draw® (Corel Corporation, Ottawa, Ontario, Canada), and Sony Picture Gear Lite® (Sony Corporation, Tokyo, Japan) allow the clinician to apply techniques to images that enhance the diagnostic information available. Even simple contrast and brightness adjustments will enhance the data, and often bring out diagnostically useful information otherwise not apparent to the naked eye. Computer radiography (CR) is now commonly used for examination of human patients (Artz, 1997). The basic principles of CR are the same as those of conventional radiography; the main difference is that a reusable, storage phosphor imaging plate that can be photostimulated is used as the image receptor, instead of the standard film/screen combination (Kantor, 1997). The imaging plates are mounted inside a cassette, and are used in the same way. When the imaging plate is exposed to X-rays, phosphor electrons are excited to various energy levels and a latent image is formed on the plate. The imaging plate is then inserted into a “reader,” 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
557
0839_frame_C25 Page 558 Tuesday, May 22, 2001 11:53 AM
558
CRC Handbook of Marine Mammal Medicine
where it is scanned by a red laser, which induces photostimulated luminescence. This light is collected and converted into electronic voltage by photomultiplier tubes. An analog-todigital converter (ADC) converts the electronic signals into digital data, which can be stored in a computer. CR images can then be viewed directly on a monitor, or printed to produce “hard” copies similar to conventional radiographs. CR offers considerable potential advantages over conventional radiography for imaging marine mammals. As for other computerized imaging modalities, such as computed tomography (CT), it allows the use of postprocessing manipulation procedures, including interactive windowing, tonal reversal, and image filtering, to enhance particular features. The imaging plates are reusable and their linear response provides a wider dynamic range and higher contrast resolution than standard film. This increased range and contrast allow for a reduction in the exposure required, which improves image quality in larger patients. Although the spatial resolution of CR images is poorer, the increased contrast and edge enhancement more than compensate, particularly when imaging the dolphin’s lungfields (Figures 1A and B). Studies are under way to evaluate this technique more fully, and further investigation is required to assess whether or not it offers the same advantages in animals larger than dolphins. Newer technologies of CT and magnetic resonance imaging (MRI) are less likely than radiology to be familiar to veterinary practitioners. These diagnostic imaging tools are becoming more available, and a brief introduction to their physics and language may prove informative and promote their use. The authors are not nuclear physicists, and although they have used these techniques in practice, they are not specialists in radiology, CT, or MRI. They hope, however, that their clinical explanation of these emerging methods will be useful to others with opportunities to utilize these methods in providing for marine mammals. CT, arguably the biggest advance in diagnostic imaging since Roentgen’s discovery of X-rays, was developed by a British electronics engineer, Godfrey Hounsfield, in 1967. The first installation and use of a medical CT scanner was at the Mayo Clinic in 1973. Today, the speed, flexibility, and resolution of the technique have improved dramatically, and CT scanners are in common usage in the human medical community. Although the size, expense, and technical requirements of the scanners limit their use by veterinarians, they are becoming increasingly available. The scientific literature includes several papers describing the use of CT in experimental or anatomical investigations of four pinniped species and three cetacean species (Nordoy and Blix, 1985; Hillmann, 1991; Woodhouse and Rennie, 1991; Ponganis et al., 1992; Endo et al., 1999a). In addition, although there are unique limitations presented by marine mammal patients, CT scans of several clinical conditions have been successfully produced (Haulena et al., 1998; Van Bonn, unpubl. data). CT is a specialized imaging technique that combines X-ray production, projection, and reception with computerized digital signal processing to produce a clinical image data set. An array of sensors and an X-ray source are rotated around the patient or specimen to be imaged, resulting in data collection from multiple incident angles. A microprocessor subjects the data to algebraic reconstruction, and displays the output as two-dimensional “slice” images. The source, beam collimators, and receptors are mounted on a mechanical frame called a gantry. For medical diagnostic applications, the patient is positioned within an opening in the gantry between the source and receptor arrays. Because the X-ray beam is projected at multiple angles around the patient’s body and the data integrated, the technique is not subject to the limitations of summation and silhouette experienced in conventional radiology. This tool has also been widely recognized to have much improved detection rates of pathological conditions, when compared with conventional radiology. For more detailed information on CT scanning, refer to: http://www.radiologyresource.org/content/ct_of_the_body.htm
0839_frame_C25 Page 559 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
559
A
B
FIGURE 1 (A) D-V computerized radiographic image of the thorax of a bottlenose dolphin (Tursiops aduncas). (B) Lateral computerized radiographic image of the thorax of a bottlenose dolphin (Tursiops aduncas.) (Image credits: F. Brook.)
The digital output of a CT scan can be displayed on a monitor or sent to a film recorder to produce “hard-copy” films. The digital nature of the data is perhaps the most valuable aspect of the technology. CT brings a third dimension to diagnostic X-rays. Image analysis software routines allow the data to be “postprocessed” in a myriad of manipulations, resulting in the ability to perform three-dimensional constructions, subtraction techniques, colorizations to enhance image data of interest, and more.
0839_frame_C25 Page 560 Tuesday, May 22, 2001 11:53 AM
560
CRC Handbook of Marine Mammal Medicine
MRI is the medical application of nuclear magnetic resonance (NMR) spectroscopy. The technique has its origin in structural determination methods of organic chemistry. MRI does not involve ionizing radiation of any type. Rather, MRI employs powerful magnetic fields and radio frequency electromagnetic energy to produce images. All nuclei with odd-numbered masses, or even-number masses but odd atomic numbers, exhibit magnetic properties; that is, they behave as though they are spinning about an axis. Normally, the orientation of the spin, and therefore the magnetic moment, is random. However, when subject to sufficiently powerful magnetic fields, the nuclei will align. If, while aligned, the nuclei are exposed to appropriate radio frequency electromagnetic pulses, they will absorb energy and achieve “resonance” with the frequency (Fessenden and Fessenden, 1979). These resonance shifts can be detected by sensitive radio frequency receivers and mapped to the image space as it corresponds to their position. Like CT scans, the output of MRI scans is digital, and can be extensively postprocessed. Unlike CT, the image data largely reflect the behavior and position of protons within water molecules in the tissues. The strength of an MRI signal depends on three parameters: proton density, T1 (the time it takes for resonating nuclei to return to an equilibrium state parallel to the magnetic field), and T2 (the time to an equilibrium state perpendicular to the magnetic field). The strength is directly proportional to the proton density of the tissue. These times (T1 and T2) differ markedly for different tissue types, and thus these properties are responsible for the excellent soft-tissue contrast of MRI images. The digital output of MRI scans is said to be “weighted” to emphasize one or the other of the above properties. Readers with interest in learning more of the physics of MRI applications are encouraged to consult the following resource sites http://www.cis.rit.edu/htbooks/mri/inside.htm http://www.med.harvard.edu/AANLIB/sigsors.html
Most CT and MRI scanners are fixed installations at human medical centers, although “portable” truck-mounted scanners and magnets do exist. For most facilities caring for marine mammals, the availability of these technologies is quite restricted. However, as they become increasingly routine in human clinical use, it is expected they will continue to be more and more available to veterinarians working with marine mammals. Even conventional X-ray equipment and facilities are limited in many veterinary practices, thus establishing a working relationship with local human medical centers is a good way to gain potential access to diagnostic imaging equipment and expertise. There are, of course, numerous restrictions to the use of human hospital resources for animal imaging, but one can often find personnel willing to accommodate “late-night” scans or radiography of marine mammal patients. Combining the expertise and equipment of these facilities with the specialized knowledge of the husbandry, handling, anatomy, physiology, and anesthesia of marine mammals possessed by the attending veterinarian is valuable. This approach can be an extremely rewarding source of information in the clinical management of these animals. In spite of their limited availability and other limitations outlined later, the value of CT and MRI data sets cannot be overemphasized. Their superior quality and the additional information they provide make them extremely valuable reference sets for use when performing other diagnostic imaging techniques. The availability of CT or MRI scans when performing sonography or surgical planning, for example, is perhaps their greatest immediate use for marine mammal practitioners. Scans of cadaver specimens are worth the effort and cost to obtain for use as clinical references to guide other diagnostic, therapeutic, and experimental techniques and procedures.
0839_frame_C25 Page 561 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
561
Indications The most frequent radiographic procedure in the authors’ practice is thoracic radiology of dolphins. Radiographs are indicated any time that respiratory disease is suspected, especially when sonograms fail to demonstrate abnormalities. Radiographs are also extremely valuable for assisting with prognosis determination of those cases where pathology is apparent on sonograms of the thorax. Pulmonary disease is one of the most common suspected and/or confirmed causes of morbidity and mortality in dolphins (see Chapter 40, Cetaceans). Sonography and endoscopy can provide very valuable diagnostic information about the lungs, airways, lymph nodes, and other intrathoracic anatomy; but X-rays are the only readily available means of imaging the lung parenchyma and small airways, and the only currently readily available, reliable method of imaging the entire heart. One specific indication for thoracic radiographs is to overcome an inherent limitation of sonography. Sonography of intrathoracic anatomy is limited by reverberation artifact created by the large acoustic impedance mismatch of the soft tissues and ventilated lung parenchyma. If air-filled lung is present in the plane of scan between the transducer and a lesion, the lesion cannot be imaged with sonography. Transesophageal sonography will sometimes overcome this limitation, but not always, and it has its own limitations (see Chapter 26, Ultrasonography). Properly produced radiographs will visualize the region of interest and often demonstrate important changes due to disease of the structures deep within the thorax. Figures 2 and 3 illustrate this limitation of sonography and the importance of radiology to evaluate deep thoracic anatomy. Radiography of the pinniped thorax is less commonly performed in the authors’ practice, but the indications and advantages over other imaging modalities are the same. Plain radiography (survey radiography) of the abdomen is much less frequently indicated, because of the limitations of the technique discussed below. Positive contrast (barium, iodinated media) and negative contrast (gases) studies are useful to evaluate motility of the intestinal tract, and in situ position of abdominal viscera (Figures 4 and 5). This technique is indicated
FIGURE 2 Dorsal plane sonogram of left lung of 180-kg male bottlenose dolphin (Tursiops truncatus). Note prominent pleural reverberation artifact. (Image credit: U.S. Navy.)
0839_frame_C25 Page 562 Tuesday, May 22, 2001 11:53 AM
562
CRC Handbook of Marine Mammal Medicine
FIGURE 3 D-V radiograph of left hemithorax of the same animal as in Figure 1. Note deep foci of increased radiopacity and normal air-filled lung at surface. (Image credit: U.S. Navy.)
whenever intraluminal or extraluminal space-occupying lesions are suspected, but not imaged with endoscopy or sonography. Sonography is a superior imaging technique for the abdomen of marine mammal species, and endoscopy is a superior method to evaluate the forestomach lumen (see Chapter 27, Endoscopy). The most common indications for taking bone films in marine mammals are assessments of the head and jaws of cetaceans and the head, jaws, and extremities of pinnipeds. Whenever lesions of skeletal structures are included in the rule-out list, a radiograph of the region should be produced using high-contrast technique to illustrate bone. In the authors’ experience, fractures, osteomyelitis, spondylosis, and foreign bodies are the most frequently detected lesions. Reduced-contrast “soft-tissue” techniques may be useful in demonstrating soft-tissue lesions
0839_frame_C25 Page 563 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
563
FIGURE 4 Lateral positive-contrast radiograph of 225-kg female bottlenose dolphin. Image was captured approximately 47 min after feeding liquid barium–injected fish. Note fluid lines in forestomach, and fundic and pyloric chambers. (Image credit: U.S. Navy.)
such as fistulae, sinuses, or abscesses in these skeletal regions. The use of positive-contrast media to highlight the extent of the abscess, fistula, or sinus is frequently the most effective method of imaging these conditions. The use of positive-contrast media is useful to illustrate the orientation of vascular structures. Angiography has most often been used as a research tool in marine mammal species rather than as a clinical tool. A very eloquent and elaborate description of the somatic thoracic and cranial circulation of the living Atlantic bottlenose dolphin ( Tursiops truncatus) was performed using positive-contrast angiography (Viamonte et al., 1968). Figures 6 and 7 were produced by Dr. Forrest Townsend. Both contrast studies used 61% Iopamidol® (Bracco Diagnostics, Inc., Princeton, NJ) to illustrate vascular networks. The dorsal fin in Figure 5 was obtained post-mortem from an adult Atlantic bottlenose dolphin, while Figure 6 illustrates the vasculature of an unborn, near-term, dolphin calf, after infusion of contrast media into the aorta. The use of contrast agents to explore the route and/or extent of draining tracts and sinuses can be very useful in clinical situations. CT and MRI, when available, are superior techniques for imaging internal soft-tissue anatomy. They are indicated whenever sufficient diagnostic information is not available by using conventional radiology or ultrasonography. The use of MRI in marine mammal investigations has been limited largely to experimental use with post-mortem specimens (Hillmann, 1991; Matassa et al., 1994; Endo et al., 1999b). Images produced by MRI scans of post-mortem specimens are extremely valuable as reference materials to guide the use of other imaging modalities.
0839_frame_C25 Page 564 Tuesday, May 22, 2001 11:53 AM
564
CRC Handbook of Marine Mammal Medicine
FIGURE 5 Ventrodorsal radiograph of 115-kg castrated male California sea lion (Zalophus californianus) under general anesthesia. Image was captured approximately 30 min after administration of liquid barium via stomach tube. Note electrocardiographic electrodes and leads. (Image credit: U.S. Navy.)
FIGURE 6 Lateral positive-contrast angiogram of cadaver dorsal fin of mature bottlenose dolphin (Tursiops truncatus). (Image credit: F. Townsend.)
0839_frame_C25 Page 565 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
565
FIGURE 7 Lateral positive-contrast angiogram of fetal bottlenose dolphin (Tursiops truncatus). A radiopaque urinary catheter has been sutured in place in the dorsal aorta. (Image credit: F. Townsend.)
Limitations The use of all X-ray machines poses some radiation exposure hazard to personnel. The radiation area during operation must be carefully defined. Only qualified, knowledgeable, and conscientious individuals may operate the equipment or assist. Personal protection and dosimetry monitoring must be practiced. Time, distance, and shielding concepts must be considered in the radiation safety plan. For a good review of overall radiation safety and safety plan development, readers are encouraged to consult Barber and Lewis (1982). The authors are not aware of any currently used techniques for obtaining radiographs, CT scans, or MRI images of marine mammals while the animals are in water. The attending clinician must carefully weigh the benefits against the risks of removing animals from the water to obtain any diagnostic images. The risks include respiratory compromise, disturbances of thermoregulation, significant preexisting disease, and psychosocial stress, especially in animals with limited experience of being handled out of water. The techniques described in this chapter are used only when other modalities have not resulted in a diagnosis. A technique chart used at one facility or by one practitioner is often totally inappropriate for use by another. The development of a specific chart under the specific conditions of use by the specific facility ensures the production of high-quality diagnostic images with minimum time, effort, and risk to patient and personnel. Portable handheld diagnostic X-ray units have voltage (kVp) and current (mAs) capabilities that limit the size of the body or body part that can be imaged. The adult size of many marine mammal species will exceed these limits. The use of portable systems at locations remote from film-processing equipment creates the need for careful planning. One must plan exposure and technique to avoid excessive patient restraint time or time out of water, and to avoid repeated trips to and from film-processing areas. These machines are designed primarily for large-animal extremities in barns or clinics, not for use in or around open-water environments. Care must be taken to avoid damage to the equipment and to minimize electrical shock hazard to patients and personnel.
0839_frame_C25 Page 566 Tuesday, May 22, 2001 11:53 AM
566
CRC Handbook of Marine Mammal Medicine
Large “hospital-type” portable units are capable of imaging much larger animals and body parts. However, these units are less easily moved to remote sites. Fixed-site units are often capable of imaging large dolphins, pinnipeds, and even small whales, although those units require the patient to be brought to the machine. The close proximity to film-processing equipment often outweighs the risk associated with patient movement in well-planned facilities with appropriate patient support. With careful attention to the principles of safe animal transport (see Chapter 39, Transport), it is often more efficient to move the patient to a facility with adequate-size machines and on-site film-processing capabilities. Film screen combinations are generally not designed for use in and around open-water environments either. Cassettes must be protected from contamination with water used to keep animals wet during exposures. The authors’ facility has had good success with a custom-made clear bag of radiolucent plastic within which the cassette is placed during exposures. Pinnipeds, unless trained for the purpose, will often not remain still long enough for appropriate positioning and exposure. Moderate sedation or general anesthesia is frequently required (see Chapter 29, Anesthesia) to avoid motion artifact and obtain high-quality diagnostic films. Motion artifact is usually not a problem in cetacean species. Most cetaceans out of the water can be safely restrained, and the normal breath hold during the respiratory cycle reduces complications due to movement during inspiration and expiration. The many and varied limitations of individual facilities and equipment combinations have hindered the standardization of radiographic positioning of marine mammals. There are several published radiographs of a variety of marine mammal species, including those in the first edition of this book, but detailed anatomical interpretation is lacking. A given facility or practitioner should strive to standardize studies as much as possible. This will allow for at least in-house comparisons A
B
FIGURE 8 (A) Dorsolateral-ventromedial oblique radiograph of 90-kg male bottlenose dolphin (Tursiops truncatus) right pectoral flipper. Compare with Figure 8B. (B) Dorsolateral-ventromedial oblique radiograph of 90-kg male bottlenose dolphin (Tursiops truncatus) left pectoral flipper. Note degenerative joint disease of scapulo–humeral, humeral–radial, and humeral–ulnar joints. Compare with Figure 8A. (Image credits: U.S. Navy.)
0839_frame_C25 Page 567 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
A
567
B
FIGURE 9 (A) Lateral radiograph of skull section of bottlenose dolphin ( Tursiops truncatus) cadaver. Note radiopaque catheter within nasal passage, auditory tube, and middle ear. (B) D-V radiograph of cervical and rostral thoracic vertebrae of bottlenose dolphin ( Tursiops truncatus) cadaver. (Image credits: U.S. Navy.)
of images. The use of a control, clinically normal animal or a normal paired body part, is valuable when interpreting suspect lesions (Figures 8A and B). Radiology of necropsy specimens, followed by dissection, is another powerful technique that may be useful in support of anatomical or clinical investigations (Figures 9A and B). Even with fixed machines capable of high kVp and mAs outputs, it is difficult to produce diagnostic images of cetacean abdominal soft tissues. Cetaceans have a thick layer of fatladen blubber surrounding an abdomen with little visceral fat. This produces large attenuation of the beam as it enters the abdomen and reduces the contrast. The use of contrast agents for gastrointestinal or urogenital studies can produce valuable films (Figures 10 and 11). Pinnipeds will more often have gas visible on radiographs of the abdomen and more anatomy can be imaged in pinnipeds than in cetaceans; however, survey abdominal radiology is often inconclusive. The gantry and table of most CT scanners are limited to a patient weight of approximately 136 kg (300 lb). The equipment is also subject to water damage; this presents challenges for keeping patients cool and wet during scans. Newer, high-speed CT scanners are now in use and have the capability of scanning approximately half the body length of a small dolphin or porpoise between breaths. Thus, it may be possible to get whole-body CT data
0839_frame_C25 Page 568 Tuesday, May 22, 2001 11:53 AM
568
CRC Handbook of Marine Mammal Medicine
FIGURE 10 Lateral radiograph of caudal abdomen of 180-kg male bottlenose dolphin (Tursiops truncatus). Note lack of contrast and anatomical detail. Also note transitional vertebrae and radiopaque foreign body. (Image credit: U.S. Navy.)
sets of live dolphin patients with two scans, minimal time out of water, and minimal water damage hazard. The largest recognized risk with diagnostic MRI is the projectile effect. Any ferromagnetic object will be attracted to the magnet, and may cause serious trauma to the patient. In human medicine this is usually encountered with shrapnel or implants. This will likely not be a frequently observed effect in marine mammal medicine. However, the clinician is cautioned to be aware of this potential, and screen patients for the presence of ferromagnetic implants or foreign bodies prior to MRI scanning. Magnets also have limited aperture ranges restricting the size of the patient that can be scanned. Newer U-shaped or open magnets may overcome this limitation.
Technique Regardless of the species or anatomy to be imaged, whenever possible two views on perpendicular axes should be produced for every radiographic study. Two views on perpendicular axes will often avoid summation and silhouette sign that may obscure anatomy and pathological lesions, and will localize structures in three dimensions (Figures 12 and 13). When displaying and interpreting films from any species, the procedure should be
0839_frame_C25 Page 569 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
FIGURE 11 Lateral positive-contrast radiograph of 225-kg female bottlenose dolphin (Tursiops truncatus). Image was captured approximately 17 min after oral administration of liquid barium. Compare with Figure 4. (Image credit: U.S. Navy.)
FIGURE 12 D-V radiograph of caudal peduncle of 190kg female bottlenose dolphin (Tursiops truncatus). Note absence of obvious bony change. Compare with Figure 14. (Image credit: U.S. Navy.)
569
0839_frame_C25 Page 570 Tuesday, May 22, 2001 11:53 AM
570
CRC Handbook of Marine Mammal Medicine
FIGURE 13 Lateral radiograph of caudal peduncle of same 190-kg female bottlenose dolphin (Tursiops truncatus) as in Figure 12. Note mineral densities dorsal to spine. Oblique radiodense lines ventral to these densities are artifact created from support sling. Compare with Figure 12. (Image credit: U.S. Navy.)
FIGURE 14 Animal positioned for D-V radiograph in sling. Note minimal personnel requirement. (Image credit: U.S. Navy.)
0839_frame_C25 Page 571 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
571
standardized to promote efficiency and completeness. Radiographic films of marine mammal species should be displayed with the head to the left on lateral views and the left aspect of the animal to the right on the dorsoventral (D-V) view, as is conventional for domestic species (Thrall, 1986). A sling suspension device for radiology of small cetaceans facilitates more rapid procedures that are safer for the patient and personnel who are involved (Figure 14). With the animal suspended in a sling within a frame, similar to those used for transportation, the tube head and cassettes can be moved around the patient and any view required can be obtained. Positioning the animal in sternal recumbency and using a horizontal beam is much preferred, compared with having the animal in lateral recumbency and placing the cassette underneath the animal. This promotes standardization of views, and avoids complications and inconsistencies when repositioning the animal between exposures. Repeated rolling of an animal from side to side is often not well tolerated. Placing conventional cassettes underneath the animal may lead to abrasions and contusions, as well as damage to equipment. For mature animals of most species, generally one hemithorax is imaged at a time on D-V views, and two lateral views are required to image the entire air-filled lung with the animal out of the water. Diagnostic films can be produced by placing the animal on the floor or deck surface, positioning the cassette underneath the animal, and utilizing a vertical beam. For lateral views, the animal is rolled into lateral recumbency with the side of interest down. Although this is possible, and in many facilities necessary, in the authors’ experience this method is more time-consuming, causes more equipment failures, is less well tolerated by the patient, and often leads to inferior image quality. It should be noted, however, that the patient, experiences less thoracic compression than when suspended. This may be clinically important in animals with respiratory disease. When interpreting thoracic radiographs, the clinician must always be cognizant of the technique used. Films produced with patients suspended will show evidence of compression when compared with films produced with patients on the floor or deck (Figures 15A and B). This compression may be of clinical value, since it will increase contrast, all other factors being equal. When interpreting thoracic radiographs of dolphins, the clinician must keep in mind the constraints imposed by the unique anatomy of the cetacean lung and the effects of compression under gravity. Cetacean lungs have a large amount of elastic connective tissue and a cartilage component in all airways to the level of respiratory bronchioles (Simpson and Gardner, 1972; Britt and Howard, 1983). These factors contribute to a prominent interstitial pattern of the normal cetacean lung on radiographs. The clinician must also be aware that the peripheral lung fields of cetaceans out of the water may be atelectic, and cannot be thoroughly assessed with radiographs alone. The caudal sector of the thorax extensively overlaps the cranial sector of the abdomen. The artifacts of summation and silhouette sign obscure the intrathoracic anatomy in much of the caudal sector of the thorax. The technique and interpretation of pinniped thoracic radiographs is essentially the same as that for large dogs. Dental radiographs are useful in managing tooth and jaw diseases of marine mammals. Cetaceans will often tolerate open-mouth oblique views, and cassettes can be placed in the mouth to image individual arcades (Figure 16). Pinnipeds usually require general anesthesia to produce diagnostic dental films. The entire axial and appendicular skeleton of small cetaceans and pinnipeds can be reliably imaged with X-ray devices. Patient positioning and views indicated are dictated by the clinical situation, and should be tailored to promote standardization and accommodate the special requirements of marine mammals outlined in the above sections.
0839_frame_C25 Page 572 Tuesday, May 22, 2001 11:53 AM
572
CRC Handbook of Marine Mammal Medicine
A
B
FIGURE 15 (A) D-V radiograph of left hemithorax of 150-kg male bottlenose dolphin (Tursiops truncatus). Image captured with animal suspended in sling. Compare with Figure 15B. (B) D-V radiograph of left hemithorax of same 150-kg male bottlenose dolphin (Tursiops truncatus). Image captured with animal in sternal recumbency on deck. Compare with Figure 15A. (Image credits: U.S. Navy.)
0839_frame_C25 Page 573 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
573
FIGURE 16 Right and left dorsolateral-ventromedial oblique radiographs of mandibular arcades of 100-kg male bottlenose dolphin (Tursiops truncatus). (Image credit: U.S. Navy.)
0839_frame_C25 Page 574 Tuesday, May 22, 2001 11:53 AM
574
CRC Handbook of Marine Mammal Medicine
Clinical Applications To promote standardization of views and assist in diagnostic image collection, the figure icons in this chapter illustrate the position of the X-ray beam with respect to the patient. The cassette is always positioned perpendicular to the beam, and as close to the region of interest as possible. Recall that two views in perpendicular planes should always be produced.
Dolphin Normal Radiographic Anatomy
To produce D-V views of the thorax, with the patient in sternal recumbency, the cassette is positioned ventral to the patient and a vertical beam is used. In all but the smallest animals, each hemithorax is imaged separately. The air-filled lung field extends one to two rib spaces rostral to the shoulder joint at the rostral insertion of the pectoral flipper, and the beam is centered accordingly (Figure 17). The ventilated lung field extends caudal to the level of the dorsal fin, and overlays the cranial sector of the abdomen. It is usually necessary to divide each hemithorax into cranial and caudal sectors to assess the entire lung adequately (Figure 18). To produce lateral views of the thorax, with the patient suspended in sternal recumbency, the cassette is positioned lateral to the patient, and a horizontal beam is used. The cassette is
FIGURE 17 D-V radiograph of left cranial hemithorax of 170-kg male bottlenose dolphin (Tursiops truncatus). Image captured with animal suspended in sling. (Image credit: U.S. Navy.)
0839_frame_C25 Page 575 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
575
FIGURE 18 D-V radiograph of right caudal hemithorax of 170-kg male bottlenose dolphin (Tursiops truncatus). Image captured with animal suspended in sling. (Image credit: U.S. Navy.)
positioned as close as possible along the side of the animal where the suspected pathology exists, to reduce geometric artifact. Again, the lung field is divided into two sectors and images are produced of the cranial and caudal sectors separately. Recall that the dorsal aspect of the trunk of the animal is composed of the epaxial muscle complex, and that the air-filled lung field extends dorsally only to the level of the spine (see Chapter 9, Anatomy). The beam for the two lateral projections is centered accordingly. The tendency is to center the beam too high on the animal, and not on the thoracic cavity itself (Figures 19 and 20). High-quality diagnostic skull films can be obtained with the patient positioned in sternal recumbency on the floor or deck, or with the animal suspended. The cassette is positioned ventral to the head and a vertical beam used. Recall that the temporo-mandibular joint is coincident with the caudal extent of the orbit, and summation and silhouette sign will often complicate interpretation of radiographic anatomy of this region on this projection (Figure 21). Lateral projection images of the skull can also be obtained with the patient on the deck or suspended. If the patient is on the deck and positioned in lateral recumbency, care must be taken to elevate the rostrum so the long axis of the skull is parallel to the plane of the film and perpendicular to the beam. With the animal suspended, a horizontal beam is used. Oblique views of the skull are often useful or required to offset the clinical region of interest (Figure 22).
FIGURE 20 Lateral radiograph of caudal aspect of thorax of mature 170-kg bottlenose dolphin (Tursiops truncatus). (Image credit: U.S. Navy.)
576
FIGURE 19 Lateral radiograph of rostral aspect of thorax of mature bottlenose dolphin (Tursiops truncatus). Note apices of lung fields extending cranial to scapulae. (Image credit: F. Townsend.)
0839_frame_C25 Page 576 Tuesday, May 22, 2001 11:53 AM
CRC Handbook of Marine Mammal Medicine
FIGURE 21 D-V radiograph of skull of 275-kg bottlenose dolphin (Tursiops truncatus). Image was captured with animal in sternal recumbency on deck. (Image credit: U.S. Navy.)
FIGURE 22 Right dorsolateral left ventromedial oblique radiograph of skull of 275-kg bottlenose dolphin (Tursiops truncatus). (Image credit: U.S. Navy.)
0839_frame_C25 Page 577 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging 577
0839_frame_C25 Page 578 Tuesday, May 22, 2001 11:53 AM
578
CRC Handbook of Marine Mammal Medicine
The cranial sector of the abdomen underlies the caudal sector of the thorax. The abdominal field on a lateral projection extends to a point approximately two rib spaces caudal to the caudal insertion of the pectoral flipper. The caudal limit of the abdominal field is the anus. The dorsal aspect of the trunk of the animal comprises the epaxial muscle complex. In the abdomen, the hypaxial muscle complex is positioned ventral to the spine beginning at the level of the dorsal fin. The tendency is to center the beam too high on the patient. The beam must be centered on the abdomen, not the entire animal. Survey D-V projection images of the dolphin abdomen are frequently not of diagnostic usefulness when evaluating intra-abdominal viscera (see Figures 4, 10, 11, 20, 23, and 24). Flippers are largely compressed in two dimensions. Summation, silhouette, and geometric distortion are less problematic during diagnostic interpretation of radiographs of this anatomy. High-quality diagnostic films can be produced by positioning the cassette along the inferior or superior aspect of the flipper with careful attention to keeping the incident beam perpendicular to the cassette. Oblique beam angles are often necessary, as seen in Figures 8A and B. The entire spine of a dolphin can be imaged with the animal suspended in sternal recumbency. A vertical beam with the cassette positioned ventral to the area of interest is used for D-V projections, and a horizontal beam with the cassette positioned laterally for lateral projection images. The beam must be adequately collimated to avoid scatter artifact from adjacent structure mass (Figure 24).
FIGURE 23 D-V radiograph of caudal abdomen of 180-kg male bottlenose dolphin (Tursiops truncatus). Note lack of contrast and anatomical detail. Also note transitional vertebrae and radiopaque foreign body. (Image credit: U.S. Navy.)
0839_frame_C25 Page 579 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
579
FIGURE 24 Lateral radiograph of caudal peduncle mature male bottlenose dolphin (Tursiops truncatus). Note pelvic bones. (Image credit: U.S. Navy.)
Radiographic Pathology
In Figure 25, note the multiple foci of increased interstitial and alveolar pattern in deep lung parenchyma and air-filled lung at the superficial aspect. Ultrasound-guided fineneedle aspiration of lesions confirmed the presence of Cryptococcus neoformans-like fungal elements. Figure 26 demonstrates the appearance of severe pneumonitis. Note the presence of air bronchograms in the caudal ventral aspect, the hallmark of an alveolar pattern. Consolidation of the ventral lung was also obvious on sonography, and was confirmed at necropsy (Figure 27). Figure 10 is a survey lateral radiograph of a mature 170-kg dolphin abdomen. The presence of several transitional vertebrae and the radiopaque foreign body (consistent with a bullet) dorsal and lateral to the spine were incidental findings and not associated with clinical signs. Note the poor contrast and lack of detail of soft tissues. Figures 28 and 29 demonstrate the pre- and postfixation radiographic appearance of multiple open, comminuted fractures of both the mandible and maxilla of a 4.5-year-old male Atlantic bottlenose dolphin. Figure 29 shows placement of external fixation using IMEX
0839_frame_C25 Page 580 Tuesday, May 22, 2001 11:53 AM
580
CRC Handbook of Marine Mammal Medicine
FIGURE 25 D-V radiograph of left hemithorax of 170-kg male bottlenose dolphin (Tursiops truncatus). Note the multiple foci of increased interstitial and alveolar pattern in deep lung parenchyma and air-filled lung at the superficial aspect. (Image credit: U.S. Navy.)
0839_frame_C25 Page 581 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
581
FIGURE 26 Lateral radiograph of rostral aspect of thorax 170-kg male bottlenose dolphin (Tursiops truncatus). Note air bronchograms in ventral lung field. Severe bronchopneumonia confirmed at necropsy. (Image credit: U.S. Navy.)
(IMEX Veterinary, Inc., Longview, TX) threaded pins. Note the presence of clinical union but not bony union after fixator removal 9 weeks postoperatively in Figure 30. In Figure 12, note that the pathology, soft-tissue mineralization, is not visualized on the D-V view due to summation artifact. This demonstrates the requirement for two views in perpendicular planes whenever possible.
Pinniped Normal Radiographic Anatomy
As discussed previously, pinniped patients are not often trained to position themselves and remain motionless long enough to produce diagnostic radiographs. The highest-quality images are obtained with the patient under moderate sedation or light anesthesia. Most radiographs of pinnipeds are thus produced with methods similar to those for other veterinary species under anesthesia. The use of a bucky table expedites cassette positioning beneath the patient,
0839_frame_C25 Page 582 Tuesday, May 22, 2001 11:53 AM
582
CRC Handbook of Marine Mammal Medicine
FIGURE 27 Transverse sonogram of caudal sector of the left thorax of the same 170-kg male bottlenose dolphin (Tursiops truncatus) depicted in Figure 26. Image generated by 3.5-MHz curvilinear transducer. Note lack of reverberation artifact. Cursors indicate path of transcutaneous fine-needle aspiration. (Image credit: U.S. Navy.)
FIGURE 28 Lateral radiograph of rostrum of a juvenile bottlenose dolphin (Tursiops truncatus). Note open comminuted maxillary and mandibular fractures. At initial oral examination, multiple bone fragments were present at the fracture sites. (Image credit: F. Townsend.)
0839_frame_C25 Page 583 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
FIGURE 29 D-V radiograph postfixation of fractures depicted in Figure 28. (Image credit: F. Townsend.)
A
FIGURE 30 Lateral (A) and D-V (B) radiographs of animal depicted in Figures 28 and 29, 9 weeks after removal of fixation. (Image credits: F. Townsend.)
B
583
0839_frame_C25 Page 584 Tuesday, May 22, 2001 11:53 AM
584
CRC Handbook of Marine Mammal Medicine
FIGURE 31 Lateral radiograph of the thorax and cranial abdomen of a juvenile harbor seal (Phoca vitulina). (Image credit: U.S. Navy.)
FIGURE 32 D-V radiograph of the thorax and cranial abdomen of a juvenile harbor seal (Phoca vitulina) depicted in Figure 31. (Image credit: U.S. Navy.)
0839_frame_C25 Page 585 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
585
which is then positioned parallel to the film plane with the aspect of interest as close to the cassette as possible. The reader should recall the advantages of columnation and grid use to reduce mass scatter artifact. A reference set of normal radiographs produced with a California sea lion (Zalophus californianus) under anesthesia was presented in the first edition of this book. The author included excellent line drawings to assist in interpretation and the reader is referred to this work for examples of routine positioning and radiographic anatomy of the California sea lion (Sweeney, 1990). The reader is cautioned to note that the orientations of several of the images in that edition were incorrectly labeled. Figures 31 and 32 are normal survey lateral and D-V radiographs of a juvenile harbor seal (Phoca vitulina). Note the relatively large diameter of the aortic bulge in clinically normal seals. The following section of images is included to illustrate pathology identified on radiographs of sea lions. These cases were chosen for inclusion, because the pathology results in increased radiographic contrast and normal anatomy is well demonstrated. Radiographic Pathology
Figures 33 and 34 illustrate the radiographic changes due to subcutaneous and retroperitoneal emphysema in a juvenile female California sea lion. It was postulated that the animal’s recent restraint in a squeeze cage had resulted in ruptured pulmonary parenchyma or a ruptured airway, causing air leakage into the mediastinal space and migration of air subcutaneously and retroperitoneally. Note the radiolucent appearance surrounding both kidneys due to air in the retroperitoneal space.
FIGURE 33 Lateral radiograph thorax of juvenile female California sea lion (Zalophus californianus). Note enhanced contrast of mediastinal anatomy due to presence of free gas. (Image credit: F. Townsend.)
0839_frame_C25 Page 586 Tuesday, May 22, 2001 11:53 AM
586
CRC Handbook of Marine Mammal Medicine
FIGURE 34 D-V radiograph of the abdomen of a juvenile female California sea lion (Zalophus californianus). Note prominent cranial borders of both kidneys, due to gas in the retroperitoneal space. (Image credit: F. Townsend.)
Radiographs depicting a case of severe mechanical ileus secondary to fecal impaction in a 100-kg castrated male California sea lion are shown in Figure 35. The stomach has been displaced dorsocranially as a result of gas-filled bowel loops.
Computed Tomographic Anatomy As discussed previously, most CT images of marine mammals have been produced as portions of an experimental design or opportunistically on cadaver specimens. Figure 36A and B illustrates the powerful technique of three-dimensional reconstruction that is possible with contemporary CT scanners and image management software. These images are especially useful in planning imaging studies with other techniques or for detailed anatomical investigations. The acquisition of a dolphin whole-body CT scan using a high-speed spiral scanner is currently under investigation by the U.S. Navy marine mammal program.
0839_frame_C25 Page 587 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
587
FIGURE 35 Lateral radiograph of the cranial abdomen of 100-kg male California sea lion (Zalophus californianus). Note distended gas-filled bowel loops. (Image credit: U.S. Navy.)
Magnetic Resonance Imaging Anatomy, Dolphin A whole-body MRI data set was collected 5 hours post-mortem from an 8-year-old, 237cm, 150-kg female Atlantic bottlenose dolphin. Images were produced using a Genesis Signa® diagnostic unit (General Electric Medical Systems, Milwaukee, WI) with T2 weighted technique. The animal was positioned on the magnet gantry in left lateral recumbency. Two perpendicular scan planes were used. Three dorsal plane images and three transverse plane images demonstrating clinically relevant anatomy have been selected, digitally reproduced, and are included here. Figure 37 images A through C were captured in a dorsal plane. Image A visualizes the intrathoracic anatomy at the level of the heart base and also includes the cranial sector of the abdomen. The spatial relationship of the pyloric chamber, pancreas, spleen, forestomach, fundic chamber, and
0839_frame_C25 Page 588 Tuesday, May 22, 2001 11:53 AM
588
A
CRC Handbook of Marine Mammal Medicine
B
FIGURE 36 (A) Three-dimensional CT reconstruction of skull of 200-kg male bottlenose dolphin (Tursiops truncatus). Skull, mandibles, and hyoid as seen from posterior aspect. Note dense ear bones. (Image credit: U.S. Navy.) (B) Three-dimensional CT reconstruction of juvenile northern elephant seal (Mirounga angustirostris) head as viewed from rostral aspect. (Image credit: M. Haulena.)
small intestine is clearly imaged and of particular clinical value. Image B clearly shows the intrathoracic anatomy at the level of the primary bronchi and the cranial sector of the abdomen at the level of the fundic chamber and forestomach. Dependent fluid accumulation is seen in the left aspect as increased signal strength. The animal was positioned in left lateral recumbency during the immediate post-mortem period and scan acquisition. Recently ingested fish is visible within the forestomach lumen. Image C demonstrates the cranial and middle sectors of the abdomen. Note the ventral flexure of the pyloric chamber, the pancreas, the spleen, and the position of both kidneys. Figure 37 images D through F were captured in a transverse plane. Image D shows the intrathoracic anatomy at the level of the heart. Dependent fluid accumulation is again evident as increased signal strength. Also note the compression of the dependent left lung and the position of the margin of both lungs with respect to the heart. Recently ingested fish is seen within the esophageal lumen. Image E demonstrates the overlap of the caudal sector of the thorax with the cranial sector of the abdomen. This is of particular clinical importance when utilizing other imaging techniques. Reverberation artifact during sonography reduces the acoustic window to the cranial sector of the abdomen and summation sign in radiographs will often obscure caudal pulmonary anatomy. Image F illustrates the complex gastrointestinal anatomy of the cranial sector of the abdomen. The ampulla of the duodenum is visible dorsal to the ventral flexure of the pyloric chamber, and can be distinguished from the latter by the presence of luminal folds. This is also clearly evident on sonography. A dependent fluid line is seen within the ampulla. Luminal folds are also clearly visible within the esophagus as it enters the forestomach.
Acknowledgments The authors thank the animal care specialists assigned to Navy Element Army Activities who have been instrumental in developing the standard techniques discussed in this chapter, specifically, MSG Bill Applegate, SFC Daniel Jermier, CWO Kevin Knight, SPC Rushawn Orth, SPC Cheryl Short, SPC Ian Mathey, and PVT Lisa Tanner. The authors also thank the entire animal care staff of Science Applications International Corporation.
0839_frame_C25 Page 589 Tuesday, May 22, 2001 11:53 AM
589
CRC Handbook of Marine Mammal Medicine
A
D
B
E
C
F
FIGURE 37 Select MRI images of 150-kg female bottlenose dolphin (Tursiops truncatus). Images A through C were captured in a dorsal plane through the thorax and cranial abdomen. Images D through F were captured in a transverse plane through the thorax and cranial abdomen. Note presence of fish in forestomach lumen and dependent changes in signal due to animal’s left lateral recumbency during scanning. (Image credits: U.S. Navy.)
0839_frame_C25 Page 590 Tuesday, May 22, 2001 11:53 AM
590
CRC Handbook of Marine Mammal Medicine
References Artz, D.S., 1997, Computed radiography for the radiological technologist, Semin. Roentgenol., 32: 12–24. Barber, D.L., and Lewis, R.E., 1982, Guidelines for Radiology Service in Veterinary Medicine, produced by the American Veterinary Medical Association, 24 pp. Britt, J.O., and Howard, E.B., 1983, Anatomic variants of marine mammals, in Pathobiology of Marine Mammal Diseases, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 7–46. Bryden, M.M., and Felts, J.L., 1974, Quantitative anatomical observations on the skeletal and muscular systems of four species of Antarctic seals, J. Anat., 118: 589–600. Endo, H., Sasaki, H., Hayashi., Y., Petrov, E.A., Amano, M., Suzuki, N., and Miyazaki, N., 1999a, CT examination of the head of the Baikal seal (Phoca sibirica), J. Anat., 194: 119–126. Endo, H., Yamagiwa, D., Arishima, K., Yamamoto, M., Sasaki, M., Hayashi, Y., and Kamiya, T., 1999b, MRI examination of trachea and bronchi in the Ganges River dolphin (Platanista gangetica), J. Vet. Med. Sci., 61: 1137–1141. Felts, J.L., and Spurrell, F.A., 1966, Some structural and developmental characteristics of cetacean (Odontocete) radii. A study of adaptive osteogenesis, Am. J. Anat., 118: 103–133. Fessenden, R.J., and Fessenden, J.S. (Eds.), 1979, Organic Chemistry, Willard Grant Press, Boston, MA, 332 pp. Haulena, M., Gulland, F.M.D., DeCock, H.E.V., and Harman, J.W., 1998, The use of computerized tomography for diagnosis of osteomyelitis in pinnipeds, in Proceedings of the 30th International Association for Aquatic Animal Medicine Conference, May 2–6, San Diego, CA, 133–134. Hillmann, D.J., 1991, Anatomy of the fetal bowhead whale (Balaena mysticetus) using magnetic resonance images, Abstr., Proceedings of the 22nd International Association for Aquatic Animal Medicine Conference, 172. Kantor, C., 1997, Computed radiography, Biomed. Instrum. Technol. 31: 73–75. Kooyman, G.L., Hammond, D.D., and Schroeder, J.P., 1970, Bronchograms and tracheograms of seals under pressure, Science, 169: 82–84. Matassa, K., Early, G., Wyman, B., Rommel, S., and Krum, H., 1994, The stranding of a neonate male pilot whale (Globicephala melaena): A case study, Abstr., Proceedings of the 25th International Association for Aquatic Animal Medicine, 166. Morris, E.L., 1990, Radiography of dolphins, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 193–201. Nordoy, E.S., and Blix, A.S., 1985, Energy sources in fasting grey seal pups evaluated with computed tomography, Am. J. Physiol., 249: 471–476. Ogden, J.A., Conlouge, G.J., and Rhodin, A.G.J., 1981, Roentgenographic indicators of skeletal maturity in marine mammals (Cetaceae), Skeletal Radiol., 7: 119–123. Ponganis, P.J., Kooyman, G.L., Sartoris, D., and Jobobis, P., 1992, Pinniped splenic volumes, Am. J. Physiol., 262: 322-325. Simpson, J.G., and Gardner, M.B., 1972, Comparative microscopic anatomy of selected marine mammals, in Mammals of the Sea: Biology and Medicine, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 298–418. Sommer, L.S., McFarland, W.L., Galliano, R.E., Nagel, E.L., and Morgane, J.P., 1968, Hemodynamic and coronary angiographic studies in the bottlenose dolphin (Tursiops truncatus), Am. J. Physiol., 215: 1498–1505. Sumner-Smith, G. Pennock, P.W., and Ronald, K., 1972, The harp seal (Pagophilus groenlandicus, Erxleben, 1777): XVI epiphyseal fusion, J. Wildl. Dis., 8: 29–32. Sweeney, J.C., 1990, California sea lion radiology, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 203–213. Sweeney, J.C., and Ridgway, S.H., 1975, Procedures for the clinical management of small cetaceans, J. Am.Vet. Med. Assoc., 167: 540–545.
0839_frame_C25 Page 591 Tuesday, May 22, 2001 11:53 AM
Radiology, Computed Tomography, and Magnetic Resonance Imaging
591
Thrall, D.E., 1986, Introduction to radiographic interpretation, in Textbook of Veterinary Diagnostic Radiology, Thrall, D.E. (Ed.), W.B. Saunders, Philadelphia, 1–11. Viamonte, M., Morgane, P.J., Galliano, R.E., Nagel, E.L., and McFarland, W.L., 1968, Angiography in the living dolphin and observations on blood supply to the brain, Am. J. Physiol., 214: 1225–1249. Woodhouse, C.D., and Rennie, C.J., 1991, Observations of vaginal calculi in dolphins, J. Wildl. Dis., 27: 421–427.
0839_frame_C25 Page 592 Tuesday, May 22, 2001 11:53 AM
0839_frame_C26 Page 593 Tuesday, May 22, 2001 11:54 AM
26 Ultrasonography Fiona Brook, William Van Bonn, and Eric Jensen
Introduction The advantages and clinical applications of diagnostic ultrasound in the field of veterinary medicine are now well established, and this technique plays a valuable role in the care of marine mammals. However, few clinicians appear to have utilized the diagnostic capabilities of B-mode sonography, and even fewer have reported their experiences, so that there is little available reference material. Every potential practitioner should study the basic physics of ultrasound, since this is essential to understanding how ultrasound interacts with different tissues and interfaces, how the image is formed, and how to recognize image artifacts; there are several good, simple reference texts available (Hykes et al., 1992; Sanders, 1998). The purpose of this chapter is to provide hands-on sonographic information and practical marine mammal guidelines. Equipment selection, scanning techniques, common errors and limitations, and normal sonographic anatomy are described, along with illustrations of lesions identified in marine mammals using ultrasound. Much of the sonographic information gathered to date has been in dolphins and the information in this chapter is no exception; more work on other marine mammals is needed.
Indications Sonography is a valuable adjunct to information provided from physical examination, radiology, and endoscopy. It is indicated any time that more information about internal structure is desirable. This technique has proved more valuable than plain radiographs in marine mammal species. The inclusion of sonography in routine health surveys should be considered in a comprehensive preventive medicine program. As a diagnostic technique, sonography is simple, safe (AIUM, 1988), and cost-effective, and its noninvasiveness and ease of adaptability to any working environment make it an ideal addition to the diagnostic armamentarium of the marine mammal clinician. Ultrasound can provide a vast amount of morphological information, because it is able to differentiate soft tissue structures, accurately assess morphology and morphometry, and locate and characterize many conditions that alter echopatterns, shape, and position. Animals can be trained to cooperate for examination, chemical restraint is not required, and examinations can be repeated as often as necessary without risk, enabling monitoring of disease and the course of therapy. Pulmonary disease is prevalent in both wild and captive marine mammals worldwide (Brown et al., 1960; Duignan et al., 1992; Reeves et al., 1994), and is considered by many to be the
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
593
0839_frame_C26 Page 594 Tuesday, May 22, 2001 11:54 AM
594
CRC Handbook of Marine Mammal Medicine
most common cause of death in dolphins (see Chapter 40, Cetaceans). The high incidence of such disease is probably due to the specific adaptations in the anatomical structure of the cetacean respiratory apparatus (Wislocki, 1929), plus the lack of a cough reflex, preventing emission of exudates and particulates. If available, sonography should be part of the first line of investigation of suspected pulmonary disorders or thoracic trauma in marine mammals. Chest sonography is also indicated following trauma; pleural effusion, chylothorax, hemothorax, and/or rib fractures are all diagnosable using sonography.
Limitations Ultrasound examinations of marine mammals are often performed outdoors, in direct sunlight, and it may be difficult to see the image adequately. This can be overcome by using (1) a darkcolored blanket, somewhat like an old-fashioned photographer’s hood, to shield the monitor or (2) a heads-up video display, which allows remote viewing of the screen. A sick animal may not be cooperative and this may necessitate removal of the dolphin from the water in order to conduct an adequate examination. When this is necessary, or if animals are accustomed to such handling, an appropriately prepared deck area should be available at, or near, the side of the enclosure and the dolphin placed on a wet foam mattress. In the authors’ experience, this has had no serious adverse effect on any dolphin. However, it should be noted that sick animals, especially those with respiratory disease and significant lung disease, may ˙ ⁄Q ˙ ) inequalities out of water, and thus caution experience significant ventilation/perfusion ( V should be taken in working with dolphins so affected. When necessary, these effects can be minimized by scanning the animal in sternal recumbency. The sheer size of some marine mammals may prohibit useful ultrasonographic examination of some structures. The relatively thick integument and blubber layers in animals such as the walrus are very difficult to penetrate without serious loss of resolution and image detail. Having said that, it is always worth a try!
Technique Equipment and Preparation Any standard diagnostic ultrasound unit with a “scroll” or “zoom” capability, allowing visualization of deeper structures, can be used to examine marine mammals. Transducer selection depends on the size of the patient and the area of interest. A 3.5-MHz linear/curvilinear array transducer provides a wider field size, suitable for examination of larger animals and deeper structures. A 3.5-MHz linear array, with a 10-cm footprint is best for larger animals, or to visualize large or deep organs and advancing pregnancies. A 5-MHz curvilinear array is suitable for smaller animals and superficial structures. For eye examinations, a 10-MHz transducer is ideal, although a reasonable examination can be conducted using 7.5 MHz. It may be necessary to use up to 2.5 MHz for examination of deep structures in very large animals, such as big killer whales (Orcinus orca), but it must be noted that image detail and, therefore, diagnostic accuracy will be sacrificed. Sector probes are not as suitable, as the field of view close to the transducer is limited, the lateral resolution is poorer, and edge artifacts are more pronounced. The noise and vibrations of mechanical sector transducers also may disturb some animals. Modern curved array probes provide adequate access, even when acoustic windows are small. Gone are the days of having to worry about fixed focal zones and changing transducers in the middle of an examination. All modern transducers have dynamic focusing, which means the depth of the focal zone is constantly changed along the beam axis, effectively extending the
0839_frame_C26 Page 595 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
595
focal zone along the entire depth of the image. Many machines also have manual focusing options, which allow for fine tuning of layers of the image at particular depths. An effort should be made to ensure consistent use of power, or “gain,” levels, both for comparison between animals and within individuals. It can be all too easy to mislead oneself into thinking a structure is “brighter” or “darker” than it was before, if care and attention are not paid to this. This must also, obviously, be considered in light of the animal’s weight—significant weight gain or loss will affect the acoustic power of the ultrasound beam, and thereby the brightness of the image. All modern transducers are waterproofed, but may be further protected by covering them with a sheath or plastic bag containing some acoustic gel, and sealed with waterproof tape around the cable. If animals are trained for enclosure-side examination, the area being examined is often at or above the surface and no covering is needed. Coupling gel may be needed when examining pinnipeds. However, because of the nature of the dolphin’s skin, there is no surface air layer to act as a barrier to the ultrasound beam, and gel is not required. Animal training for medical procedures, including ultrasound scanning, is now an integral part of husbandry and management of marine mammals in many facilities. With training, these animals quickly accommodate and cooperate in examinations. Pinnipeds can be trained to station erect, or to lie down and roll into a supine or semisupine position for examination. Both positions are necessary—the erect for examination of the chest, upper abdomen, head, and neck, and the supine for examination of the lower abdomen and pregnancy. They may also be scanned while lying in a tub of water, which can eliminate the need for coupling gel.
Image Orientation Consistent orientation and labeling of images is essential for systematic interpretation of scans and for image transfer between clinicians. Consistent positioning of animals aids this and also reinforces the animal’s training. There are two main methods of orienting ultrasound images. The first is most commonly used by sonographers and some veterinarians. The top of the image represents the skin surface on which the transducer is placed. For sagittal, parasagittal, and/or coronal (also termed dorsal) scans, the animal will be in sternal, dorsal, or lateral recumbency, with the head to the left, or in front, of the operator, or erect. The cranial aspect of the image is to the left of the monitor, and the caudal aspect is to the right. For transverse views, when the animal is in dorsal recumbency, the animal’s head should be to the left of the operator. The left side of the abdomen should be on the right of the image/monitor and right side on the left (i.e., the image is oriented to the viewer in the same way as the animal’s anatomy while scanning). For transverse views through the side of the abdomen (e.g., when scanning the kidney, ovary, and testis), the animal is in lateral recumbency, with the head to the operator’s left, or erect, for the left side and the head to the operator’s right for the right side. The side being examined is at the top of the image/monitor, the dorsum is on the right, and the ventrum on the left. The second method is the one that has been most commonly used by veterinarians, particularly for small-animal studies, and standardizes transducer placement to produce sonograms with the following orientations: for scans parallel to the long axis of the body (sagittal, parasagittal, coronal /dorsal, oblique), the head of the animal is at the right of the sonogram; for scans perpendicular to the long axis of the animal (transverse, oblique), the dorsal aspect of the animal is to the right on the sonogram. The authors believe the method used is a matter of personal preference, and that operators should choose the one they are most comfortable or familiar with—but they emphasize the need for consistency and accurate labeling. Standardization of image labeling and orientation reduces confusion when interpreting sonograms, allows for consistency between studies, and facilitates ease of transducer placement with animals routinely positioned in or out of the water.
0839_frame_C26 Page 596 Tuesday, May 22, 2001 11:54 AM
596
CRC Handbook of Marine Mammal Medicine
Clinical Applications Thoracic Imaging Sonography of the chest can be difficult because of the air-filled lungs, but is a valuable method of detection and characterization of changes in the pleura, pleural cavity, and pulmonary parenchyma. Pleural effusion, pneumonia, lung abscessation, and thoracic lymphadenopathy are all detectable on sonography. Ultrasound-guided thoracentesis can facilitate collection of aspirates for culture or cytology, providing valuable information for specific therapy, while also providing relief of physical signs in the animal with large pleural effusions, respiratory distress, tachycardia, and altered buoyancy (Rhinehart et al., 1995). Little is known of cardiac disorders in marine mammals and, to the authors’ knowledge, there has been only one report of congenital, structural anomalies in a dolphin fetus (Gray and Conklin, 1974). Congestive heart failure and amyloidosis have been reported in dolphins and seals (Miller and Ridgway, 1963; Reeves et al., 1994; Cowan, 1995) (see Chapter 23, Noninfectious Diseases). Parasitosis is another concern, and cardiac nematode infestation has been reported (Brown et al., 1960) and depicted using ultrasound in harbor seals (Phoca vitulina) (Stone, 1990).
Heart and Mediastinum Echocardiography is commonly used in veterinary care of many species (Nyland and Mattoon, 1995), but is not as easily applied in marine mammals, particularly cetaceans. Although an experienced echographer could probably detect gross abnormalities using standard techniques, further data collection is necessary to document normal cardiac parameters for meaningful echocardiography in all marine mammal species. B-mode echocardiography provides a two-dimensional (2D) image, which allows comprehensive evaluation of the anatomy, spatial relationships, and motion of cardiac structures. M-mode echocardiography is an older technique that uses a much narrower ultrasound beam to detect only axial motion in a very small section of the heart. A B-mode transducer is used to produce a 2D image, which is used to guide and orient the M-mode examination. Axial measurements of the dynamics of different structures can be made. The advantage of M-mode is that it has a much faster sampling rate than B-mode, which is limited by lower frame rates and poorer image resolution. M-mode recordings are much closer to “real time” and so are better able to detect rapid movement and subtle changes, which may be missed on B-mode alone. Meaningful M-mode examinations require standardization of techniques, as well as knowledge and understanding of beam paths and normal and pathological cardiac anatomy. M-mode echocardiography is currently being used by one of the authors (E.J.) to monitor and establish norms for fetal heart rates in hopes of using this as a tool to monitor overall fetal health and fetal stress. Often the entire cardiac surface area in the dolphin is surrounded by lung tissue. The large sternum obstructs access from the ventral midline and the heart is therefore not accessible. In some animals, the lungs are not so intrusive and a low parasternal approach may allow access to the heart. This is best done with a dolphin remaining in the water in dorsal recumbency. This position may help to remove the lungs from more of the cardiac surface and increase available window size. Trans-sternal echocardiography is possible in the very young calf. Transesophageal echocardiography is possible in larger animals, with the endoscopic transducer inserted, through a mouth guard, in the same way as a standard endoscope or gastric tube. Doppler echocardiography uses changes in frequency of a sound beam, which is reflected from moving cellular elements in blood, to depict and assess various blood flow parameters. There are several Doppler techniques, including color flow imaging, in common clinical use
0839_frame_C26 Page 597 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
597
FIGURE 1 Left coronal/dorsal view of the aortic arch in a dolphin (head to the left; left side up).
for human cardiac evaluation. Doppler echocardiography provides further information on hemodynamics, including direction and characterization of flow patterns. Again, applications in marine mammals have not been explored. Similar to the heart, the mediastinum is not easily visualized in dolphins; nevertheless, visualization should be attempted if clinically indicated. An enlarged thymus or mediastinal lymphadenopathy may be seen, and benign mediastinal tumors, lymphangiomyomatosis, cystic transformation, and malignant lymphomas have been reported in cetaceans (Cockrill, 1960; Rawson et al., 1992; Cowan, 1994; Bossart et al., 1997). The aortic arch and associated vessels of the smaller dolphin species can sometimes be demonstrated by placing the transducer immediately above the sternum, in the transverse plane, and angling the beam approximately 30 to 45° caudally, or by scanning in the coronal/dorsal plane, just above the pectoral flipper (Figure 1). This view is also useful for evaluating possible aortic arch nodes, which are associated with the thyroid and thymus. Pinnipeds are easier subjects for echocardiography. Whether trained or restrained, surrounding lung, as in dolphins, does not obstruct access to the heart. Intercostal spaces are large and an oblique parasternal approach from either side allows good visualization of the entire heart. Again, gross abnormalities may be detected, but normal parameters are, as yet, unknown. More work needs to be done in this area.
Lungs The anatomy and surface landmarks of the lungs are described in Chapter 9. A full survey of both lungs should be carried out, with particular attention paid to the ventrolateral bases, as these are most dependent and commonly involved in cases of pulmonary infection. With the dolphin in lateral recumbency, the transducer is placed in the transverse plane and perpendicular to the subject, below the pectoral flipper. It is then moved caudally toward the base of the lung, along the pectoral line. Moving in parallel sweeps, the examination continues until the entire lung has been assessed in a grid-type pattern. The procedure is repeated with the transducer in a sagittal/parasagittal plane, aiming toward the midline, and then repeated again for the contralateral lung. The surface of the parietal pleura can be examined through the intercostal spaces and lies just deep to a thin, echolucent muscle layer beneath the ribs. Many dolphins show evidence of apparent pleural irregularity with multiple associated, highly echogenic “streaks” on the image. These are “comet-tail” artifacts, a type of reverberation artifact produced at an air interface and a phenomenon
0839_frame_C26 Page 598 Tuesday, May 22, 2001 11:54 AM
598
CRC Handbook of Marine Mammal Medicine
FIGURE 2 Sagittal scan showing longitudinal section through mid-abdomen of a young T. t. aduncas. Highly echogenic “comet-tail” artifacts are seen at air/bowel wall interfaces.
FIGURE 3 Sagittal, midline scan of caudal thorax showing suspected hemothorax following trauma in a false killer whale (Pseudorca crassidens).
FIGURE 4 Left coronal/dorsal view of the left lung base in a bottlenose dolphin (T. truncatus), demonstrating extensive consolidation. Lack of aeration in the affected area allows penetration of the ultrasound beam. Cursor marks show guide for needle placement and required depth for UGFNAB. (Image credit: U.S. Navy.)
0839_frame_C26 Page 599 Wednesday, May 23, 2001 12:26 PM
Ultrasonography
599
well recognized in sonography (Laing, 1983; Thickman et al., 1983). This appearance has been observed in animals with no subsequent evidence of pleural abnormality at necropsy and is not indicative of pathology, but may be age associated. Other interfaces that commonly produce this type of artifact are the diaphragm-lung and bowel wall-bowel gas (Figure 2). Normal, air-filled lung presents the typical reverberation artifact of an air-to-tissue/fluid interface. Pulmonary infiltrate may be diagnosed by absence of the normal reverberation pattern and penetration of the ultrasound beam to reveal the presence of fluid within the thorax (Figure 3), pulmonary lesions (most commonly abscesses), or consolidation (Figure 4). The appearance of echolucent, edematous, or consolidated lung tissue surrounding air-filled bronchioles is pathognomic of pneumonia. Smaller, more discrete, echolucent or heterogenous lesions (Figure 5) are more likely to be sites of focal abscessation, which are commonly reported in both pinnipeds and cetaceans. Care must always be taken to image any suspected abnormality in at least two planes, to ensure a focal lesion is indeed present. The sonographic appearances of abscesses are variable and these may be difficult to identify, particularly if they are isoechoic to surrounding tissue, or echogenic because of the presence of gas or chronic changes, such as encapsulation or mineralization. Borders may be irregular and poorly demarcated, and the appearance of contents may be misleading. Where there is
FIGURE 5 Right coronal/dorsal views of the pleural surface showing (top) small granulomata in a killer whale calf and (bottom) subpleural abscess in a bottlenose dolphin (T. truncatus). (Image credits: T. Robeck.)
0839_frame_C26 Page 600 Tuesday, May 22, 2001 11:54 AM
600
CRC Handbook of Marine Mammal Medicine
doubt, chest radiography should also be performed for further information. Again, it is important to scan and repeat-scan systematically, and to ensure the position of any lesion is accurately determined and recorded for follow-up examination to be most useful. In Figure 3, there are multiple rib fractures and a probable hemothorax in a young, male false killer whale (Pseudorca crassidens) attacked by an adult female killer whale. At least one fracture was confirmed radiographically, but it was not possible to obtain adequate radiographs of all sites in this large animal. Removal of a large animal with suspected rib fractures from the weight-bearing environment of the water to a hard poolside surface may, indeed, be contraindicated, because of the risk of lung puncture by a displaced fracture or fracture fragments. Routine sonography of this animal, under trained behavior, allowed monitoring of the fluid collection, callous formation, and the degree of fracture healing, and assisted in determination of when to allow the animal to return to performance training. Ultrasound-guided fine-needle aspiration biopsy (UGFNAB) of pulmonary lesions can provide valuable information on cause and for determining appropriate therapy. UGFNAB has been performed successfully in dolphins (see Figure 4), including repeat procedures on the same animal at intervals of several weeks, without complications (Van Bonn, pers. obs.). The region of interest is visualized, and the skin aseptically prepared as for surgery. Sterile lubricating gel is used if coupling is necessary. Needle placement is guided by the sonogram to avoid vital structures and vasculature. A regional “wash” is also rewarding. Introduce 10 to 15 ml of sterile saline via the needle and apply suction immediately to recover as much diagnostic material as possible. Cytology, microbiology, and virology of the aspirated material is extremely valuable in the diagnosis of several respiratory diseases (see Chapter 20, Cytology). Always remember that lesions deep to normal lung and not extending to the pleura will not be able to be visualized. Sonography, therefore, may identify the presence of pulmonary disease, but cannot exclude it, and may not demonstrate the full extent of the affected area. Chest radiography should always be performed, if clinically indicated and feasible.
Thoracic Lymph Nodes A dolphin has numerous lymph nodes within the thorax, which often enlarge markedly in the presence of pulmonary infection, presenting an amorphous, echolucent “mass” on sonography (Figure 6, left), which may be difficult to differentiate from an area of consolidation.
FIGURE 6 Parasagittal views of diaphragmatic lymph nodes in a bottlenose dolphin (T. t. aduncas), showing (left) acute lymphadenopathy and (right) chronic appearance with mineralized foci.
0839_frame_C26 Page 601 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
601
These may also contain mineralized foci, particularly if chronically affected (Figure 6, right). Thoracic lymphadenopathy is most commonly seen in the caudoventral aspect of the thorax, just above the diaphragm and ventral to the heart (diaphragmatic nodes), and in the apical region, where the prescapular (or superficial cervical) nodes are situated. Chest radiography in these large animals may be difficult, and radiographs are often of poor quality, although the newer technology of computed radiography (CR) holds much promise (see Chapter 25, Radiography). Pulmonary changes may also be subtle and not clearly demonstrated on radiographs. Therefore, the visualization of affected lymph nodes is very important and may be the only detectable sign of chest infection on imaging. UGFNAB of prescapular nodes has been performed without complications by one of the authors (W.V.B.). Although not as yet reported in the literature, UGFNAB of intrathoracic lymph nodes may also be a valuable diagnostic procedure in cases of suspected thoracic disease with demonstrated lymphadenopathy.
Abdominal Imaging In the dolphin, scanning of the upper abdomen is relatively easy. However, the more acute rounding of the caudoventral abdomen and the dense musculature make a midline approach difficult, particularly in the lower abdomen. Scanning in the parasagittal and transverse planes, through the lateral abdominal wall, provides the best contact and access to the entire abdomen. For standardization, the abdomen can be divided into three sections: (1) cranial, containing the liver, biliary system, spleen, pancreas, forestomach, second (or fundic) stomach, duodenal ampulla, and the pylorus; (2) middle, containing the kidneys, small bowel, and gravid uterus; and (3) caudal, containing small bowel, rectum, urinary bladder, and reproductive organs. Animals vary in body size and shape; similarly, the size and “normal” echopatterns of each individual’s organs vary. Thus, a baseline record of each individual in the normal state should be obtained. Abdominal scanning in the smaller seals, particularly in trained animals, is not difficult, although a thorough examination can take time, since the animal will need to be repositioned several times to allow access to the whole abdomen. Ultrasound may not always be possible in larger animals, such as big males, or walruses, due to their size and thick blubber layers. The liver, biliary system, spleen, and urinary and reproductive tracts can be separately identified and assessed for normal size and echopatterns. The gastrointestinal (GI) tract can also be visualized, and the motility of the bowel and appearance of bowel contents can prove to be clinically significant. Unless grossly affected, the normal pancreas is very difficult to visualize adequately.
Liver and Biliary System Liver disorders are relatively common in wild and captive marine mammals, particularly parasitosis, hepatic lipidosis, hepatitis, and diffuse and focal cirrhosis (Cockrill, 1960; Brown et al., 1960; Baker, 1992). The large dolphin liver is molded to the inferior surface of the diaphragm, extends across the entire upper abdomen, and is larger on the right side. The liver is examined by placing the dolphin left side up initially, with the transducer perpendicular and placed longitudinally and parallel to the pectoral flipper, along the pectoral plane. Scan from the diaphragm to the caudal hepatic margin. Sequential sagittal/parasagittal and transverse scans of the whole liver are then carried out. The procedure is repeated with the dolphin right side up. Baseline records should include the distance of the caudal extent of the liver above (U.L. + x cm) or below (U.L. − x cm) the umbilicus, in the midline. This is done with the animal in the supine position. The caudal margin of the liver usually lies rostral the umbilicus in a dolphin; however, in a large animal, the normal liver may extend several centimeters caudal
0839_frame_C26 Page 602 Tuesday, May 22, 2001 11:54 AM
602
CRC Handbook of Marine Mammal Medicine
FIGURE 7 (Left) Transverse oblique scan showing a normal gallbladder and right lobe of liver in California sea lion (Z. californianus) and (right) right coronal/dorsal section of normal liver in a bottlenose dolphin (T. truncatus).
the umbilicus. Measurements will, of course, vary with degree of inspiration, or flexion/extension of the body. Care should be taken to ensure the dolphin inhales in the same way before rotating and is as relaxed as possible, to maximize repeatability of measurements. Although not strictly accurate, this technique does provide a useful indicator of significant increase or decrease in liver size, and is a useful part of routine examination and records. In pinnipeds, the entire liver can be examined in the supine position, although it will probably be necessary to reposition the animal to assess the whole liver adequately. The upper liver can be examined with the animal erect, but the caudal margin extends very low in the abdomen and is not adequately visualized in this position. The gallbladder is relatively large and easy to evaluate. The outline of the liver is smooth with a sharp caudal edge. The normal echotexture is similar to that seen in most liver tissue, presenting a homogenous, finely speckled pattern of midlevel intensity (Figure 7), frequently interrupted by hepatic and portal vascular branches. The large hepatic veins in dolphins and venous sinuses in pinnipeds are easily visualized, coursing toward the vena cava. Thicker, echogenic borders identify the portal veins, due to surrounding Glisson’s capsule. Nowadays, with higher-resolution equipment, intravascular flow within the normal hepatic and portal veins is frequently observed, and must not be interpreted as pathological. The flow is imaged as moving echoes within the lumen of the vessel, and is in phase with the cardiac cycle. Flow may be seen to be bidirectional or even turbulent, especially at the low end of the cardiac cycle. The signal is similar to that seen due to rouleaux formation in humans and is thought to be a result of low flow velocities in the central veins of diving mammals. This phenomenon is currently being investigated by one of the authors (W.V.B.). Ideally, if the biliary system is to be evaluated, subjects should not be fed for about 6 hours prior to examination, to allow maximal accumulation of bile and full distension of the biliary tree. The pinniped gallbladder is large and easily seen when distended. Dolphins do not have a gallbladder, but the distal bile ducts are relatively large and similar in size, position, and echopattern to the portal veins, so care must be taken to differentiate these. Bile ducts tend to run deep to the portal veins and are larger in diameter at the porta hepatis, but taper quickly within the liver (Figure 8) and are not normally seen within the peripheral parenchyma. Disease of the biliary system is apparently rare in marine mammals. A “starry-sky” pattern of the liver, with transitory dilatation of the central bile ducts, has been seen in a proven case of fluke infestation (Figure 9). On one occasion, mobile, echogenic foci were seen within the lumen of the ducts of this animal. These disappeared following treatment and the ducts regressed to normal size.
0839_frame_C26 Page 603 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
603
FIGURE 8 Radiograph of an excised bottlenose dolphin ( T. truncatus) liver. The bile duct has been cannulated and injected with contrast medium to delineate the biliary tree.
FIGURE 9 Transverse section of liver showing abnormal sonographic appearance of intrahepatic bile ducts in a case of fluke infestation in a young, male bottlenose dolphin (T. t. aduncas).
Changes in size, shape, and echogenicity of the liver may indicate diffuse lesions. Blunting of the caudal margin and “bulging” of the liver between the ribs has been seen in acute hepatitis. Diffuse increases in parenchymal echogenicity have been observed with subsequent biochemical indicators of hepatopathy (see Chapter 19, Clinical Pathology). If liver disease is suspected, an attempt should be made to examine the area around the porta hepatis and the para-aortic regions, for lymphadenopathy (Figure 10). A general increase in liver echogenicity is common in pregnant dolphins, as increased fat deposition occurs (Figure 11, left). It is advisable to monitor the degree of “fatty liver” in pregnancy, particularly in females with a history of liver dysfunction, since this has been known to preempt hepatic failure due to excessive steatosis in one bottlenose dolphin (Tursiops truncatus aduncas) with a prior history of acute hepatitis (Brook, pers. obs.). Cirrhotic changes, with lobular fibrosis and decreased size, may be observed in dolphins with a history of hepatitis (Figure 11, right). If this is suspected, the animal should be monitored for disease progression, even if liver function appears clinically normal. Discrete hepatic lesions are relatively rare in
0839_frame_C26 Page 604 Tuesday, May 22, 2001 11:54 AM
604
CRC Handbook of Marine Mammal Medicine
FIGURE 10 Longitudinal section of normal portal vein and periportal lymph nodes in a bottlenose dolphin (T. truncatus).
FIGURE 11 (Left) Fatty liver in a pregnant bottlenose dolphin (T. t. aduncas). (Right) Cirrhotic changes in the liver of an adult, male bottlenose dolphin (T. t. aduncas) with a history of repeated episodes of hepatopathy. The parenchymal echopattern is hyperechoic and coarsened, and intralobular fibrosis can be seen at the boundary between the liver parenchyma and the large intrahepatic vessels.
marine mammals, although granulomas, fibromas, lipomas, and hemangiomas have been reported (Cockrill, 1960; see Chapter 23, Noninfectious Diseases).
Spleen The spleen in a bottlenose dolphin is relatively small and rounded, or slightly ovoid, lying just to the left of the midline, medial and deep to the forestomach, and often behind the second stomach (fundic chamber) and duodenum (Figure 12). It is ideal to restrict food for 6 hours prior to sonographic examination, to facilitate visualization of the spleen without the food-filled stomachs and duodenum getting in the way. Because of the dolphin spleen’s variable position and the gut that surrounds it, sonographic identification may be difficult. The spleen is most often seen using a parasagittal approach through the ventral abdomen from the right side, using the liver as an acoustic window. As it is small, the normal spleen can be visualized in its entirety and presents a very even, speckled
0839_frame_C26 Page 605 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
605
FIGURE 12 Left coronal/dorsal views of (left) a hyperechoic spleen in a stranded bottlenose dolphin (T. truncatus). (Image courtesy of T. R. Robeck.) (Right) A normal bottlenose dolphin (T. t. aduncas) spleen. (The stranded animal was diagnosed with pneumonia, and it is possible that the spleen was affected, but this was not determined.)
echo pattern, usually isoechoic or slightly more echogenic than the normal liver. Reported normal sizes are variable within the same species (Figure 12)—3.5 to 6.0 cm in T. t. aduncas; 6.0 to 8.0 cm in T. t. truncatus (Stone, 1990)—and normal baselines should be documented for each individual when possible. Because of the mobility of the spleen, it is difficult to access the largest diameter reliably, and this may account for some variability of sizes noted. Splenic size appears to increase only slightly, or remain unchanged, in cases of systemic infection in dolphins. The size appears to be most dramatically affected in cases of septicemia, where splenic congestion usually presents as increased echolucency of the parenchyma. If hemorrhagic, the sonographic appearance of the spleen may vary from diffusely echolucent to diffuse or focal areas of increased echogenicity. In seals, the sonographic appearance of the spleen is similar to that of the dog, although it is more difficult to access, because of the elongated rib cage of the pinniped. It is also obscured by the stomach and the small and large bowel. It may be accessed through scanning in a subcostal oblique plane, allowing the transducer to be guided by the left costal margin, or by transverse or coronal scans from the left side, through the intercostal spaces. The subject should be in an erect position, if possible. A curvilinear array is the best choice of transducer for examination of the spleen in pinnipeds, as it allows a reasonable contact area but with the flexibility needed to angle up and between ribs.
Pancreas It is difficult to visualize the margins of the normal marine mammal pancreas, because of its elongated and irregular shape, lack of definitive border, and its position, closely affixed to the duodenum, often behind the second stomach and the bowel. The gland is finely lobulated, with no capsule to provide differentiation or delineation on sonography. Sections of the normal pancreas may sometimes be identified when surrounded by fluid-filled bowel loops. Pancreatic pathology in dolphins is most commonly confirmed in cases of septicemia (Brown et al., 1960).
Gastrointestinal Tract Gastrointestinal disorders are common in marine mammals, especially dolphins. With relatively less intestinal gas than most species, sonography can provide useful information about much
0839_frame_C26 Page 606 Tuesday, May 22, 2001 11:54 AM
606
CRC Handbook of Marine Mammal Medicine
FIGURE 13 Sonographic appearance of (left) a normal, empty fundic stomach and (right) thickened rugal folds in an endoscopically confirmed gastritis in Tursiops.
of the GI tract. The appearance and thickness of the gut lumen and wall can be assessed in cases of suspected gastroenteritis or obstruction, peristalsis can be observed in real time, and hyper- or hypomotility identified. GI masses can also be identified and peritoneal effusions detected and evaluated. Ultrasound does not provide much useful information about the main stomach; that is, forestomach. The proximal wall may be imaged for gross pathology. However, there is insufficient information about normal appearance and mural thickness at present for this to be very useful. The stomach is rarely empty and is also hugely distensible, making measurement unreliable. The stomach usually contains free gas, which, even in small amounts, will rise to the transducer side, scattering the ultrasound beam and preventing imaging of the lumen. Sternal recumbency may be useful for examination of the forestomach. Gas rises dorsally, so scanning in the coronal/dorsal plane, ventral to the gas, can improve visualization. The fluid in the forestomach can provide a good acoustic window to the deeper anatomy of the cranial abdomen. Administration of water via stomach tube prior to the examination may also be useful. Besides distending the GI tract and stimulating motility and gastric contraction, it allows visualization of the pylorus and duodenum, and the fluid-filled gut highlights adjacent anatomy (spleen, pancreas). Gastroscopy, however, remains the method of examination of choice for the stomach (see Chapter 27, Endoscopy). In dolphins, the second, or fundic, stomach contains very little free gas and can be seen quite well. It is easily identifiable by its position, thick wall, and distinctive rugae (Figure 13). A marked increase in rugal thickness may indicate gastritis or parasitosis (Woodard et al., 1969) in an animal in which normal appearances are known, although normal ranges have yet to be published. The transition of hyperechoic gastric contents through the second stomach is easy to monitor and may be useful to identify functional abnormalities or mass lesions. The pylorus is obvious on ultrasound, and can be seen extending laterally from the fundic stomach, then reflecting at the medial border of the right liver, to run caudomedially for a short distance within the cranial abdomen. The size, motility, and transition of contents through the pyloric chamber may provide useful information. In one case of clinically suspected obstruction, marked and prolonged distension of the pyloric chamber, with hypermotility and increasing hyperechogenicity of contents, possibly due to microbubble formation, has been seen (Figure 14). A partial obstruction was suspected. After a period
0839_frame_C26 Page 607 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
607
FIGURE 14 Distended, possibly partially obstructed pylorus/duodenum in an adult male bottlenose dolphin (T. t. aduncas). Hyperechogenicity of contents is probably due to microbubble formation.
of some weeks, this suddenly resolved and a few days later the dolphin passed several large, hard objects in its feces. These were composed of impacted sand, which was later discovered to be entering the enclosure through a faulty filter. This dolphin had regularly been observed hovering around the affected inlet. The sonographic appearance of inflammatory GI disorders varies depending on the pathology involved, the site and extent of involvement, and associated complications, such as peritonitis. The relatively fluid caudal bowel contents of marine mammals provide a useful “contrast medium” when examining the GI tract. Gas content in caudal bowel is normally low when compared with that of other species, and changes in mural thickness are more easily seen. Similarly, dense fecal matter seen in other species does not obscure the rectum. Unlike in humans and small animals, it is not easy sonographically to differentiate bowel wall layers in marine mammals, even with the improved resolution of modern equipment. Normal mural thickness values have not been determined, but any pathological increase is usually obvious to the experienced eye (Figure 15). Similarly, hypermotility can be seen and monitored. Acute hypomotility appears to be less common, but in the authors’ experience has been iatrogenically induced by overzealous oral fluid administration and is easily recognizable. Luminal patterns seen depend on contents, although it is rare to see empty bowel segments, particularly in dolphins. The normal appearance in the caudal bowel is that of relatively hypoechoic fluid contents. The presence of large amounts of gas within the bowel lumen or hyperechogenicity of feces may be indicative of disease, but can also be seen when diet is varied, in some cases of stress, or in clinically normal animals. The large pararectal lymph nodes may also be examined (Figure 16). Mesenteric nodes are more difficult to evaluate, possibly because they are invested by smooth muscle, which likely increases their echogenicity, making it almost impossible to differentiate nodes from surrounding tissue. Further research is required to determine the role of ultrasound in assessment of the GI tract in marine mammals. Ultrasound is the least invasive and most accurate method for detecting the presence of ascites in marine mammals (Figure 17). Evaluation of the fluid for particulates, and assessing whether the small bowel is seen to float freely or appear matted and adhered may give further information. Ultrasound can identify a suitable location for paracentesis and guide safe needle placement. The development of ascites in dolphins is generally not a good prognostic indicator.
0839_frame_C26 Page 608 Tuesday, May 22, 2001 11:54 AM
608
CRC Handbook of Marine Mammal Medicine
FIGURE 15 Progressive enteritis in the same animal shown in Figure 14. (Top left) Normal appearance of small bowel; images (top right) and (bottom) show inceasing bowel wall thickness and hypoechogenicity (note same scale).
FIGURE 16 Mildly enlarged, hypoechoic pararectal lymph nodes in two dolphins with clinical signs of enteritis.
0839_frame_C26 Page 609 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
609
FIGURE 17 (Left) Gross ascites in a bottlenose dolphin (T. t. aduncas) calf in hepatic failure following severe weaning difficulties, and (right) mild ascites seen in a pregnant female dolphin with excessive hepatic steatosis and progressively abnormal liver function tests/liver parameters.
Urinary Tract Focal lesions, such as cysts, abscesses, adenomas, mineral deposits, and frank calculi, have been reported in kidneys (Brown et al., 1960; Miller and Ridgway, 1963; Migaki et al., 1978; Howard et al., 1983), but diffuse glomerular disease appears to be rare. Massive interstitial corticomedullary amyloidosis has also been noted (Cowan, 1995). Both dolphin and pinniped kidneys can be examined using ultrasound. In both, the kidneys are large, with renal lengths loosely correlated with body length. The dolphin kidneys are easy to find, located caudal to the dorsal fin, and lying with their medial aspects closely aligned to either side of the midline, tucked up between the large hypaxial muscles. They are located by scanning along the pectoral line, with the transducer in the transverse position and moving either caudally from the dorsal fin or cranially from the cranial end of the genital slit. Kidneys are multilobulated. The individual renules can be seen, each with its own collecting system and a hyperechoic boundary, the overall appearance resembling that of a bag of marbles (Figure 18). The multiple interfaces presented by these anatomical
FIGURE 18 Sonographic appearance of bottlenose dolphin kidneys in (left) a 7-year-old male and (right) a 27-year-old male.
0839_frame_C26 Page 610 Tuesday, May 22, 2001 11:54 AM
610
CRC Handbook of Marine Mammal Medicine
FIGURE 19 (Top) Two small renal calculi are seen in a male bottlenose dolphin (note the acoustic shadows behind each of the foci), and (bottom) a simple renal cyst is shown in the right kidney of another male. (Image credits: U.S. Navy.)
structures produce high-amplitude echoes, which appear to become more pronounced with age (Figure 18, right), and may even produce acoustic shadowing and mimic small calculi or mineral deposits. Scanning in more than one plane should eliminate shadowing if mineralization is not present (Figure 19, top). Pinniped kidneys are more similar in appearance to those of the pig or bear, and are anatomically located in much the same place as in the dog. Because of its more caudal position, the left kidney is easier to see; overlying bowel often obscures the right, which also lies higher under the rib cage and may be inaccessible. With the animal in lateral or dorsal recumbency, the transducer is placed on the ventral abdomen, at the lower costal margin and in the sagittal or coronal/dorsal plane. Firm pressure is usually required to displace overlying bowel loops and visualize kidneys adequately. Once the kidney is located, full sagittal and transverse scans are performed. Sometimes a ventrally located spleen may provide a useful acoustic window for examination of the left kidney in seals. Large amounts of perirenal fat may also help to differentiate kidneys. Relative echogenicity of the kidneys to other reference organs has not been established for these species. The medullae cannot be reliably differentiated in dolphins and, because of the size of the average patient and the anatomy of the dolphin kidney, subtle changes in parenchymal appearance
0839_frame_C26 Page 611 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
611
and echogenicity are unlikely to be detected in marine mammals with currently available equipment. Apart from identification of focal lesions, the value of ultrasound in the evaluation of renal disease in marine mammals is not yet clear. UGFNABs of kidneys have been successfully performed by two of the authors (W.V.B., E.J.). The normal ureters are not seen in these large animals. However, the oval, urine-filled bladder is easy to examine (see Figure 21, right). The bladder wall can be assessed for thickness and any lesions. Wall thickness should be ≤ 3 mm in bottlenose dolphins (T. t. aduncas) when fully distended, but normal measurements have not been documented in other species. The bladder lumen should be anechoic, although, rarely, crystalline debris may be seen circulating in the urine. Bladder calculi have been reported in marine mammals and should present as mobile foci with highly echogenic, curved surfaces and marked distal acoustic shadowing.
Reproductive Tract As for many species, much of the available literature about reproductive tracts relates to the use of ultrasound during pregnancy (Leopold, 1977; Stone et al., 1984; Williamson et al., 1990; Taverne, 1991; Young and Grantmyre, 1992; Barr et al., 1994; Brook et al., 1994; Brook, 1994; Jensen, 1999; Stone et al., 1999; Van Bonn, 1999). Sonographic appearances of the reproductive tract in the male (Brook et al., 1991; 1996; Brook, 1997; 1999) and female (Brook et al., 1992; 1994; 1996; 1999; Robeck et al., 1998) bottlenose dolphin have also been documented. Males
Ultrasound can be used to examine the intra-abdominal testes, epididymes, vasa deferentia, penis, bulbourethral and bulbocavernosal muscles in dolphins, and the scrotum and its contents in the California sea lion (Zalophus californianus). Phocids have a scrotal structure, but inguinal testes, which are not accessible for transcutaneous examination. Dolphin testes are situated in the caudal abdomen, approximately midway between the midline and the ventral edge of the hypaxial muscles. They are easily located and assessed using ultrasound. The testes are elliptical and elongated, with a well-defined border (Figure 20, left). In cross section the testes are rounded (Figure 20, right). Testicular length increases dramatically during the
FIGURE 20 (Left) Coronal/dorsal scan showing longitudinal section, and (right) transverse scan showing transverse section of mature dolphin testes. Note small parenchymal cysts in the right image; these are common and usually resolve spontaneously.
0839_frame_C26 Page 612 Tuesday, May 22, 2001 11:54 AM
612
CRC Handbook of Marine Mammal Medicine
FIGURE 21 Coronal/dorsal scans showing longitudinal section of the testes in (left) a subadult and (right) a juvenile bottlenose dolphin. The partially filled, oval urinary bladder is seen in the midline in the right image.
process of sexual maturation and does not appear to regress in those species without a clearly defined seasonal breeding pattern, such as the bottlenose dolphin, false killer whale (Pseudorca crassidens), and killer whale (Orcinus orca). Three distinct sonographic patterns can be seen in dolphin testes, and these can be used to estimate reproductive status (Brook, 1997; Brook et al., 2000). Mature testes are homogeneous and moderately to highly echogenic. This is the characteristic appearance of mature testes seen in many species of mammal. A lobular pattern may be visible in the parenchyma of older animals (Figure 20, left). The sonographic appearance of the testicular parenchyma in subadults is less well defined (Figure 21, left). The parenchymal echopattern is homogenous, but less echogenic than in mature animals. Immature testes are relatively small, and the testicular parenchyma is hypoechoic and poorly differentiated (Figure 21, right). The epididymis is clearly visualized, from the triangular head at the cranial end to the dilated and convoluted distal end, where it is not possible to distinguish where the epididymis joins the vas deferens. In mature males, rounded protrusions of convoluted tubules may be seen adjacent to the epididymis. The reported incidence of pathology of the dolphin reproductive tract is low, with orchitis apparently the most common disorder. Changes in size, shape, and/or sonographic appearance of the testicular parenchyma should be detectable if orchitis is clinically suspected. This usually presents focal areas of decreased echogenicity. Small, simple cysts, probably due to obstruction of tubules, may occasionally be seen within the testes (Figure 20, right); these usually resolve spontaneously. Sonographic examination of the male reproductive tract in pinnipeds is more of a challenge and usually requires sedation. Scrotal swelling is not uncommon in pinnipeds and ultrasound is very useful to identify scrotal edema (Figure 22), or hydrocele, or to detect testicular lesions. Females
The ovaries in cetaceans are relatively superficial, tucked high in the dorsolateral aspect of the abdomen, to lie in the angle formed by the hypaxialis lumborum (hlm) and rectus abdominus (ram) muscles, against the wall of the abdominal cavity, at a variable distance from the genital slit. There is a palpable depression in the flank where these muscles meet, which can also provide a guide to transducer placement. The transducer is held in the transverse position and placed at the junction of the hlm and ram, at the midpoint of the genital slit, then moved cranially until the transverse axis of the ovary is identified (Figure 23, left). The transducer can
0839_frame_C26 Page 613 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
613
FIGURE 22 Sagittal scan through the distended scrotum of a fur seal (Callorhinus ursinus) showing massive edema.
FIGURE 23 (Left) LS and (right) transverse section images of the ovary in a bottlenose dolphin. Note smaller follicle in the right image.
then be rotated 90° to visualize the long axis of the ovary (Figure 23, right). It may be necessary to angle the transducer slightly dorsally. The position of the ovary varies quite widely, and they may be seen anywhere from the caudal pole of the ipsilateral kidney down to the level of the uterine body. This variation in position is particularly seen in females that have given birth, and is likely due to loss of integrity of suspensory structures. It is possible routinely to identify and assess the ovaries of all species of cetaceans studied, including harbor porpoises (Phocoena phocoena), bottlenose dolphins, white-sided dolphins (Lagenorhynchus obliquidens), belugas (Delphinapterus leucas), false killer whales, and killer whales (Figure 24). To the authors’ knowledge, there is no report of the sonographic appearance of the ovaries in any pinniped. Sonography can be more difficult in larger, older, or fatter females, or when ovaries are inactive due to lactation or prolonged periods of anestrus, yet visualization can usually be achieved with perseverance. The ovarian cortex in older females is generally more echogenic than in younger animals, and the outline is more irregular. In the long axis, a double, echogenic “tram line” appearance, representing the reflections of the mesovarium in the
0839_frame_C26 Page 614 Wednesday, May 23, 2001 12:25 PM
614
CRC Handbook of Marine Mammal Medicine
FIGURE 24 LS images of the ovary in (top) a killer whale and (bottom) a white-sided dolphin. (Image credits: T. Robeck.)
FIGURE 25 Sonographic appearance of follicles in bottlenose dolphins. Note thickened, hyperechoic wall in the left image. This developed into an anovulatory follicular cyst of 5.0-cm diameter before spontaneous resolution. Multiple follicular development seen in the right image is common, especially in older females.
0839_frame_C26 Page 615 Tuesday, May 22, 2001 11:54 AM
615
Ultrasonography
hilum, is a useful marker for identifying the ovary (see Figures 23 and 24). Small, echolucent areas seen within the hilum of some ovaries may represent blood or lymphatic vessels. Free fluid occasionally seen within the hilum represents normal peritoneal fluid. Small, quiescent, antral follicles as large as 4 mm in diameter are occasionally seen, whether a dolphin is exhibiting ovarian activity or not. Developing follicles are always prominent and easily identifiable by ultrasonography, even in older and larger dolphins, when it is sometimes difficult to distinguish the ovary clearly. A developing follicle presents the classic, rounded, cystic appearance seen in most mammals (Figure 25). The corpus luteum (CL) can also be seen and monitored. The usual appearance of a CL in a nonconceptive cycle is a rounded, solid mass with unclear margins at the site of the ruptured follicle, which may appear isoechoic or slightly more echogenic than the ovarian parenchyma. Rarely, a nonconceptive CL may appear hypoechoic. Hypoechoic CLs can be distinguished from follicles by their larger size, thicker walls, and lack of distal acoustic enhancement. CLs of pregnancy in bottlenose dolphins tend to be larger than those of nonconceptive cycles, and appear hypoechoic, organized, and more regular in outline. This appearance is a useful sign for diagnosis of early pregnancy before the conceptus is visualized. Occasionally, and particularly if associated with pregnancy, the CL may cavitate (Figure 26). The use of ultrasound to monitor the ovarian cycle in dolphins has added much to knowledge of reproductive physiology in some species and has allowed the application of controlled, selective breeding programs and artificial insemination (see Chapter 11, Reproduction). Pathology of the ovaries, including ovarian carcinoma, mucinous cystadenoma, and granulosa cell tumors, has been reported in cetaceans (Cockrill, 1960), and ovarian cysts (Munson et al., 1998; Robeck et al., 2000) are not unusual. Fibromyoma appears to be the most commonly identified uterine abnormality (Cockrill, 1960).
A
B
C
D
FIGURE 26 Sonographic appearances of CLs in bottlenose dolphins (T. t. aduncas): (A) CL of a nonconceptive cycle; (B) persistent, “cystic” CL (note lack of acoustic enhancement behind); (C) CL of early pregnancy (2 to 3 weeks gestation); (D) cavitated CL in later pregnancy (26 weeks).
0839_frame_C26 Page 616 Tuesday, May 22, 2001 11:54 AM
616
CRC Handbook of Marine Mammal Medicine
To examine the uterus, the transducer is placed in the longitudinal plane, just above the genital slit and moved dorsally until the cervix can be visualized lying just dorsal to the urinary bladder. The transducer is then moved cranially, with short dorsoventral sweeps to examine the uterine body and ipsilateral horn. Again, the procedure is repeated for the other side. The uterine cervix can be seen to angle dorsally, immediately behind and/or to one side of the urinary bladder. There may be mucus present in the cervical canal, which may delineate the internal architecture and allow visualization of the pseudocervix and spermothecal recess. The uterus is bicornuate and relatively large and the uterine horns curve backward along their length, toward the ovaries. The uterus is usually surrounded by gut, and therefore not always easy to examine. The normal uterus is soft and very malleable, with a thin, poorly demarcated myometrium. It presents as an amorphous, soft tissue “mass” on ultrasonography. Sections of the endometrium may also be seen. Further study is required to be able to characterize the ultrasonographic appearance of the endometrium in all species and to compare changes during periods of ovarian activity. Sonography of the pregnant uterus is described in Chapter 11 (Reproduction).
Eyes B-mode ultrasonographic examination of the eye has been performed in veterinary medicine for more than two decades, and there are numerous references about sonographic ophthalmic techniques and appearances of intraocular and retrobulbar pathology in the literature (Johnston and Feeney, 1980; Dziezyc et al., 1987; Fielding, 1987; Nyland and Mattoon, 1995). This technique can also be used for examination of marine mammals (Cartee et al., 1995) using a standard ultrasound unit, in conjunction with a 7.5- to10-MHz transducer. A 5-MHz transducer may be needed for the orbit. Scanning is performed through the eyelid, using an acoustic coupling gel. Both pinnipeds and dolphins can be examined without anesthesia if trained to station for examination. In the authors’ experience, direct contact with the corneal surface of the eye, as carried out for small animals, is not as acceptable to marine mammals. Indications for eye sonography include trauma, photophobia, retraction of the globe, complete or partial closure, or any ocular opacity. A transonic standoff pad is useful for examination of the cornea, anterior chamber, or eyelids, and improves visualization of the lens, as it decreases reverberation artifacts in the near field. Full sagittal and transverse scans of both eyes for comparison need to be carried out. The transducer is placed on the optic axis and then “swept” in an arc across the whole globe. There are no established normal axial measurements for these species. Retinal detachments (Figure 27), intraocular hemorrhage, subluxation of the lens, and lacrimal gland hyperplasia (Figure 28) have been demonstrated using this technique in marine mammals.
Musculoskeletal System Ultrasonography is an ideal method for diagnosing and monitoring muscular injury, and can also be used to evaluate superficial skeletal structures. This can be very useful when radiography is not available or even, in some cases, to add further information when nonspecific radiological abnormalities are found. Muscle pathology, particularly abscess formation, is common in cetaceans (Brown et al., 1960; Cockrill, 1960) and may have serious consequences. Abscesses, granulomas, osteolytic lesions (Figure 29) and fractures have all been diagnosed in dolphins using sonography.
0839_frame_C26 Page 617 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
617
FIGURE 27 Retinal detachments shown in a (left) California sea lion (Z. californianus) and (right) bottlenose dolphin (T. truncatus).
FIGURE 28 Sonographic images of (top) collapsed globe with subluxation of the lens, retinal detachment, and hemorrhage and (bottom) lacrimal gland hyperplasia in an elderly male T. t. aduncas.
0839_frame_C26 Page 618 Tuesday, May 22, 2001 11:54 AM
618
CRC Handbook of Marine Mammal Medicine
FIGURE 29 Sonographic appearance of (left) normal vertebral body and spinous process and (right) severe spondilysis and degeneration resulting from Staphylococcus aureus infection.
Body Condition Sonography is an excellent means of measuring and tracking blubber and subcutaneous adipose tissue thickness. It is important that the site of determination be standardized between examinations in a given individual, and between individuals, if inferences are to be drawn about differences. The blubber and subcutaneous fat layers tend to be thickest in the cervical region of cetaceans, and if one site is to be chosen, this may be the most valuable. However, this site is also very dynamic, and correlations with factors such as caloric intake, age, gender, season, and water temperature have not been completely tested to date, but hold promise as excellent indicators of body condition change.
Conclusion The use of ultrasonography in the care and management of marine mammals has increased during the last decade. Many now recognize it as being safe, noninvasive, cost-effective, and very valuable as a means of diagnosis and monitoring of many conditions in these large animals, often when other types of imaging are impossible or unavailable. Modern units are relatively inexpensive and image quality has improved dramatically. What is needed now is for more marine mammal clinicians to practice utilizing this technology and for those who already have to better share their experiences and the information they are gathering, so that others can utilize that experience to the benefit of the animals in their care.
Acknowledgments The authors thank all of the colleagues who work and have worked with them, facilitating their role as “imagers,” and Reimi Kinoshita and Natalie Rourke for reviewing this chapter.
References AIUM (American Institute of Ultrasound in Medicine), 1988, Bioeffects considerations for the safety of diagnostic ultrasound, J. Ultrasound Med., 7: 51–53.
0839_frame_C26 Page 619 Tuesday, May 22, 2001 11:54 AM
Ultrasonography
619
Baker, J.R., 1992, Causes of mortality and parasites and incidental lesions in dolphins and whales from British waters, Vet. Rec., 130: 569–572. Barr, L.L., Gillespie, D., Campbell, M.K., and Babcock, D., 1994, Prenatal sonography: A useful adjunct in the management of the gravid captive walrus (Odobenus rosmarus divergens), J. Ultrasound Med., 13: 485–486. Bossart, G.D., Ewing, R., Herron, A.J., Cray, C., 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: 454–459. Brook, F.M., 1994, Ultrasound diagnosis of anencephaly in a bottlenose dolphin, J. Zoo Wildl. Med., 25: 569–573. Brook, F.M., 1997, The Use of Diagnostic Ultrasound in Assessment of the Reproductive Status of the Bottlenose Dolphin, Tursiops aduncas, in Captivity and Applications in Management of a Controlled Breeding Programme, Ph.D. thesis, The Hong Kong Polytechnic University, Kowloon, Hong Kong, 339 pp. Brook, F.M., 1999, Sonographic ovarian and testicular evaluation in bottlenose dolphins, Tursiops truncatus aduncas, in Proceedings of Bottlenose Dolphin Reproductive Workshop, 207–222. Brook, F.M., Chow, D.C.M.A., and Schroeder, J.P., 1991, Ultrasound imaging of the reproductive tract of the male bottlenose dolphin, Br. J. Radiol., 763: 645. Brook, F.M., Chow, D.C.M.A., and Schroeder, J.P., 1992, Ultrasound imaging of the reproductive tract of the female bottlenose dolphin, Tursiops aduncas, Br. J. Radiol., 775: 628. Brook, F.M., Kinoshita, R., Chan, S.Y., and Hui, S.W., 1994, Sonographic assessment and monitoring of the reproductive status of the bottlenose dolphin, Tursiops aduncas, in captivity, Abstr., Proceedings of the 4th Annual Conference on Southeast Asian Zoology, 179. Brook, F.M., Kinoshita, R., Chan, S.Y., and Hui, S.W., 1996, Applications of sonography in controlled breeding of captive bottlenose dolphins at Ocean Park, Hong Kong, Proceedings of the 6th Annual Conference on Southeast Asian Zoology, 48. Brook, F.M., Kinoshita, R., Brown, B., and Metreweli, C., in press, Sonographic anatomy of the testes and epididymis of the bottlenose dolphin, Tursiops truncatus aduncas, J. Reprod. Fertil. Brown, D.H., McIntyre, R.W., Delli Quadri, C.A., and Schroeder, R.J., 1960, Health problems of captive dolphins and seals, J. Am. Vet. Med. Assoc., 137: 534–538. Cartee, R.E., Brosemer, K., and Ridgway, S.H., 1995, The eye of the bottlenose dolphin (Tursiops truncatus) evaluated by B mode ultrasonography, J. Zoo Wildl. Med., 26: 414–421. Cockrill, W.R., 1960, Pathology of the cetacea. Part 1, Br. Vet. J., 116: 133–144. Cowan, D.F., 1994, Involution and cystic transformation of the thymus in the bottlenose dolphin, Tursiops truncatus, Vet. Pathol., 31: 648–653. Cowan, D.F., 1995, Amyloidosis in the bottlenose dolphin, Tursiops truncatus, Vet. Pathol., 32: 311–314. Duignan, P.J., Geraci, J.R., Raga, J.A., and Calzada, N., 1992, Pathology of morbillivirus infection in striped dolphins (Stenella coeruleoalba) from Valencia and Murcia, Spain, Can. J. Vet. Res., 56: 242–248. Dziezyc, J., Hager, D.A., and Millichamp, N.J., 1987, Two-dimensional real-time ocular ultrasonography in the diagnosis of ocular lesions in dogs, J. Am. Anim. Hosp. Assoc., 23: 501–508. Fielding, J.A., 1987, Ultrasound imaging of the eye through the closed lid using a non-dedicated scanner, Clin. Radiol., 38: 131–135. Gray, K.N., and Conklin, R.H., 1974, Multiple births and cardiac anomalies in the bottlenosed dolphin, J. Wildl. Dis., 10: 155–157. Howard, E.B., Britt, J.O., and Simpson, J.P., 1983, Neoplasms in marine mammals, in Pathology of Marine Mammal Diseases, Vol. 2, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 95–162. Hykes, D.L., Hedrick, W.R., and Starchman, D.E., 1992, Ultrasound Physics and Instrumentation, Mosby Year Book, St. Louis, MO, 295 pp. Jensen, E., 1999, Early embryonic loss, in Proceedings Bottlenose Dolphin Reproductive Workshop, 273–278.
0839_frame_C26 Page 620 Tuesday, May 22, 2001 11:54 AM
620
CRC Handbook of Marine Mammal Medicine
Johnston, G.R., and Feeney, D.A., 1980, Radiology in ophthalmic diagnosis, Vet. Clin. North Am. Small Anim. Pract., 10: 317–337. Laing, F.C., 1983, Commonly encountered artifacts in clinical ultrasound, Semin. Ultrasound, 4: 27–43. Leopold, G.R., 1977, Role of diagnostic ultrasound in evaluating pregnancy in Tursiops truncatus, in Breeding Dolphins; Present Status, Suggestions for the Future, Ridgway, S.H., and Benirschke, K. (Eds.), MMC report 76/07, 217–219. Migaki, C., Woodward, J.C., and Goldston, R.T., 1978, Renal adenoma in an Atlantic bottlenose dolphin (Tursiops truncatus), Am. J. Vet. Res., 39: 1920–1921. Miller, R.M., and Ridgway, S.H., 1963, Clinical experiences with dolphins and whales, Small Anim. Clin., 3: 189–193. Munson, L., Calzada, N., Kennedy, S., and Sorensen, T.B., 1998, Luteinized ovarian cysts in Mediterranean striped dolphins, J. Wildl. Dis., 34: 656–660. Nyland, T.G., and Mattoon, J.S., 1995, Veterinary Diagnostic Ultrasound, W.B. Saunders, Philadelphia, 357 pp. Rawson, A.J., Patton, G.W., and Brooks, J.S., 1992, Lymphangiomyomatosis in the Atlantic bottlenose dolphin (Tursiops truncatus), J. Wildl. Dis., 28: 323–325. Reeves, R.R., DeMaster, D.P., Hill, C.L., and Leatherwood, S., 1994, Survivorship of odontocete cetaceans at Ocean Park, Hong Kong, 1974–1994, Asian Mar. Biol., 11: 107–124. Rhinehart, H., Townsend, F., Gorzelany, J., and Broecker, S., 1995, Ultrasound-aided thoracentesis of a bottlenose dolphin, unreferenced paper. Robeck, T.R., McBain, J.F., Mathey, S., and Kraemer, D.C., 1998, Ultrasonographic evaluation of the effects of exogenous gonadotropins on follicular recruitment and ovulation induction in the Atlantic bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 6–13. Robeck, T.R., Jensen, E., Brook, F., Rourke, N., Rayner, C., and Kinoshita, R., 2000, Preliminary investigations into ovulation manipulation techniques in delphinids, in Proceedings of Joint Meeting of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine, New Orleans, LA, 222–225. Sanders, R.C., 1998, Clinical Sonography. A Practical Guide, 3rd ed., Lippincott, Philadelphia, 613 pp. Stone, L.R., 1990, Diagnostic ultrasound in marine mammals, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L. (Ed.), CRC Press, Boca Raton, FL, 235–264. Stone, L.R., Phillips, B., and Sweeney, J.C., 1984, Diagnostic ultrasound of the bottlenose dolphin, Abstr., Annual Proceedings of the American Association of Zoo Veterinarians, 94. Stone, L.R., Johnson, R.L., Sweeney, J.C., and Lewis, M.L., 1999, Fetal ultrasonography in dolphins with emphasis on gestational aging, in Zoo and Wild Animal Medicine. Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 501–506. Taverne, M.A.M., 1991, Applications of two-dimensional ultrasound in animal reproduction, Wien. Tierärztl. Monatsschr., 78: 341–345. Thickman, D.I., Ziskin, M.C., Jacobs Goldenberg, N., and Linder, B.E., 1983, Clinical manifestations of the comet tail artifact, J. Ultrasound Med., 2: 225–230. Van Bonn, W., 1999, Infectious diseases and late term abortions, in Proceedings Bottlenose Dolphin Reproductive Workshop, 279–288. Williamson, P., Gales, N.J., and Lister, S., 1990, Use of real-time B-mode ultrasound for pregnancy diagnosis and measurement of fetal growth rate in captive bottlenose dolphins (Tursiops truncatus), J. Reprod. Fertil., 88: 543–548. Wislocki, G.B., 1929, On the structure of the lungs of the porpoise (Tursiops truncatus), Am. J. Anat., 44: 47–72. Woodard, J.C., Zam, S.G., Caldwell, D.K., and Caldwell, M.C., 1969, Some parasitic diseases of dolphins, Pathol. Vet. (Swiss), 6: 257–272. Young, J.S., and Grantmyre, E.B., 1992, Real-time ultrasound for pregnancy diagnosis in the harbour seal (Phoca vitulina concolor), Vet. Rec., 130: 328–333.
0839_frame_C27 Page 621 Tuesday, May 22, 2001 11:55 AM
27 Flexible and Rigid Endoscopy in Marine Mammals Samuel R. Dover and William Van Bonn
Introduction Endoscopy means “to look inside.” The term is usually reserved for examination of the interior of hollow viscera such as bronchi or the intestinal tract, but it can also be applied to examination of the abdominal cavity (laparoscopy) or the thorax (thoracoscopy) (Freeman, 1999). Endoscopes can be flexible or rigid. Flexible endoscopes have been used in marine mammal medicine for many years, primarily for examination of the gastrointestinal and respiratory systems, and should be considered a required diagnostic tool in any marine mammal practice. The rigid endoscope (or telescope) always requires a straight path to the organs being examined. It is the veterinarian and/or the indication that determines the type of scope to be used (Lunemann, 1999). Flexible endoscopes utilize either fiber-optic or video chip technology. The primary difference between the two is a trade-off between image qualities vs. price. Video technology provides images that are far superior to fiber optic; however, they are two to three times more expensive to purchase. The authors’ advice is to buy the best-quality scope one’s budget will allow, and avoid purchasing old fiber-optic scopes with damaged fibers. The quality of the image obtained can easily affect the diagnosis and outcome of the treatment; it is important to be sure one is collecting the best information possible. Minimally invasive surgery (MIS), including laparoscopy and thoracoscopy, is a collection of surgical techniques designed to minimize the extent of the anatomical approach while still maintaining precision and efficiency. Endoscopic surgery involves performing a minimally invasive surgical procedure with visualization provided by an endoscope (Freeman, 1999). Noninvasive endoscopy vs. MIS involves the progression from flexible to rigid scopes. One may also use rigid scopes in some applications where flexible scopes were commonly used, or vice versa. In most cases, the use of rigid endoscopes requires higher planes of anesthesia to prevent injury to the animal and/or damage to the scope. Laparoscopy has a long history of use in veterinary medicine, including use in exotic and zoo animals (Murray et al., 1998). In comparison with most traditional open surgical procedures, MIS induces relatively minor tissue trauma, which, in most cases, results in shorter postoperative recovery periods, decreased postoperative care, and fewer postoperative complications 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
621
0839_frame_C27 Page 622 Tuesday, May 22, 2001 11:55 AM
622
CRC Handbook of Marine Mammal Medicine
(Cook and Stoloff, 1999). These factors make MIS ideally suited for marine mammal applications. Although open surgical techniques have been used successfully in a variety of marine mammals, there are frequent complications (Sweeney, 1990). MIS allows the surgeon to overcome several of the complications associated with difficult anatomical approaches, as well as wound healing and dehiscence. Additionally, in the case of cetaceans, there is a limited length of time that animals can be safely kept out of water. The small surgical wounds (5 to 12 mm or less) produced during MIS procedures are easily rendered impervious to leakage, which can allow any marine mammal to be returned rapidly to the water postoperatively. This chapter discusses the types of equipment required for both flexible endoscopy and laparoscopy. The indications, limitations, techniques, and clinical applications of both flexible and rigid endoscopy are presented with examples of successfully performed procedures in several marine mammal species.
Indications Flexible endoscopy is indicated for examination and treatment of the gastrointestinal, respiratory, and urogenital tracts. The most common clinical indication for flexible endoscopy in marine mammal care is the examination of the esophagus and forestomach of the dolphin. Detection and removal of foreign materials, thorough evaluation of the mucosa and motility, and collection of specimens for cytology, microbiology, and histological or special studies are readily accomplished in most species. The primary differences among species are in their physical restraint and handling. Respiratory tract endoscopy is rapidly proving itself to be an equally valuable and safe technique in marine mammals (Hawkins et al., 1997; Van Bonn et al., 1997). Rhinoscopy, pharyngoscopy, tracheoscopy, and bronchoscopy are surprisingly well tolerated in most cetaceans, with little or no sedation. These procedures usually require the use of general anesthesia for marine mammals other than cetaceans (see Chapter 29, Anesthesia). The increasing use of these techniques will advance the knowledge of respiratory pathology and the response of cetacean lungs to disease. Examinations of the urogenital tract in females can be performed. Although this is usually performed under sedation and restraint, several Atlantic bottlenose dolphins (Tursiops truncatus) have been trained to tolerate this procedure while stationed in the water. The use of flexible endoscopes to guide advanced reproductive techniques such as artificial insemination of small and large cetaceans is also rapidly emerging as a valuable clinical method (Robeck et al., 1994; Robeck, 2000). Although less commonly encountered in practice, the authors have had reason to perform endoscopic examinations of the rectum and colon in several bottlenose dolphins and California sea lions (Zalophus californianus). All may be performed with little difficulty and low risk to the patient, and all provide valuable information to support clinical decision making. The most common indication for MIS in veterinary medicine is exploratory visualization of internal organs and subsequent biopsy collection. However, with training and experience, one can include surgical treatment options as well. MIS has been used in zoological and wildlife medicine for many years, most commonly for reproductive procedures (Cook and Stoloff, 1999). Laparoscopic appendectomy, antireflux surgery, adhesiolysis, small bowel resection, hernia repair, splenectomy, lymphadenectomy, and liver biopsy are routinely performed in humans (Freeman, 1999). For certain procedures, such as cholecystectomy, the laparoscopic approach is performed more commonly than the conventional laparotomy technique. New MIS techniques for conventional surgical procedures are continually being developed and refined, and one can apply these advances to improve the care of marine mammal patients.
0839_frame_C27 Page 623 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
623
In most cetacean species, practitioners have been limited to noninvasive procedures for the medical management of critically ill animals. Techniques such as radiology, ultrasound, and laboratory work can provide various pieces of the diagnostic puzzle, but many times will miss the one critical piece that can complete the picture, a visual examination of the suspected location of a lesion. While conventional surgical approaches have been used successfully in odontocetes, complications are frequently encountered. The surgical approach to the abdominal cavity in both pinnipeds and cetaceans offers a host of complicating factors to the surgeon (Sweeney, 1990). As described in the introduction, MIS techniques overcome many of these problems.
Limitations The primary limitation of endoscopy in marine mammals is the physical dimensions of the equipment available to the practitioner. The adult bottlenose dolphin requires a minimum scope length of 150 cm for a thorough gastroscopy. Several veterinary fiber-optic endoscope manufacturers have models in the 1.5 to 2.0 m lengths (Karl Storz, Pentax, Schott). Larger marine mammals, including killer whales (Orcinus orca), belugas (Delphinapterus leucas), pilot whales (Globicephala spp.), adult walrus (Odobenus rosmarus), and manatees (Trichechus manatus), usually require a flexible gastroscope longer than 2 m. However, scopes designed for specific human procedures are commercially available in a wide variety of configurations, and many can be adapted for various applications in marine mammals. Human colonoscopes are typically 2 m in length, but their large diameters limit their use in smaller pinnipeds or mustelids. Endoscopes <1 m (3.3 ft) in length can be used in smaller pinnipeds and mustelids. The diameter of most commercially available scopes of sufficient length to perform bronchoscopy on mature dolphins limits their introduction to the level of approximately sixthgeneration bronchi. Smaller-diameter scopes can be introduced into deeper airways, but have much smaller working channel diameters, and restrict lavage fluid flow rates. This has been a limiting factor to establishing acceptable standardized bronchoalveolar lavage techniques. The use of smaller-diameter custom equipment is expected to provide more diagnostic lavages, and will lead to standardization of cell counts and technique. Environmental limitations occur when these procedures are performed poolside or outdoors. The use of AC powered equipment near salt water or during inclement weather carries some electrocution hazard risk; therefore, only power sources with ground-fault interrupters should be used. Video monitors and cameras that allow for real-time viewing are designed for use with indoor lighting, so they can be difficult to view with sun glare. This can be overcome by the use of “heads-up” displays that are worn as headgear, or in the case of fiber-optic endoscopes, viewing through the eyepiece or ocular lens. There are several newer models of video recorders that contain an integrated screen to allow poolside viewing and recording. Physical limitations on restraining and anesthetizing marine mammals are frequently encountered. In many cetaceans, these can be overcome through training and conditioning. The anatomy of the cetacean upper gastrointestinal tract restricts endoscopic examination to the pharynx, esophagus, forestomach, and limited evaluation of the glandular (second) stomach. Endoscopic examination of the intestinal tract proper is limited to the retrograde introduction of the scope via the rectum and the length and diameter of the scope itself. The indications for endoscopy in pinnipeds are more limited, and these animals are not usually trained for the procedure. Gastrointestinal endoscopy can be performed in sedated, well-restrained sea lions; however, a more complete examination is possible with the animal under general anesthesia. Pharyngoscopy, tracheoscopy, and bronchoscopy can be performed successfully in conscious cetaceans. These examinations require anesthesia in pinnipeds.
0839_frame_C27 Page 624 Tuesday, May 22, 2001 11:55 AM
624
CRC Handbook of Marine Mammal Medicine
There are several additional limitations for the use of MIS techniques in marine mammals. The equipment and instrumentation can be costly to purchase and in most cases are not designed for larger species. Anesthetic challenges vary among the species, and can be prohibitive in many cases. The creation of the pneumoperitoneum or pneumothorax, required for many MIS procedures, can result in hypercarbia and inadequate ventilation, reduced cardiac output, and gas embolism (Quandt, 1999). These physiological limitations can be reduced through the use of positive-pressure ventilation and careful monitoring. Hemorrhage can be challenging to manage if the surgeon is not properly prepared. Although electrosurgery can treat minor bleeding, some complications require conversion from a laparoscopic procedure to an open laparotomy. Training and experience can minimize most of the limitations associated with MIS. Contraindications for MIS include a known ruptured diaphragm, hemodynamic instability, an uncorrected coagulopathy, generalized peritonitis, severe cardiopulmonary disease, abdominal wall infection or defect, multiple previous abdominal procedures, and pregnancy.
Equipment There are many manufacturers and suppliers of new and used endoscopic systems. All systems have at least three components: (1) the endoscope (flexible or rigid), (2) a light source, and (3) a light-transmitting cable. All endoscopic procedures should be documented through the use of still or video photography. These images will prove valuable for following the progression of lesions or response to treatment and should be included in the animal’s individual record (see Chapter 24, Overview of Diagnostic Imaging). There are numerous accessories designed for both diagnostic and therapeutic purposes. These include instruments for biopsy, grasping, aspiration, cytology, electrosurgery, and laser surgery. Pumps are required for suction, insufflation, and irrigation (Chamness, 1999).
Flexible Endoscopes The two types of flexible endoscopes are the fiber-optic and the video endoscope. The difference between the two relates to the systems for acquiring and transmitting images. Higher image quality is the primary advantage of a videoscope over the fiber-optic scope. The fiber-optic image is carried from the distal optics of the scope to the eyepiece through bundles of optical glass fibers. Video endoscopes transmit the image electronically from the tip of the endoscope to the video monitor by using a solid-state silicon computer chip or CCD (charge-coupled device). Recent developments in technology allow for affordable CCD cameras that can be directly attached to the objective of fiber-optic scopes, converting the optical image to an electronic signal that is displayed on a video monitor and recorded for further examination. Cameras used in endoscopy can also utilize digital enhancement and filters to improve the image quality of fiber optics, approaching that of videoscopes (Chamness, 1999). There is a significant cost advantage to this approach, since the same camera, monitor, and recording devices can be used for both flexible and rigid systems. Flexible endoscopes are available in diameters ranging from 14 mm to less than 1 mm. For all but the smallest marine mammals, the larger-diameter scopes are commonly used. The length of the endoscope required depends on the species and procedure to be performed. Unfortunately, there is no one, multipurpose scope that will meet all species needs. Most larger-diameter scopes are equipped with an accessory channel and a deflectable tip, as well as a water channel used to clean the distal optics. Several excellent fiber-optic scopes are made specifically for veterinarians in the 1- to 2-m lengths. Additionally, there are many scopes made specifically for human procedures that will meet a variety of needs, such as colonoscopes or bronchoscopes.
0839_frame_C27 Page 625 Wednesday, May 23, 2001 10:52 AM
Flexible and Rigid Endoscopy in Marine Mammals
625
FIGURE 1 (Top) Typical gastrointestinal fiber-optic endoscope; (Bottom) position in which the handpiece is held. (Courtesy of Karl Storz Veterinary Endoscopy-America, Inc., Goleta, CA.)
Several manufacturers will custom-build endoscopes to specified lengths and diameters, but this can be quite costly. A typical flexible endoscope has three primary sections (Figure 1): the insertion tube, the handpiece, and the umbilical cord. The knobs on the handpiece control tip deflection. Although most larger-diameter scopes will deflect in two planes (up/down, left/right), some smaller scopes, such as bronchoscopes, will only deflect in one plane. Two-plane deflection is desirable for performing a complete gastrointestinal examination. The handpiece also contains the ocular lens, air/water, and suction control buttons, and the opening for the accessory channel. Endoscopes must be handled carefully to avoid damage to the fibers or control cables within the insertion tube. The lens on both the ocular and distal tip must be handled and cleaned carefully to avoid scratches or cracks. The insertion tube must be protected to prevent breaks in the outer surface; failure to identify and repair leaks in this covering can lead to damage of the internal components. The umbilical cord contains the light guide cable that attaches to the light source
0839_frame_C27 Page 626 Tuesday, May 22, 2001 11:55 AM
626
CRC Handbook of Marine Mammal Medicine
FIGURE 2 Hopkins rod lens telescopes. (Top) 10-mm diameter, 30° angled view; (Middle) 5-mm diameter, 0° view; (Bottom) 2.7-mm diameter, 30° angled view. (Image credit: S. Dover.)
and the connections for the insufflation/irrigation pump. Disinfection of the endoscope should be performed prior to and following all examinations, using manufacturer’s recommendations.
Rigid Telescopes There are many manufacturers of rigid telescopes. However, the Hopkins rod lens system is considered by many to be the best available. Light is transmitted through a series of glass rods with lenses at the ends, rather than through air as in other lens systems. This improves light transmission, allowing for higher resolution and a wider and larger image. The development of rod-shaped lenses has allowed for superior photographic and video documentation. There are a wide variety of diameters of rigid telescopes available, with outer diameters typically ranging from 1 to 10 mm (Figure 2). Larger-diameter telescopes have greater light-carrying capacity and larger image size. As with flexible endoscopes, no one size of rigid telescope is suitable for all procedures. For most marine mammals, a 5- or 10-mm diameter telescope will be required for large abdominal spaces. The viewing angle of the telescope affects both the surgeon’s view and orientation. Forwardviewing telescopes (0°) provide the simplest bearings, but the most limited view. Although more challenging to operate, 30° viewing angles allow a much larger field of view by rotating the telescope along its long axis (Chamness, 1999). Telescopes with other viewing angles, including 90° and reverse angles, are available. The choice of telescope will be largely dependent on the procedure and the experience of the surgeon.
Light Sources The quantity and quality of light is a critical factor in obtaining diagnostic images. The two parameters that influence quantity and quality of light from a source are the power (wattage) and the type of illumination technology. The three most common types of light sources are xenon, metal-halide, and tungsten-halogen. They range in power from 25 to 500 W. In general terms, the higher the wattage, the brighter the illumination; however, the type of light source will determine the amount of wattage required for a given procedure. Wattage (in these terms) is not a measure of how much light is produced, but, rather, how much power is consumed. For example, a 40-W incandescent bulb produces less light than a 40-W fluorescent bulb, while consuming the same amount of power. The same principle applies to the difference between tungsten-halogen vs. xenon light sources. A 150-W xenon light source is much brighter than
0839_frame_C27 Page 627 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
627
a 150-W tungsten-halogen bulb. Additionally, the xenon light produces a much truer color replication, because the spectrum of light produced by the bulb closely approximates that of pure sunlight. The primary advantage of tungsten-halogen light sources is the lower purchase price and bulb replacement cost, as compared with xenon. The disadvantages of tungsten-halogen bulbs are that they do not produce as true color reproduction, and there is loss of much of their intensity after only a fraction of the estimated life span (approximately 100 hours). The superior quality and quantity of light produced by a 300-W xenon light source make it the preferred option (Chamness, 1999). The type of scope used will affect the amount of light reaching the area of interest (e.g., gastrointestinal tract, abdominal cavity). The image seen through a large-diameter flexible fiberoptic scope will appear much brighter than a small-diameter scope, even if the same power and light source is used. Similarly, when using a rigid scope, the larger-diameter scope has a much brighter image. Practically speaking, a larger animal requires more light because the area to be illuminated is much larger. This is especially true in laparoscopic procedures on large marine mammals. Laparoscopy on a walrus requires a 300-W xenon source, a 10-mm-diameter telescope, and a large-diameter light guide cable. The same procedure on a sea otter (Enhydra lutris) could be performed with a 150-W halogen light source and a 2.7-mm-diameter telescope. It is also possible to use too much light, which causes the color of the tissue to appear washed out. This problem is easily remedied by paying careful attention to the intensity setting of the light source and adjusting as needed, even during a procedure, as the situation warrants.
Accessories and Instruments There are a variety of accessory instruments available for use with both rigid and flexible endoscopes. The most common uses of flexible instrumentation are for biopsy and foreign body retrieval, and there is a wide selection of available styles (Figure 3). The instrument required will depend on the task at hand and the experience of the operator. Care must be taken to avoid damage to the operating channel of the scope. Damage can occur by using instruments that are too large for the operating channel, using excess force when manipulating the instrument, passing the instrument through a deflected tip, or attempting to remove foreign bodies through the scope, rather than removing the entire scope after the object has been grasped. Specimens for cytology, microbiology, histology, and other special tests can be collected using standard equipment and methods described for other veterinary species or humans (Jones, 1990). Foreign material retrieval, especially in cetaceans, often requires creative construction of custom tools designed around specific material and species presentations (Pieterse et al., 1998; Stetter et al., 1999; Applegate et al., 2000). The method of manually removing items from the forestomach of dolphins with the bare arm is an old technique and not recommended (Beroza and Barclay, 1981). This method significantly compromises the patient and has several limitations. Animals with an arm down the esophagus are not able to breath due to displacement of the larynx; the operator’s arm is often too short to reach the forestomach lumen and places pressure over the base of the heart during the procedure. The authors have also seen several animals that demonstrated elevated serum transaminase activity following this type of procedure (unpubl. data). The type and variety of rigid instrumentation is continually changing as the needs of the market demand. The two primary categories are disposable instruments (commonly used in human procedures) and reusable instruments. Typically, instruments will be passed through trocars that are strategically placed to allow access to the organ of interest. The most common indications are exploratory laparoscopy followed by biopsy of selected organs or lesions.
0839_frame_C27 Page 628 Tuesday, May 22, 2001 11:55 AM
628
CRC Handbook of Marine Mammal Medicine
FIGURE 3 Flexible instruments for use with endoscopes that have an accessory channel. (Courtesy of Karl Storz Veterinary Endoscopy-America, Inc., Goleta, CA.)
Although there are specific biopsy forceps created for these techniques, the use of forceps for tissue manipulation (curved or straight, traumatic or atraumatic), suture material or staples for control of hemorrhage, and scissors (curved or straight, sharp or blunt) for removing tissue may also be required. Some instruments are insulated and can be attached to electrosurgical generators to facilitate cutting or coagulation of tissue. A complete description of all instruments available is beyond the scope of this chapter. However, be aware that there may be a specific instrument available for almost any need (Figures 4 and 5). Specific training and experience is required to use the more-advanced instruments and techniques safely. There is ample documentation to suggest that, in the case of human medicine, most surgical complications
0839_frame_C27 Page 629 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
629
FIGURE 4 Disposable laparoscopic instruments, all insulated for electrosurgery. (Top) scissors; (Middle) irrigation, suction, and electrosurgery probe; (Bottom) Babcock forceps. (Image credit: S. Dover.)
FIGURE 5 Nondisposable instruments. (Top to Bottom) Tissue forceps, biopsy forceps, insulated scissors, insulated tissue forceps. (Image credit: S. Dover.)
occur early in the learning curve. One report that evaluated predictors of laparoscopic complications after formal training found that at 3 months after training, surgeons who performed clinical procedures without additional training were 3.39 times more likely to have at least one complication compared with surgeons who sought additional training. Additionally, an ongoing clinical association with surgeons performing similar procedures decreases long-term complication rates (See et al., 1993).
Cameras The handling characteristics of video endoscopes are very similar to fiber-optic scopes; yet video endoscopes have the advantage of a camera as an integral part of the system. Video cameras allow the images to be displayed on a video monitor allowing the endoscopist to work more comfortably, as well as allowing the information to be shared with observers or recorded for documentation. The basic endoscopic video camera system consists of the endoscopic adapter, camera head, camera control unit, and monitor. The adapter attaches directly to the eyepiece of the scope where it focuses and magnifies the image. The amount of magnification is dependent on the focal length
0839_frame_C27 Page 630 Tuesday, May 22, 2001 11:55 AM
630
CRC Handbook of Marine Mammal Medicine
of the adapter; the higher the focal length, the higher the magnification. The camera head contains the computer chip that transforms the optical image into an electronic signal, which is processed by the camera control unit and displayed on the monitor. These cameras are made using a solid-state silicone computer chip, consisting of either a single chip or three chips. The chip contains an array of light-sensitive silicon elements. Silicon emits an electrical charge when exposed to light; these charges are then amplified, transmitted, displayed, and recorded. The three-chip camera, in addition to its higher price, has better resolution and color than the single-chip cameras. This occurs because each silicon element contributes one unit (labeled a pixel) and three-chip cameras contain more silicon elements. The videoendoscope uses wires rather than fiber optics to transmit the signal to the camera control unit. Most videoendoscopes have the light source and camera control unit in one system, whereas fiber-optic systems usually have a separate light source and camera control unit.
Video Monitors and Recorders The monitor is the last electronic factor affecting the quality of the image ultimately seen by the operator. Recall that the light source, the endoscope, the camera head, and the camera control unit affect the image quality, all before it reaches the monitor. Therefore, make sure that the monitor is at least as good as the weakest link in the video chain. Do not expect a high-quality monitor to render an excellent image when the scope has damaged fibers and a dirty objective lens. Standard consumer-grade video monitors have 350 lines of horizontal resolution. The minimum monitor resolution should be 500 lines for single-chip systems, and 600 lines for threechip cameras. The monitor should also have S-video (Y/C) inputs for single-chip cameras, and RGB inputs for three-chip cameras. S-video formats separate the brightness and the color into two distinct signals, whereas RGB separates to four channels to maximize the advantages of these formats. Composite video-format monitors offer the least detail and lowest price because they combine all the elements required for a clear image into one signal (Chamness, 1999). Video recorders and printers should be required accessories for any endoscopic system. These recorded images are invaluable for communicating information for clinical, scientific, and educational purposes. A variety of videotape formats are available; VHS and 8 mm are acceptable, but inferior in quality to S-VHS (S-video), Betacam, and Hi-8. Digital video recorders provide higher quality than any analog system, but are significantly more expensive. The best-quality still images are photographs taken with a 35-mm camera attached directly to the eyepiece of the endoscope, but this is inconvenient and time-consuming. It also requires a light source with an integrated flash generator. Video printers, image-capture systems, and slide makers can quickly document still photographs of procedures. By matching the resolution of each component of the system, one can achieve the best overall quality images possible.
Clinical Applications in Cetaceans Cetacean Gastroscopy There are a variety of acceptable variations in the techniques for performing an endoscopic examination of the upper gastrointestinal tract of the species of cetaceans commonly kept in zoos or aquaria. Sedation of the animal with diazepam or midazolam may be indicated. However, esophagoscopy and gastroscopy are commonly performed safely without medication. Fasting for at least 6 hours prior to the examination is recommended. Normal gastric emptying time of healthy dolphins is less than 4 hours, and food items observed at 6 or more hours indicate delayed gastric emptying.
0839_frame_C27 Page 631 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
631
Cetaceans not trained for in-water endoscopy are typically restrained in a stretcher or placed on a foam pad. Care must be taken to ensure the safety of both the attendants and the animal. The preferred technique is to place the animal in sternal recumbency on a padded surface. The dolphin’s head is restrained by placing attendants on either side of the head. Prior to introducing the scope, clean terry cloth bath towels are placed around the maxillary and mandibular rostrum. The upper (maxillary) towel should be placed first because the mandibles are delicate and may be fractured if the animal thrashes with no restraint of the maxilla in place. Never restrain the mandible without first controlling the maxilla. Two towels each on the maxilla and mandible may be needed in larger animals. An additional towel wrapped around the hand introducing the scope is often useful as protection from the sharp teeth of small cetaceans. It is useful to orient the scope image prior to passing into the pharynx, although the internal anatomy of the gastrointestinal tract is less confusing than the respiratory tract. It is valuable to have one person passing the scope and another “driving” the distal optics at the scope controls. Oral speculums made from formed PVC tubes can be used, although many practitioners discourage their use. This rigid speculum should only be used if it is heavily padded to prevent damage to the oral mucosa of the patient. Larger cetaceans will require appropriately matched equipment for keeping the mouth open for examination. The authors have used molded rigid foam to make custom-fitting speculums that are gentle on the teeth and oral mucosa. Some clinicians prefer the use of large-animal endotracheal tubes as speculums. The endoscopist is positioned at the head of the animal (Figure 6), holding the control housing in the dominant hand. The opposite hand advances the insertion tube, or as mentioned above, an assistant is used to advance the tube. The distal scope tip is passed over the dolphin’s tongue and into the oropharynx. As the scope enters the esophagus, air insufflation will be required to obtain an image. With the scope in place, the most efficient moves in order are (1) slow withdrawal; (2) rotation; (3) deflection of the tip; and (4) advancement. Withdrawing the scope is the safest move and increases perspective. This is often enough to orient the endoscopist and direct further movement. Rotation is generally safe, will also increase perspective, and is often required prior to advancing. Indiscriminate advancing of the scope will lead to confusion, misdirection, and can potentially traumatize the patient. The most common cause for an obscured
FIGURE 6 Endoscopist positioned at head of dolphin during gastroscopy. (Image credit: U.S. Navy.)
0839_frame_C27 Page 632 Tuesday, May 22, 2001 11:55 AM
632
CRC Handbook of Marine Mammal Medicine
view occurs when the distal lens is placed directly on the mucosa, which appears as a blurred reddish-pink image. Slowly withdrawing the endoscope tip will correct the problem. The oral cavity and esophagus of cetaceans contain a thick tenacious mucus that will readily adhere to the lens and blur the image; ensure that there is a properly functioning water irrigation valve present to prevent or correct this problem. As the scope is advanced into the oropharynx, numerous punctate openings are visible in the mucosa. These are the openings of mucous glands, which lubricate food items prior to swallowing. Within the oropharynx, the larynx can often be seen, effectively bifurcating the proximal esophagus (Color Figure 2A).* The scope is advanced on either side of this bifurcation into the esophagus, insufflating as necessary. The normal cetacean esophagus is seen as longitudinal folds of mucous membrane to the level of the lower esophageal sphincter. A cardiac pulse is often observed with the scope at midthorax level. The lower esophageal sphincter of cetaceans is not as pronounced as in pinnipeds, and there is not a clear delineation between the mucosa of the forestomach and esophagus (Color Figure 2B). The bottlenose dolphin, like most small odontocetes, has a three-chambered stomach (see Chapter 9, Anatomy). The first chamber (forestomach) has thick squamous epithelium-lined rugal folds that often appear lighter in color than the esophagus and are easily distinguished by their appearance (Color Figure 2C). The second chamber is the fundic portion of the stomach, which is glandular and has a mucosal surface that is deep pink to red in color. The ostium between the forestomach and the second chamber is located cranially in the left ventral quadrant of the first compartment and is approximately 2.5 cm in diameter in a mature bottlenose dolphin (Color Figure 2D). The third chamber is the pyloric region with a thin mucosal lining and mucous glands. A small tubular structure, the connecting channel, connects the fundic and pyloric compartments. Once in the lumen of the forestomach, a prominent fold is seen along the right wall; the lumen of the forestomach lies to the left side of the animal (on the right side of the properly oriented image). By advancing the scope to the fundus and retroflexing the distal tip, the entire lumen of the forestomach can be examined. A moderate amount of cloudy, greenish-brown fluid is usually present in the forestomach (see Color Figure 2C). The gross appearance and cytology of a sample of this fluid may aid in diagnosis. Aspirating this fluid or carefully repositioning the animal may be required to perform a complete examination. It is also often not unusual in clinically normal animals, to observe a waxy material floating on the fluid surface and coating the mucosa. The peristaltic motion of the forestomach is unique in cetaceans examined to date. A primary wave moves from the lower esophageal sphincter to the fundus and a secondary wave proceeds from the fundus to the lower esophageal sphincter. In those dolphins examined thus far, a rate of three to four cycles per 1-min period has been recorded. Reduced peristalsis is frequently seen along with other indications of digestive disturbance (e.g., delayed gastric emptying, persistence of bones, elevated forestomach pH). Hypermotility has been observed with forestomach impactions and the presence of foreign materials. During the primary wave, gastric fluid from the glandular (second) stomach refluxes into the forestomach through the opening just distal to the lower esophageal sphincter on the left inferior aspect of the forestomach. With practice, this reflux and the opening can be visualized by slow withdrawal of the scope with the scope tip retroflexed. The glandular (second) stomach is nondistensible and the mucosa is organized in a distinct arrangement of roughly circular crypts. The mucosa is seen as a velvety appearing deep red color (see Color Figure 2D). A complete evaluation of the lumen and mucosa is not possible, due to the unique anatomy.
*Color Figures follow page 462.
0839_frame_C27 Page 633 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
633
FIGURE 7 Graphic of forestomach and fundic stomach of Atlantic bottlenose dolphin. (Image credit: U.S. Navy.)
The connecting channel between the second and third (pyloric) stomach compartments is intramural, small diameter, and J-shaped. Therefore, it is not possible to examine the remainder of the stomach compartments or proximal small bowel of cetaceans with current technologies (Figure 7).
Colonoscopy With an animal in lateral recumbency and liberal lubrication, small-bore endoscopes can be introduced through the anus. Evaluation of the lumen and mucosa of the rectum and colon is therefore possible. The authors recommend using scopes intended for use in fluid media like those for evaluation of the urinary tract of humans, and to distend the bowel with sterile isotonic fluids for these examinations. The mucosa of the lower gastrointestinal tract of cetaceans appears to be more friable than that of terrestrial species. This is postulated to be because marine mammals do not pass solid stools, and have no requirement to store feces in the colon or rectum. As a result, trauma to the mucosa is easily induced unless caution is used during endoscopic procedures. The indications for lower gastrointestinal endoscopy are infrequent, and the extent of the evaluation, i.e., how far the scope is advanced into the bowel, must be weighed against the potential for trauma. The farther the scope is advanced, the more difficult control becomes, and the more potential exists for difficulty visualizing tissue or inducing trauma. As the scope is introduced through the anus and advanced proximally, the lining is observed as a continuation of the skin for several centimeters. An obvious distinct demarcation is then seen as the mucosa changes to typical velvety red tissue (Color Figure 2E). Samples for cytology, microbiology, or histopathology are easily obtained through the working channel of the scope.
Respiratory Endoscopy Pharyngoscopy, tracheoscopy, and bronchoscopy can be performed successfully in conscious cetaceans. Some small cetaceans have been trained to accept the bronchoscope while still stationed in the water. However, these procedures are most commonly performed with the animal out of the water using appropriate levels of physical restraint with or without sedation. Dolphins undergoing nasal, pharyngeal, or lower airway endoscopy are usually restrained in
0839_frame_C27 Page 634 Tuesday, May 22, 2001 11:55 AM
634
CRC Handbook of Marine Mammal Medicine
FIGURE 8 Endoscopist positioned during bronchoscopic examination of Atlantic bottlenose dolphin. (Image credit: U.S. Navy.)
sternal recumbency. Handlers positioned alongside the animal may be the only restraint required, although sedation should be used if necessary. The endoscope should be cleaned and disinfected prior to and after all uses. If introduction of the scope through the glottis is planned or expected, the scope should be sterilized and the procedure performed aseptically. It is critical to orient the image produced by the equipment with the animal’s position prior to introducing the distal tip into the nasal passage. Once beyond the blowhole plug, orientation becomes very difficult, especially for the endoscopist beginner. Initial introduction of the scope is accomplished by grasping the distal few centimeters with the sterile gloved dominant hand and bracing it on the animal’s melon (Figure 8). During a breath, the scope is quickly inserted into the airway and held still, until the animal accepts it without objection. Dolphins will often produce a series of short, forceful exhalations when the scope is first introduced. The introduction of lidocaine via the operating channel may help alleviate discomfort (Harrell et al., 1996; Van Bonn et al., 1997), although the authors have completed numerous examinations without its use. Again, with the scope in place, the most efficient moves in order are (1) slow withdrawal of the scope; (2) rotation; (3) deflection of the tip; and (4) advancement. With the scope inserted a depth of 10 to 15 cm from the blowhole surface, the nasopharynx is visualized. Along the lateral wall of the nasopharynx, just inferior to the nasal septum, the opening of the auditory (Eustachian) tube is often visible. The laryngeal cartilages are visualized within the depths of the nasopharynx (Color Figure 2F). The palato-pharyngeal muscular complex is dynamic and is often seen closing over the laryngeal cartilages obscuring them from view. In a clinically normal animal, a moderate amount of white froth is commonly seen in the dependent aspects of the nasopharynx, and may also obscure anatomy. Introduction through the glottis requires
0839_frame_C27 Page 635 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
635
positioning the scope tip just superior to the laryngeal cartilages and careful timing to pass the tip into the open glottis during a breath. In the authors’ experience with Atlantic bottlenose dolphins, scopes with insertion tube diameters of 8 to 9 mm can be advanced to a depth of approximately 70 to 80 cm (sixthgeneration bronchi), and scopes with an insertion tube diameter of 3 mm can be advanced to a depth of approximately 90 to 110 cm (Color Figure 2G). Bronchoalveolar (BAL) washes obtained from each of these sites in clinical cases have returned markedly different cytology and microbiology results (see Chapter 19, Clinical Pathology; Chapter 20, Cytology). Standardization of this technique is ongoing. A data set of cytological, microbiological, and chemical analyses (immunoglobulin and cytokine concentrations) (see Chapter 13, Immunology) from clinically normal dolphins under controlled conditions must be obtained before valid clinical interpretations of BAL in dolphins can be made. Standardization of the anatomical nomenclature of cetacean airways is also yet to be common practice, but will facilitate communication between facilities. The nomenclature system applied to dogs and horses has been applied to the bronchial tree of the harbor porpoise (Phocoena phocoena), and there are rather elaborate descriptions of the nasal structures of odontocetes (Cranford et al., 1996; Harper et al., 1998).
Urogenital Retrograde cystoscopy of the urinary bladder has been performed in female dolphins, and at least one animal has been trained to allow this procedure while stationed in the water. This procedure requires a small-bore, sterile scope, and must be performed with scopes designed for use with fluid distending media. The urethral orifice of female dolphins is located at the apex of the clitoris. The entire genital slit should be cleansed with an antiseptic prior to scope introduction. Sterile technique must be used when handling the scope during cystoscopy. To the authors’ knowledge, urethroscopy or cystoscopy of male cetaceans or pinnipeds has not been reported. It is likely that by passing a small-bore endoscope retrograde within the urethra, as one would a urinary catheter, this procedure may be performed. The lumen of the uterus of female dolphins can be examined with sterile small-bore scopes. This must be performed as a sterile procedure with preparation of the genital slit as for cystoscopy. In addition, flushing the entire vaginal vault with an antiseptic wash to remove sources of contamination that may be introduced through the cervix should be included in patient preparation. With the animal in lateral recumbency, a sterile speculum is placed into the vagina and advanced to the level of the pseudocervix. The scope is then inserted through the speculum, the true cervix visualized, and the scope advanced into the lumen of the body of the uterus. Distention of the uterus with sterile isotonic fluid and use of scopes designed for fluid media are recommended. Indications for hysteroscopy are not frequent and the above approach can be very difficult in animals that are not postpartum. However, as controlled breeding of small cetacean species becomes more critical to maintaining populations under human care, this may prove to be a valuable part of examinations for breeding soundness . The technologies and techniques developed may also find application to endangered species management (Robeck et al., 1994; Robeck, 2000).
Clinical Applications in Other Marine Mammals The largest difference in the technique of flexible endoscopy as applied to other marine mammal species is the requirement for general anesthesia. With the exception of the pharynx and larynx, the endoscopic anatomy of seals and sea lions is very similar to that of domestic dogs, and comparable techniques are used (Color Figure 2H); see Jones’s Veterinary Endoscopy (1990) for
0839_frame_C27 Page 636 Tuesday, May 22, 2001 11:55 AM
636
CRC Handbook of Marine Mammal Medicine
FIGURE 9 Positioning of sea lion for endoscopic examination. (Image credit: U.S. Navy.)
a complete description of suitable small animal techniques. The pinniped pharynx has an abundance of pendulous soft tissue that will often obscure visualization of the laryngeal cartilages. The epiglottis is also rather abbreviated when compared with dogs, and appears less prominent. Full extension of the head and neck during scope introduction will facilitate efficient examinations (Figure 9). Endonasal pharyngoscopy has been described as useful in facilitating tracheal intubation in the Florida manatee (Walsh et al., 1997).
Minimally Invasive Surgical Techniques There are several excellent references describing techniques for MIS. However, these procedures should not be performed without adequate training and experience (Cook and Stoloff, 1999; Freeman, 1999; Tams, 1999). The basic techniques for most marine mammals are similar in principle to terrestrial species. The primary differences pertain to anatomical limitations and postoperative exposure to an aquatic environment. This section describes the approach for MIS in general terms, followed by individual species considerations. As techniques evolve, some of the procedures described here will eventually become obsolete. Regardless of the species, the approach for MIS depends on the organ of interest. The most common procedure performed in marine mammals is laparoscopic exploratory examination and subsequent biopsy collection. With experience and increasing numbers of cases, more advanced procedures will be developed for the indications above. The basic procedures in MIS are the same as for open surgery: access, exploration, retraction, dissection, hemostasis, tissue apposition, tissue removal, and closure (Kolata and Freeman, 1999).
Insufflation As in flexible endoscopy, a visual space is needed for effective examination. In laparoscopy, carbon dioxide is the preferred insufflation gas. Carbon dioxide is inexpensive, readily available, nonexplosive, physiologically absorbed, and easily excreted. Nitrous oxide, air, and helium have also been used, but they all have limitations. Insufflation is performed by an automatic insufflator (Figure 10), which controls the flow rate and maintains a constant intra-abdominal pressure. In most species, a working pressure of 12 to 14 mmHg is sufficient. It is recommended
0839_frame_C27 Page 637 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
637
FIGURE 10 Mechanical insufflator, gauges measure gas bottle pressure, insufflation line pressure, flow rate, and intra-abdominal pressure. (Image credit: S. Dover.)
that the maximum pressure not exceed 15 mmHg, as serious respiratory and cardiovascular compromise can occur. In thoracoscopic procedures, the rigidity of the chest wall and the ability to collapse one lung allow the creation of a working space. Insufflation can be used in the thorax. Yet, care should be exercised to avoid iatrogenic damage. MIS requires additional equipment and instrumentation, dependent on the procedure to be performed. Insufflation needles, trocars, forceps, scissors, dissectors, staplers, clip appliers, suture, retrieval pouches, and electrosurgical and irrigation/suction probes may be needed. Most of these instruments are available in disposable and nondisposable varieties, and there are a number of suppliers. As in conventional open surgery, a clear surgical field is essential. The use of hemostasis, irrigation, and suction may be required, and should be prepared for use prior to starting the procedure. Management of hemorrhage can be difficult in MIS, and even minor bleeding can quickly obscure the operative field. Electrosurgical generators, lasers, and ultrasonic generators can all effectively provide hemostasis. The most common technique is the use of conventional electrosurgical devices that are standard equipment in most practices. Many instruments are designed to attach directly to the electrosurgical unit, allowing a variety of hemostatic methods. Lasers and ultrasonic generators are less commonly used in veterinary medicine, although there are indications for their usage. These instruments should not be used without further training or experience. Preliminary investigations involving laparoscopic procedures in marine mammals, especially cetaceans, suggest that this technology can expand current diagnostic and treatment capabilities. Advances in anesthetic techniques will increase the safety of these procedures, as general anesthesia is usually indicated. It may prove possible to perform limited procedures under heavy sedation or regional anesthesia, but to date this has not been attempted.
Access Access to the internal structures is achieved by placing ports (trocars and cannulas) to allow introduction of telescopes and instruments to perform the desired procedures. One must create an optical space to allow visualization of the organs and have sufficient space to work.
0839_frame_C27 Page 638 Tuesday, May 22, 2001 11:55 AM
638
CRC Handbook of Marine Mammal Medicine
FIGURE 11 (Left to right)Disposable Veress needle, 5-mm trocar with blade shield, 12-mm trocar with blade shield, 11-mm optical surgical obturator and sleeve. (Image credit: S. Dover.)
The techniques used are dependent on the anatomy of the patient and the discretion of the surgeon. The two most common approaches are the closed technique or the Hasson (open) technique. Both procedures are designed to establish a pneumoperitoneum, and, as with many surgical procedures, it is necessary to become familiar, and then competent, with a variety of techniques. In the closed technique, insufflation is achieved by introducing a Veress needle® (Ethicon, Inc., Somerville, NJ) into the peritoneal cavity. The Veress needle contains a spring-loaded blunt obturator that extends past the sharp tip once it has penetrated the peritoneum, thereby protecting the underlying organs (Figure 11). The gas supply line is attached to the stopcock on the needle to create the pneumoperitoneum. The Veress needle is replaced with the trocar sized for the telescope required. The Hasson technique was developed by Harrith Hasson in 1974 to avoid injury to the intra-abdominal organs encountered in the closed (blind) method. Hasson designed a cone with flanges for attaching sutures, which fits over the cannula (Kolata and Freeman, 1999). A small incision is made that extends to the peritoneum, which allows direct visualization of the organs. A blunt trocar is then inserted. Sutures are placed and tightened to create a gas-tight seal between the fascia and the flanges on the trocar. The gas supply line is attached to a stopcock on the trocar to create the pneumoperitoneum.
Trocars and Cannulas Trocars and cannulas are available in a variety of lengths and diameters, with either blunt or sharp tips (Figure 12). Some disposable trocars are made with safety shields that guard the blade after it has penetrated the peritoneum (see Figure 11). The placement of the primary port is the most critical part of most MIS procedures. If the Hasson approach is used, that cannula becomes the primary port for telescope placement. If the closed technique is used, a shielded trocar replaces the Veress needle as the primary port. The replacement of the Veress
0839_frame_C27 Page 639 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
639
FIGURE 12 Reusable trocars, 5- and 10-mm diameters, various lengths. (Image credit: S. Dover.)
needle with a trocar must be performed carefully. If the abdomen is not sufficiently inflated, the body wall will be depressed during trocar insertion, allowing the tip of the trocar to come dangerously close to the viscera (Kolata and Freeman, 1999). After the trocar and cannula penetrate the abdomen, the trocar is removed and replaced with an appropriately sized telescope. Be sure to examine the abdominal contents for indications of iatrogenic trauma from trocar placement. Secondary trocars are inserted in locations that will provide access to the organ of interest, and will allow instruments space to perform the surgical task. These ports need to be carefully planned to create angles that allow instrument manipulation without interference. Typically, these instruments are placed from 30 to 60° apart, and must be neither too close, nor too far, from the organ being manipulated. These trocars are placed under direct visualization from the telescope, allowing the surgeon to avoid severing blood vessels or organ damage. Careful planning is necessary to complete the desired procedure efficiently. Failure to plan adequately can significantly prolong the procedure, thereby adding to potential complications from additional anesthesia.
Closure After completion of the procedure, the instruments and cannulas are removed, and the surgical wounds are closed. The closure required is dependent on the species of the patient and the size of incisions created. In humans, only the skin is closed for incisions of 5 mm or less, whereas larger defects require closure of the peritoneal and fascial layers. These guidelines have been successfully followed in marine mammals. Consideration for the requirement of replacing the
0839_frame_C27 Page 640 Tuesday, May 22, 2001 11:55 AM
640
CRC Handbook of Marine Mammal Medicine
animals back in an aquatic environment should be allowed. Methylmethacrylate surgical glue is routinely used for the subcutaneous and skin layers, to assist in creating a watertight closure.
Minimally Invasive Surgery in Cetaceans The laparoscopic approach for the bottlenose dolphin requires general anesthesia. The animal is placed in lateral recumbency with the abdomen prepared for sterile surgery. Insufflation is achieved by introducing a Veress needle into the peritoneal cavity lateral to the rectus abdominus muscle at the level of the umbilicus. Approximately 3 l of sterile carbon dioxide are used to inflate the peritoneal cavity to a maximum internal pressure of 15 mmHg. For minor procedures small-diameter (5 mm or less) trocars can be used. However, larger trocars (up to 12 mm) will allow insertion of 10-mm telescopes that will provide examination of the entire abdomen. Secondary ports are placed as described above to allow manipulation and sampling of organs (Color Figure 3A). The peritoneum of many cetaceans is highly elastic and distensible (Color Figure 3B), and can be difficult to penetrate. When this occurs, it is advisable to use a blunt instrument to penetrate the peritoneum, to avoid inadvertant damage to underlying organs. The laparoscopic view of an Atlantic bottlenose dolphin with the telescope placed laterally as above allows examination of most of the abdominal organs (Color Figure 3C and D). Color Figures 3E through G illustrate the steps required to perform a renal biopsy. The renal capsule is examined for an avascular region for access (Color Figure 3E). The renal capsule is opened with blunt scissors and a single reniculus is dissected free (Color Figure 3F). A pre-tied loop of absorbable suture is placed around the pedicle of the reniculus (Color Figure 3G) to provide hemostasis, and the top of the reniculus is removed for biopsy (Color Figure 3H). The renal capsule is not closed, as the wound is minor and should heal without complications. The incisions created to accommodate the trocars are closed using large (#0 to #1 PDS) absorbable suture, followed by closing the skin with nonabsorbable suture and surgical glue (Dover et al., 1999). Although thoracoscopy has not been attempted in small cetaceans as yet, the techniques have been developed in cadavers, and should be feasible in anesthetized animals. The future of these techniques in cetaceans requires careful planning to minimize complications and optimize success. All clinicians are required to adapt and develop new procedures to further the care of their patients.
Minimally Invasive Surgery in Other Marine Mammals Procedures for MIS in pinnipeds, sea otters, and polar bears (Ursus maritimus) are not significantly different from techniques used in terrestrial species. Careful attention to the anatomical differences among the species will prevent complications and increase the opportunity for success. The Florida manatee presents additional anatomical and logistical challenges due to their unique anatomy. Both laparoscopic and thoracoscopic procedures have been successfully performed in manatees for diagnostic and treatment purposes. The thick dermis can be challenging to penetrate, and the heavy organs can be difficult to manipulate. Despite these obstacles, a variety of procedures can be accomplished successfully. Color Figures 4A and B show normal views of the abdomen and lung of the Florida manatee. Severe lung pathology is a common finding in Florida manatees, often associated with trauma. Color Figure 4C demonstrates severe pleuritis, with fibrinous adhesions. This view was obtained after draining several liters of serosanguinous fluid from the thoracic cavity. Color Figure 4D illustrates pneumothorax, pneumonia, and plueritis, all secondary to severe trauma.
0839_frame_C27 Page 641 Tuesday, May 22, 2001 11:55 AM
Flexible and Rigid Endoscopy in Marine Mammals
641
Acknowledgments The authors thank the many individuals who have participated or contributed to this work. Special thanks to their colleagues from SeaWorld, Inc., the U.S. Navy Marine Mammal Program, and other practitioners that have assisted in developing these techniques over many years of work. Jim McBain is thanked for reviewing this chapter. Ronald Kolata provided many hours of training and assistance in the development of MIS in marine mammals. Mark Todd provided invaluable assistance in image processing.
References Applegate, W.I., Jensen, E.D., Van Bonn, W.G., Ketzenberger, B.K., and Fradkin, J.F., 2000, Endoscopy assisted retrieval tools: creative solutions to challenging cases, in Proceedings of the Joint Meeting of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine, New Orleans, LA, 444. Beroza, G.A., and Barclay, W.P., 1981, Manual retrieval of a gastric foreign body in an Atlantic bottlenose dolphin, J. Am. Vet. Med. Assoc., 179: 1286–1288. Chamness, C.J., 1999, Endoscopic instrumentation, in Small Animal Endoscopy, 2nd ed., Tams, T.R. (Ed.), Mosby, St. Louis, MO, 1–15. Cook, R.A., and Stoloff, D.R., 1999, The application of minimally invasive surgery for the diagnosis and treatment of captive wildlife, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 30–40. Cranford, T.W., Amundin, M., and Norris, K., 1996, Functional morphology and homology in the odontocete nasal complex: Implications for sound generation, J. Morphol., 228: 223–285. Dover, S.R., Beusse, D.O., Walsh, M.T., McBain, J.F., and Ridgway, S., 1999, Laparoscopic techniques for the bottlenose dolphin (Tursiops truncatus), in Proceedings of the 30th International Association for Aquatic Animals Medicine, Boston, MA, May, 128–129. Freeman, L.J. (Ed.), 1999, Veterinary Endosurgery, Mosby, St. Louis, MO, 1–23. Harper, C., Borkowski, R., and Hoffman, A., 1998, Development of a standardized nomenclature for bronchoscopy of the respiratory system of harbor porpoises (Phocoena phocoena), in Proceedings of the 29th International Association for Aquatic Animals Medicine, San Diego, CA, May, 9–10. Harrell, J. H., Reidarson, T.H., McBain, J.F., and Sheetz, H., 1996, Bronchoscopy of the bottlenose dolphin (Tursiops truncatus), in Proceedings of the 27th International Association for Aquatic Animal Medicine, Chattanooga, TN, May, 33. Hawkins, E.C., Townsend, F.I., Lewbart, G.A., Stamper, M.A., Thayer, V.A., and Rhinehart, H.L., 1997, Bronchoalveolar lavage in a dolphin, J. Am. Vet. Med. Assoc., 211: 901–904. Jones, B.D. (Ed.), 1990, Veterinary Endoscopy, Veterinary Clinics of North America: Small Animal Practice, 20: 1199–1395. Kolata, R.J., and Freeman, L.J., 1999, Access, port placement and basic endosurgical skills, in Veterinary Endosurgery, Freeman, L.J. (Ed.), Mosby, St. Louis, MO, 44. Lunemann, H.J., 1999, Endoscopic documentation, in Small Animal Endoscopy, 2nd ed., Tams, T.R. (Ed.), Mosby, St. Louis, MO, 17–24. Murray, M.J., Schildger, B., and Taylor, M., 1998, in Endoscopy in Birds, Reptiles, Amphibians and Fish, Endo-press, Tuttlingen, Germany, 86. Pieterse, C.M., Landau, R., and van der Elst, C., 1998, Successful retrieval of some unusual foreign bodies from the forestomach of a sedated Tursiops truncatus with the aid of gastroscopes and modified grasping devices, in Proceedings of the 29th International Association for Aquatic Animal Medicine, San Diego, CA, May, 45. Quandt, J.E., 1999, Anesthetic considerations for laser, laparoscopy, and thoracoscopy procedures, Clin. Tech. Small Anim. Pract., 14: 50–55.
0839_frame_C27 Page 642 Tuesday, May 22, 2001 11:55 AM
642
CRC Handbook of Marine Mammal Medicine
Robeck, T.R., 2000, Advances in the understanding and manipulation of bottlenose dolphin reproduction, in Proceedings Bottlenose Dolphin Reproduction Workshop, San Diego, CA, June, 1999, 109–131. Robeck, T.R., Curry, B.E., McBain, J.F., and Kraemer, D.C., 1994, Reproductive biology of the bottlenose dolphin (Tursiops truncatus) and the potential application of advanced reproductive technologies, J. Zoo Wildl. Med., 25: 321–336. See, W.A., Cooper, C.S., and Fisher, R.J., 1993, Predictors of laparoscopic complications after formal training in laparoscopic surgery, J. Am. Med. Assoc., 270: 2689–2692. Stetter, M., Mangold, B., Miller, M., Webber, M., and Capobianco, J., 1999, Successful treatment of lead toxicosis in a bottlenose dolphin (Tursiops truncatus), in Proceedings of the 30th International Association for Aquatic Animal Medicine, Boston, MA, May, 146–147. Sweeney, J.C., 1990, Surgery, in The Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 227. Tams, T.R. (Ed.), 1999, Small Animal Endoscopy, 2nd ed., C.V. Mosby, St. Louis, MO, 510 pp. Van Bonn, W., Cranford, T., Chaplin, M., Carder, D., and Ridgway, S., 1997, Clinical observations during dynamic endoscopy of the cetacean upper respiratory tract, in Proceedings of the 28th International Association for Aquatic Animal Medicine, Harderwijk, the Netherlands, May, 38–43. Walsh, M.T., Webb, A., Bailey, J., and Campell, T.W., 1997, Sedation and anesthesia of the Florida manatee (Trichechus manatus), in Proceedings of the 30th International Association for Aquatic Animal Medicine, Harderwijk, the Netherlands, May, 10–11.
0839_frame_28fm Page 643 Tuesday, May 22, 2001 11:56 AM
28 Thermal Imaging of Marine Mammals Michael T. Walsh and Edward V. Gaynor
Introduction Thermography is the production of a visual image of the surface temperature of a live subject or an inert object. It has been used in the medical field for detecting differences in temperature of soft tissue that are undetectable to the human eye or touch. It also has law enforcement, criminal tracking, surveillance, preventative maintenance of electrical equipment, building diagnostics, energy audits, and numerous other nonmedical applications. Injured tissues experience altered circulation and increased heat, and this focal inflammation can often be detected and measured with heat-sensitive equipment. The thermal changes can then be converted into a multicolored temperature map showing differences as small as 0.05°C. Alternatively, the technique can indicate areas where circulation is decreased, such as a blood clot, scar tissue, or necrotic tissue.
Technique A visual image is dependent on a light source, such as the sun reflecting off an object, which is perceived by the eye. Visual light is just one small portion of the electromagnetic spectrum, and the human eye can only see the narrow central band of visual light that one thinks of as the colors of a rainbow. The various types of electromagnetic radiation in order of decreasing wavelength and increasing frequency are radiowaves, microwaves, infrared radiation (IR), visible light, ultraviolet radiation, X-rays, and gamma radiation. Warmer objects radiate more energy at all wavelengths, and the peak energy shifts wavelengths as the temperature increases. For example, at 600°C (1112°F) a red glow is perceived in a heated object, but at 2000°C (3632°F), such as an incandescent lamp, yellow is perceived. Normal temperature (ambient room temperature) objects do not show a visible radiation change, but they do radiate changes in the IR region of the spectrum that show up as changes in color. Within this portion of the spectrum, there are two important IR bands: mid-wave IR (3 to 5 µ m) and long-wave IR (8 to 12 µ m). For thermography purposes, the long range is generally preferred, because it is less affected by sunlight. IR, like visible light, can be focused with lenses and detected by photodetectors tuned for the IR. Lenses are made from crystalline minerals such as germanium, and some zinc salts, since glass is opaque in the IR.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
643
0839_frame_28fm Page 644 Tuesday, May 22, 2001 11:56 AM
644
CRC Handbook of Marine Mammal Medicine
History The first use of IR technology occurred in the military arena as early as the 1940s with the development of heat-seeking missiles. In the 1960s and 1970s, tactical IR night vision equipment was developed that depended on cryogenic cooling with various gases or liquids such as nitrogen. More refinements took place in the 1980s, spearheaded by the military, with only peripheral involvement of the medical field. In the 1990s, the commercial field accelerated with the advent of microchip technology. Machines the size of camcorders became available without the need for gas or liquid cooling. This portability, and the recognition that this technique can be used in broad daylight or in darkness for many applications, has made the technology more appealing for both the human and veterinary fields (Jones, 1998). It was the human field that first investigated the potential for IR in medicine with numerous works from all over the globe in the 1960s (Di Blasio, 1966). Early work suggested its use in helping diagnose breast carcinoma (Isard et al., 1969) and investigating many other areas, such as perfusion of tissues (Love, 1980), but it slowly fell out of favor, because of its reliance on first-generation cameras. Numerous investigators continued to move the field forward (Goodman et al., 1985), and there were calls for studies to validate its diagnostic efficiency (Frymoyer and Haugh, 1986). Innovations resulted in thermography even being adapted for use with laparoscopic procedures (Roberts et al., 1997). In 1998, a reappraisal of the use of IR in medicine showed its successful use in neurology, vascular disorders, rheumatic diseases, tissue viability, oncology (especially breast cancer), dermatology disorders, neonatology, ophthalmology, and surgery. The most common use of veterinary thermography to date has been in the equine field (Stromberg, 1974; Purohit and McCoy, 1980; Turner et al., 1986). Hoof conditions including laminitis, navicular disease, abscesses, and corns can be identified (Turner, 1991). Ligament and joint capsule injury can also be visualized as hot spots. Tendonitis can be detected as early as 2 weeks prior to any physical swelling or pain, as can arthritis or osteitis (Vaden et al., 1980). Back disease in horses can also be visualized, but its detection is based on vasoconstriction at the affected site that is noninflammatory in nature (Graf von Schweinitz, 1999). Thermography can also detect muscle anomalies resulting from injury or atrophy (Turner, 1989). With partial acceptance in equine medicine, it was only a matter of time before this exposure resulted in trials in other species, such as cattle (Cockcroft et al., 2000), rats, and gerbils (Wu et al., 1997). As with other more-established technologies, it was also time to investigate the diagnostic validity of thermography with controlled studies. Weil et al. (1998) compared 36 normal horses with 119 lame individuals, and concluded that thermography can show and quantitatively prove changes in skin temperature in forelimb lameness. They also emphasized that thermography is most useful in combination with a thorough clinical examination, including other diagnostic examination procedures. It was the veterinary field that prompted use of thermography in marine mammals in the mid-1980s. If a major application of this technology was for examination of the legs of horses, how does that make it useful to the clinician who takes care of species without traditional limbs, such as cetaceans or manatees? In reality, there are a number of applications that are possible in marine mammals. A thermography unit expands the idea of body temperature evaluation beyond the use of a rectal thermometer, usually considered one of the most basic tools. Now a clinician can detect many conditions before they become a source of systemic illness. Detection of infection or inflammation while still a localized problem can result in more accurate diagnosis, monitoring, and more effective treatment. Wound evaluation and resolution of skin conditions assume a very different approach; some conditions that do not show up in bloodwork evaluation can be evaluated by thermography.
0839_frame_28fm Page 645 Wednesday, May 23, 2001 10:53 AM
Thermal Imaging of Marine Mammals
645
Thermography opens visual windows, just as radiography, and provides information on body changes, just as bloodwork. The limitations are related more to lack of imagination and lack of familiarity with this complex tool.
Cameras Use of thermography techniques was previously dependent on purchase or lease of a fairly complicated piece of equipment that was gas cooled. It employed high-pressure miniature gas cylinders that required filling by a special pump, or another larger, high-pressure tank with a special step-down valve. Preparation was time-consuming and movement of the equipment required at least two trained people, and an extension cord tied to a power source. The imager portion, held by the examiner, was tied to a processor by an umbilical cord, and the processor was connected to a monitor. Pointing the camera section while attempting to observe the image on the monitor and adjusting settings with a handheld remote was more than many potential users were willing to endure. In addition, sunlight made observation of the monitor very difficult. A virtual-reality headset was added to allow the examiner to view the image without turning away from the patient, and to remove the interference of sunlight. It also allowed the examiner to look below the glasses and visualize the position of the patient. Still, the process was cumbersome and not geared to the technically faint of heart. Technological improvements in the 1990s led to the development of noncooled machines reduced in size to that of a portable handheld camera. The following discussion is based on the EmergeVision (eMERGE Interactive, Inc., Sebastian, FL) thermographic camera system. The size of a camcorder, it has a flip-out LCD screen for viewing, and can be fitted with a virtual-reality headset. Although there are other systems available, this camera was originally chosen by one of the authors (M.T.W.) because of its low relative cost and its ease of operation. The operator can choose from a number of color palettes with which to view the subject, record the name, and save the images to a small 8-megabyte disk that can be downloaded to a computer. The images can be transferred for further processing, storage, and included in a workable clinical database for future reference and comparison. Real-time temperature measurements are not available with this model, although the images can be downloaded and measurements made.
Clinical Applications As with any diagnostic technique, there are certain guidelines that should be followed to enhance the quality of images and to avoid complicating artifacts (Walsh et al., 1988). For marine mammals it is important to know that, like glass, water is opaque to IR, so one cannot view the animal beneath the water surface. Initially, the heat pattern of an animal just surfaced or removed from the water will be masked until the animal begins to warm. This may take 10 to 30 s or longer depending on the temperature of the water or the area of interest. Areas that are being abraded by swim patterns or by rubbing must not be confused with true pathological conditions. Thus, it is important that the user establish normal body heat patterns before interpreting the images (Kasting et al., 1990). The clinician can choose from a variety of color palettes to portray the image on the screen. The authors like to utilize the black-and-white image to achieve the best detail, using the color choices to highlight areas of concern (Walsh et al., 1993; see Color Figure 5*). Once the image is captured on an 8-megabyte removable card, it can be downloaded into a computer, and the *Color Figures follow p. 462.
0839_frame_28fm Page 646 Tuesday, May 22, 2001 11:56 AM
646
CRC Handbook of Marine Mammal Medicine
image further processed and placed into a patient database. It can be copied for display use or stored as part of the medical record of the patient for future reference or comparison.
Manatees For thermography procedures in manatees (Trichechus manatus), the animal is placed on the pool bottom or raised out of the water by a false-bottom floor. When possible, the procedure is done in the shade to avoid sunlight artifacts. The first use of thermography by one of the authors (M.T.W.) was in 1987 on a manatee that had recurrent drainage from abscesses over the left shoulder. Color Figure 5 (upper left) shows the first attempt to visualize the wound, shown by the large white and yellow site. The smaller dot is the animal’s eye. This was an argon gas system and a color bar to the right gives the approximate temperature relationship. The detail is poor, mainly because of a lack of knowledge about what would be the best technique in this species. Over time, it became apparent that not only could the abscess site be well delineated, but it also could be detected before the area became visually apparent. Although initial drainage would appear to resolve the visual condition, the thermogram showed the area was still very inflamed and demonstrated its tendency to recur. Antibiotic use and the surgical approach were changed, based on the thermogram. A thermogram of a second manatee with a nonresponsive draining tract in the pectoral flipper showed that the infection involved more of the proximal arm tissue than was visually apparent. A second surgical site was opened based on the thermogram, and the wound resolved (Color Figure 5 (upper right)). Manatees often incur massive wounds secondary to boat trauma (see Chapter 3, Manatee Case Study; Chapter 43, Manatees). These are difficult to manage, because of the extent of bone and soft-tissue damage and require constant debridement. Thermography can indicate necrotic or compromised tissue shown by a decrease in temperature. The clinician can also determine if grossly apparent necrotic tissue is still viable. Failure to remove this necrotic tissue before it is covered with granulation tissue has led to chronic fistulous tracts and osteomyelitis. Color Figure 5 (lower left) is a thermogram of a juvenile manatee with cold-water damage, where the tail is sloughing large amounts of necrotic tissue. While visually it appears dead, as outlined by the black line, and in need of extensive debridement, the thermogram shows there is still a good blood supply, and even highlights the scalpel cuts that were made initially to search for evidence of viability.
Pinnipeds Pinnipeds tend to portray a very even external body temperature after leaving the water, so adequate time must be given to detect abnormal areas of heat. Color Figure 5 (center) shows a frontal view of a walrus (Odobenus rosmarus) with the head to the right. Note the symmetry of the face and the numerous skin lesions that are present on the body from tusking by the male during breeding. Color Figure 5 (lower right) illustrates the facial area of a walrus with a chronic infection of the left tusk that was originally thought to be quiescent, but thermographically shows an extensive amount of inflammation, and required further surgical intervention. Inflammation from trauma is illustrated in Figure 1 showing a male California sea lion (Zalophus californianus) that was inappetent. The white areas are the sites of greatest heat. Interpreting black-and-white thermograms takes more visual concentration, but provides much better detail. The thermogram taken from a distance showed a large number of wounds that were inflicted by a more dominant male. With this information, the clinician can monitor the animal’s progress and wound response with or without antibiotics. Figure 2 is a thermogram of a male California sea lion that exhibited a swollen neck, but the caretakers did not see any evidence of wounds. A thermogram of the area pinpointed the bite areas.
0839_frame_28fm Page 647 Tuesday, May 22, 2001 11:56 AM
Thermal Imaging of Marine Mammals
647
FIGURE 1 Thermogram of a male California sea lion. Multiple white areas are secondary to breeding trauma.
FIGURE 2 California sea lion with swelling of the neck area of unknown origin. The thermogram showed bite wounds as the source of the swelling, pointed out by finger, as indicated by the white areas on the neck.
Cetaceans It is with cetaceans that thermography has been most applied. Use in these animals has illustrated the ability of thermography to detect nonvisual areas of inflammation, clarify their extent, and allow the clinician to follow closely the progress of disease and its therapy. Differentiating external trauma from internal disease can be accomplished in some cases with a thermogram. Figure 3 shows areas of heat associated with conspecific trauma in a killer whale (Orcinus orca) that was acting sluggish. Figure 4 illustrates the heat pattern in the upper left maxillary area of a killer whale that loosened two teeth, while reprimanding a conspecific. Figure 5 illustrates the heat pattern in the lower right mandible of a killer whale that developed an area of inflammation secondary to tooth extraction. Resolution of
0839_frame_28fm Page 648 Tuesday, May 22, 2001 11:56 AM
648
CRC Handbook of Marine Mammal Medicine
FIGURE 3 Black-and-white thermogram of an adult female killer whale. The animal is in a slideout area completely dry and the head is off the image to the right. Although the animal appeared visually normal, the thermogram shows obvious heat signatures related to interactions with another female, some associated with rake marks and others not.
FIGURE 4 Open mouth view of a killer whale with thermogram showing heat area in the upper left maxillary region (as outlined by white line) associated with loose teeth. Note that comparing the symmetry of the image helps to clarify its location and extent.
this heat signature may take months, but this would not be obvious with a visual inspection. Oral disease can also be well visualized in smaller cetaceans by thermography. Figure 6 shows the right mandible of a bottlenose dolphin ( Tursiops truncatus), with a hot area that is extending lingually toward the tongue. An image showing a focal area solely around the tooth would be less serious. The heat signature may also extend laterally, as well as lingually. This animal required surgical intervention and the tooth was extracted. Follow-up thermograms showed heat was present for weeks after visual resolution. Thermography is definitely a technique that requires constant use to catch aspects that might otherwise be missed. Figure 7 shows a generalized histiocytic dermatitis in a dolphin that
0839_frame_28fm Page 649 Tuesday, May 22, 2001 11:56 AM
Thermal Imaging of Marine Mammals
649
FIGURE 5 Thermogram of a heat source in the right mandible of a male killer whale after tooth extraction as outlined by white line. The heat extends lingually toward the center.
FIGURE 6 Thermogram of a bottlenose dolphin mandible with an isolated heat source originating from an infected tooth. T = tongue. Note the inflammation extending into the buccal area lateral to the row of teeth.
obviously has a great deal of inflammation present. Biopsy revealed a histiocytic dermatitis, but the animal showed no changes in any bloodwork parameters.
Other Marine Mammal Species Thermography has not been adequately investigated for its clinical applications in most other marine mammal species. Figure 8 shows a thermogram of the right foot of a limping polar bear (Ursus maritimus). There is obvious involvement of the outer digit that could be visualized without handling the animal, and was monitored and resolved through antibiotic therapy. Zoological institutions are currently examining the use of thermography in many other nondomestic species.
0839_frame_28fm Page 650 Tuesday, May 22, 2001 11:56 AM
FIGURE 7 Thermogram of a dolphin with a severe, generalized, histiocytic dermatitis. Rostrum is to the right and leading edge of right pectoral flipper to the lower left. There were no changes in inflammatory indicators of the blood in spite of this dramatic image.
FIGURE 8 Thermogram of the right hind limb of a polar bear that was limping. The image clearly shows the area of involvement of the outer toe without handling or anesthesia. Heat print left behind on the concrete is to the left.
Web Sites More information on thermography can be viewed at a number of Web sites. eMergeVision.com (http://www.emergevision.com) has pioneered equine and exotic animal use. Sierra Pacific Infrared Inspection Services (SPI) has an extended explanation of the physics of thermography on its site at http://www.x26.com.
0839_frame_28fm Page 651 Tuesday, May 22, 2001 11:56 AM
Thermal Imaging of Marine Mammals
651
Conclusion As new diagnostic techniques become available, each goes through successive periods of research, exposure, initial application, acceptance, then utilization. Whether a technique is widely utilized is often dependent on its level of complication, its cost, and its economic benefits. At this point, thermography has not received wide enough exposure to gauge its true future in marine mammal medicine, but its use to date shows great potential as a research and diagnostic tool. Thermography can document heat related to inflammation or active infection before these processes are clinically detectable during examination. With early detection, this knowledge can allow the clinician to prevent more serious injury and begin proper corrective therapy early. In addition, the clinician can follow therapy for more rapid adjustment or extension of antimicrobials based on the thermal response of the tissues. Thermography is a very useful complementary technique to radiography and physical examination, especially when the site of involvement is unknown or uncertain.
References Cockcroft, P.D., Henson, F.M., and Parker, C., 2000, Thermography of a septic metatarsophalangeal joint in a heifer, Vet. Rec., 146: 258–260. Di Blasio, U., 1966, Thermography, Rass. Int. Clin. Ther., 46: 5548–5550. Frymoyer, J.W., and Haugh, L.D., 1986, Thermography: A call for scientific studies to establish its diagnostic efficacy, Orthopedics, 9: 699–700. Goodman, P.H., Healset, M.W., and Pagliano, J.W. et al., 1985, Stress fracture diagnosis by computerassisted thermography, Physiol. Sportsmed., 13: 114. Graf von Schweinitz, D., 1999, Thermographic diagnostics in equine back pain, Vet. Clin. North Am., Equine Pract., 1: 161–177. Isard, H.J., Ostrum, B.J., and Shilo, R., 1969, Thermography in breast carcinoma, Surg. Gynecol. Obstet., 128: 1289–1293. Jones, B.F., 1998, A reappraisal of the use of infrared thermal imaging analysis in medicine, IEEE Trans. Med. Imaging, 17: 1019–1027. Kasting, N.W., Hayword, J., Hewlett, K.G., and Penberton, D., 1990, Surface heat loss in the killer whale (Orcinus orca) as measured by infrared thermography, in Proceedings of the 21st International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Love, T.J., 1980, Thermography as an indicator of blood perfusion, Ann. N.Y. Acad. Sci., 335: 429. Purohit, R.C., and McCoy, M.D., 1980, Thermography in the diagnosis of inflammatory disease in the horse, Am. J. Vet. Res., 41: 1167–1174. Roberts, W.W., Dinkel, T.A., Schulan, P.G. et al., 1997, Laparoscopic infrared imaging, Surg. Endosc., 11: 1221–1231. Stromberg, B., 1974, The use of thermography in equine orthopedics, J. Vet. Radiol., 15: 94. Turner, T.A., 1989, Hind limb muscle strain as a cause of lameness in horses, in Proceedings of the Annual Meeting of the American Association of Equine Practitioners, 34: 281. Turner, T.A., 1991, Thermography as an aid to the clinical lameness evaluation, Vet. Clin. North Am. Equine Pract., 2: 311–338. Turner, T.A., Purohit, R.C., and Fessler, J.F., 1986, Thermography: A review in equine medicine, in Compendium of Continuing Education for the Practicing Veterinarian, 8: 855–886 Vaden, M.F., Purohit, R.C., and McCoy, M.D., 1980, Thermography: A technique for the subclinical diagnosis of osteoarthritis, Am. J. Vet. Res., 41: 1175. Walsh, M.T., Jones, D., Rankin, D., and Turner, T. A., 1988, Diagnostic visualization techniques in marine mammals, in Proceedings of the 19th International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive.
0839_frame_28fm Page 652 Tuesday, May 22, 2001 11:56 AM
652
CRC Handbook of Marine Mammal Medicine
Walsh, M.T., Turner, T.A., Dover, S., Wood, C., and Wood, L., 1993, Thermography as a diagnostic tool in marine animals, in Proceedings of the 24th International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Weil, M., Litzke, L.F., and Fritsch, R., 1998, Diagnostic validity of thermography lameness in horses, Tierärztl. Prax. Ausgabe Grosstiere Nutztiere, 26: 346–354. Wu, B.M., Lam, F.K., Chan, F.H. et al., 1997, Computerized infrared imaging for studying thermal activation on the skull following somatic stimulation in small animals, Med. Biol. Eng. Comput., 35: 587–594.
0839_frame_C29 Page 653 Tuesday, May 22, 2001 11:57 AM
VI Medical Management of Marine Mammals
0839_frame_C29 Page 654 Tuesday, May 22, 2001 11:57 AM
0839_frame_C29 Page 655 Tuesday, May 22, 2001 11:57 AM
29 Marine Mammal Anesthesia Martin Haulena and Robert Bruce Heath
Introduction Marine mammal anesthesia is an area of ongoing interest and study; new anesthetic agents, monitoring techniques, and anesthetic protocols are regularly introduced. Several older, excellent reviews of marine mammal anesthesia have been written and should be consulted for a more complete history of marine mammal anesthesia (Gales, 1989; Williams et al., 1990). Definitive anesthetic techniques for each species are not presented here because ongoing research, specific situations, and species differences dictate that the veterinarian planning an anesthetic procedure be familiar with all choices, and select the protocol best suited to the task at hand. Anesthesia can be affected by anatomical and physiological adaptations to life at sea. The dive reflex, for example, is a complex set of physiological adaptations that allow breath holding and conservation of oxygen. Although most terrestrial vertebrates possess a dive reflex, in airbreathing vertebrates that depend upon an aquatic environment for feeding, shelter, or breeding, the dive reflex is much more developed. In marine mammals, part of the dive reflex consists of bradycardia and shunting of blood away from peripheral tissues to conserve oxygen for vital tissues such as the heart and brain. Anatomical adaptations, efficient tissue oxygen-carrying capacity, and a tolerance for high carbon dioxide levels and acidosis increase breath-holding ability. An excellent review of marine mammal physiology as it pertains to an aquatic existence has recently been written (Elsner, 1999). Many of these adaptations can complicate anesthesia, and activation of the dive reflex has been implicated as the cause of several deaths of anesthetized marine mammals (Gales and Burton, 1988; Phelan and Green, 1992).
Anesthetic Protocol Preanesthetic Examination 1. 2. 3. 4.
Begin a patient record. Note patient presentation, demeanor, and previous anesthetic history. Note any medications being given to the animal. Perform as complete a physical examination as possible while recognizing that one may be dealing with a free-ranging animal. 5. Obtain and review clinical laboratory data if possible. 6. Generate an impression of anesthetic risk. 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
655
0839_frame_C29 Page 656 Tuesday, May 22, 2001 11:57 AM
656
CRC Handbook of Marine Mammal Medicine
Choice of a Specific Anesthetic Protocol Veterinarians should ask themselves the following questions before choosing an appropriate anesthetic protocol: 1. Is the procedure required, and are there safer alternatives for the animal and personnel for reaching the desired objective? 2. Is the animal a suitable candidate for the procedure? 3. What type of procedure is planned, and what depth of anesthesia is desired? 4. To which anatomical region is the procedure confined? 5. What preanesthetic conditions exist in the animal that may affect the immobilization, metabolism of the agents, or recovery? 6. What precautions can be taken to prevent complications stemming from those conditions? 7. What type of facility is available in which to perform the immobilization? 8. What types of emergency equipment and supplies are available? 9. Which anesthetic agents have been evaluated in this species and for this procedure? 10. Which anesthetic agents are available? 11. What levels of expertise do the available personnel have with the various anesthetic agents, and with the species involved? 12. What potential complications can arise due to the anatomy and physiology of the species involved, and how can they be prevented? 13. What complications can arise due to the procedure, and how can these be prevented?
Monitoring Techniques Monitoring of physiological parameters is an attempt to recognize changes in a patient undergoing anesthesia, so that any changes can be reacted to, and adequate support provided before irreversible effects can occur. It is best to appoint one person whose sole responsibility is monitoring the patient during an anesthetic procedure. Electronic monitors should not be relied upon to the exclusion of direct assessments of the animal, such as respiration and depth of anesthesia. Trends in values are often more important indicators of physiological change than single values, and adequate record keeping assures that trends in any measured parameter are recognized and addressed. Several factors are important in choosing monitoring equipment and technique. These may include the ability to use a piece of equipment given the varied anatomy and physiology of many marine mammal species, the cost, familiarity, and experience of the personnel involved, the relative worth of the information with respect to the time and expertise required to use a piece of equipment, and the available facilities. It is possible for an assigned anesthetist to become absorbed in watching an electronic monitor or too reliant on a single parameter and not notice an important change in some other factor, such as depth of anesthesia. Multiple parameters should be monitored, and limitations of each monitor or parameter should be recognized. These parameters will be discussed as they apply to each group of marine mammals in the following sections. Unfortunately, normal values for many of the monitored parameters have not been well established in most marine mammal species, and the effects of the various anesthetic agents are even less understood. Both these areas of study deserve additional investigation. The desired depth of anesthesia for a procedure, combined with continual assessment of the depth, will avoid excessive depression of the patient. Acute changes or trends in each monitored parameter should be addressed immediately. The anesthetist must be aware of potential emergencies and be prepared for them before they happen so response is not delayed.
0839_frame_C29 Page 657 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
657
Noninvasive Techniques A: B: C: D: E: F: G: H:
Assessment of reflexes (including withdrawal, palpebral and pupillary reflexes, and jaw tone) Stethoscope Temperature probe Pulse oximeter Capnometer Electrocardiogram Indirect blood pressure Doppler flow probe
Invasive Techniques A: B: C: D:
Central venous pressure Arterial blood pressure Blood gas parameters Cardiac output (usually only used in hospital or research situations)
Support Since anesthesia usually results in a decrease in the patient’s ability to maintain homeostasis, an important role for the anesthetist is support. Patient support may be preventative in nature, or occur as a response to changing monitored parameters. Support may include relatively simple procedures, such as adequate positioning, to limit ischemia from inappropriate weight bearing, or allowing increased thoracic movements, to prevent respiratory acidosis. More complicated methods of support include continuous intravascular (IV) fluids or assisted mechanical ventilation. Patient support is discussed in the following sections for each group of marine mammals.
Cetaceans The successful anesthesia of cetaceans has been accomplished by very few individuals. The use of careful training methods with display animals has greatly enhanced the ability to perform routine diagnostic procedures without the need for chemical restraint. A combination of the use of sedative agents such as benzodiazepines or meperidine along with physical restraint can aid in accomplishing more stressful procedures such as gastroscopy or bronchoscopy (Greenwood et al., 1978; Joseph and Cornell, 1988; Hawkins et al., 1997; Reidarson et al., 1998). Local anesthesia can also be employed for various procedures and published techniques include tooth extraction using lidocaine to anesthetize the mandible (Ridgway et al., 1975). More recently, lidocaine has been successfully delivered into the periodontal ligaments for tooth extraction and into the dorsal fin for attaching instrument packages using a dental anesthesia injector (Townsend, pers. comm.). Lidocaine has also been used as an aid in intubation (Rieu and Gautheron, 1968), and tetracaine has been used to facilitate bronchoscopy (Reidarson et al., 1998). Table 1 is a summary of some agents that have been used to immobilize cetaceans.
Induction Intramuscular (IM) atropine at 0.02 mg/kg has been given as a preanesthetic agent (Ridgway et al., 1974). Injectable induction agents have been given by the IV route by administration into the fluke veins (Ridgway and McCormick, 1967; 1971) and the caudal vein (Rieu and Gautheron,
Thiopental, halothane Meperidine Thiopental
Meperidine
3
1
6
4
Pseudorca crassidens (false killer whale)
Lagenorhynchus obliquidens (Pacific white-sided dolphin) and Tursiops truncatus (Atlantic bottlenose dolphin) Orcinus orca (killer whale)
Halothane
1
Lagenorhynchus obliquidens (Pacific white-sided dolphin)
10
Meperidine
Meperidine
0.23–0.45 mg/kg
0.23–0.45 mg/kg
10 mg/kg
10–15 mg/kg 1.0–2.0% 0.23 mg/kg
0.75–3.5%
60–80%
0.23–0.45 mg/kg
7 mg/kg Unknown
Dosage
IM
IM
IV
IV IH IM
IH
IH
IM
IV IH
Route
0
0
0
0
na
0
0
0
0
Mortality, %
Maximum effect in 20 min; duration of moderate sedation, 2 to 3 h; dosedependent sedation and analgesia Maximum effect in 20 min; duration of moderate sedation, 2 to 3 h; dosedependent sedation and analgesia
Immediate apnea and loss of reflexes after thiopental only; recovery 21 min after thiopental injection Maximum effect in 20 min; duration of moderate sedation, 2 to 3 h; dosedependent sedation and analgesia Cyanosis when nitrous oxide at 80%, return to normal reflexes and spontaneous breathing at 70% Faster induction with higher halothane setting; maintenance of surgical plane with 0.75–1% halothane Mild hepatic lesions noted; euthanatized prior to recovery; maximum effect in 20 min; duration of moderate sedation, 2 to 3 h Light anesthesia with most reflexes except blowhole present; return of respiration only after 1 to 2.5 h after injection
Comments
Joseph and Cornell, 1988
Joseph and Cornell, 1988
Joseph and Cornell, 1988 Ridgway and McCormick, 1971
Medway et al., 1970
Ridgway and McCormick, 1967
Ridgway and McCormick, 1967
Joseph and Cornell, 1988
Rieu and Gautheron, 1968
Ref.
658
3
Nitrous oxide
4
Globicephala macrorhynchus (short-finned pilot whale)
Thiopental, nitrous oxide
1
Delphinus delphis (common dolphin)
Agent
n
Species
TABLE 1 Some Chemical Immobilizing Agents Used in Cetaceans
0839_frame_C29 Page 658 Tuesday, May 22, 2001 11:57 AM
CRC Handbook of Marine Mammal Medicine
Nitrous oxide Halothane
Thiopental, halothane Thiopental, halothane Diazepam Ketamine Meperidine
Propofol, isoflurane Diazepam Propofol, isoflurane
2 10
5
1 na 74
1 1 1
6
Thiopental, methoxital
1
Tursiops truncatus (Atlantic bottlenose dolphin)
Thiopental, nitrous oxide
1
Stenella styx (coeruleoalba) (striped dolphin)
3.5 mg/kg 2% 0.2 mg/kg na
0.11–0.45 mg/kg
0.11 mg/kg 1.1 mg/kg
10 mg/kg 1–3.5% 10–15 mg/kg 1.0–2.0%
0.75–3.5%
60–90%
26 mg/kg 5 mg/kg
4 mg/kg 70%
IV IH PO IV IH
IM
IM na
IV IH IV IH
IH
IH
IP
IV IH
0 0
0
0
0 0
na
0
0
0
100
0
Tranquilization with animal motionless at water surface Maximum effect in 20 min; duration of moderate sedation, 2 to 3 h; dosedependent sedation and analgesia Anesthetized for 1 h, 37 min; good recovery Given 1 h prior to bronchoscopy
Faster induction with higher halothane setting Maintenance of surgical plane with 0.75–1% halothane Induction with 2% halothane after thiopental Anesthetized for up to 24 h at surgical plane; euthanatized prior to recovery
Minimal loss of reflexes; no cyanosis
Immediate apnea and loss of reflexes after gas and injectable are given simultaneously; recovery 13 min after introduction of 100% oxygen Extremely slow recovery
Linnehan and MacMillan, 1991 Reidarson et al., 1998 Dover et al., 1999
Hawkins et al., 1997 Sweeney and Ridgway, 1975 Joseph and Cornell, 1988
Ridgway and McCormick, 1967 Medway et al., 1970
Ridgway and McCormick, 1967 Ridgway and McCormick, 1967
Nagel et al., 1964
Rieu and Gautheron, 1968
0839_frame_C29 Page 659 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia 659
0839_frame_C29 Page 660 Tuesday, May 22, 2001 11:57 AM
660
CRC Handbook of Marine Mammal Medicine
1968). Intravenous induction agents that have been used successfully include thiopental (Ridgway and McCormick, 1967; 1971) and, more recently, propofol (Linnehan and MacMillan, 1991; Dover et al., 1999). Induction has also been accomplished in intubated cetaceans by inhalant anesthetic agents such as halothane (Ridgway and McCormick, 1967). Intramuscular midazolam has been given to one stranded gray whale (Eschrichtius robustus) to produce sedation prior to euthanasia (see Chapter 32, Euthanasia).
Intubation Intubation of most cetaceans has usually been accomplished by manual dislocation of the modified larynx from the nasopharyngeal sphincter of the blowhole passage by reaching through the mouth, pulling the larynx anteroventrally, and then slipping an endotracheal tube into the trachea after inserting two fingers through the glottis (Nagel et al., 1964; Ridgway and McCormick, 1971). Equine endotracheal tubes of between 24 to 30 mm diameter have been used (Ridgway and McCormick, 1971). Care must be taken not to intubate only one main bronchi since the trachea bifurcates relatively early and an endotracheal tube should not be passed more than 20 cm past the larynx in bottlenose dolphins (Tursiops truncatus) (Ridgway and McCormick, 1971). Although intubation has been accomplished in fully awake cetaceans prior to administration of an inhalant anesthetic agent, the administration of an injectable induction agent makes the procedure much easier (Ridgway and McCormick, 1967; 1971; Linnehan and MacMillan, 1991). Smaller cetaceans, in which a hand cannot be passed easily through the mouth to disconnect the larynx, have been intubated through the blowhole after topical anesthetic (2% lidocaine) has been injected into the blowhole, different levels of the nasal passages, and applied directly through the endotracheal tube to the larynx and trachea to prevent spasm of the glottis (Rieu and Gautheron, 1968).
Inhalation Anesthesia Small cetaceans have been anesthetized with nitrous oxide and halothane (Ridgway and McCormick, 1967) and isoflurane (Linnehan and MacMillan, 1991; Dover et al., 1999) via inhalation (IH). Nitrous oxide did not produce sufficient loss of peripheral reflexes without first resulting in cyanosis and is not thought to be an adequate anesthetic agent in cetaceans (Ridgway and McCormick, 1967; 1971). Halothane has been found to produce a reliable surgical plane of anesthesia (Ridgway and McCormick, 1967). However, there has also been some hepatic toxicity reported as a result of prolonged use of halothane (Medway et al., 1970). Isoflurane is at present in more common use (Ridgway, pers. comm.).
Monitoring Depth of anesthesia is assessed in much the same way as in other marine and terrestrial mammals. Reflexes that have been used to indicate depth of anesthesia include the palpebral, corneal, and gag reflexes (Ridgway and McCormick, 1967; 1971). In addition, withdrawal of the tongue, movement in response to distension of the anus, movement of the peduncle, movement of the pectoral muscles in response to scratch or needle stick, movement of the blowhole in response to touch, or movements of the vagina or penis when they were stimulated, have also been used to assess the depth of anesthesia (Ridgway and McCormick, 1967; 1971). Loss of the swimming motions made by the tail flukes was found to be the most reliable indicator of a surgical plane of anesthesia (Ridgway and McCormick, 1967). However, many restraint devices make the determination of that reflex difficult as they do not allow the animals
0839_frame_C29 Page 661 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
661
to show any peduncle movement (Ridgway and McCormick, 1971). Return of the blowhole reflex indicates that extubation can take place (Ridgway and McCormick, 1967; McCormick, 1969; Sweeney and Ridgway, 1975). The heart rate of halothane-anesthetized bottlenose dolphins is approximately 108 to 120 beats/ min (bpm) (Ridgway and McCormick, 1967). Electrocardiograms (ECGs) have been monitored in anesthetized cetaceans (Ridgway and McCormick, 1967; Ridgway et al., 1974; Linnehan and MacMillan, 1991). A heart rate below 60 bpm is considered to be of concern in an anesthetized cetacean (Ridgway and McCormick, 1971). Temperature has been monitored using flexible temperature probes inserted at least 20 cm into the rectum (Nagel et al., 1964; Ridgway and McCormick, 1967; Ridgway et al., 1974). However, the anesthetist should be aware that, in males, placing the temperature probe in the area of the gonads may result in the recording of a false hypothermia due to the presence of a vascular plexus used for cooling the testes (Rommel et al., 1994). A core temperature of between 36.0 and 37.5°C has been considered adequate in anesthetized bottlenose dolphins (Ridgway et al., 1974). Mild hypothermia with a temperature of 35.3°C was noted during the recovery of a bottlenose dolphin anesthetized with isoflurane (Linnehan and MacMillan, 1991). Blood gas parameters, including PO2, PCO2, and pH, have been recorded from central arteries and veins found in the flukes, and values correlate well with those taken from the carotid artery (Ridgway and McCormick, 1967; Rieu and Gautheron, 1968). Values have been maintained at 95 to 120 mmHg, 30 to 45 mmHg, and 7.2 to 7.4 for PO2, PCO2, and pH, respectively, in anesthetized bottlenose dolphins (Ridgway et al., 1974). Oxygen from inspired and expired air has been measured via an oxygen analyzer (Ridgway and McCormick, 1967). End-tidal carbon dioxide (EtCO2) has also been monitored (Ridgway and McCormick, 1967). Pulse oximetry has been used in isoflurane-anesthetized bottlenose dolphins to monitor O2 saturation via a lingual clip, with recorded values of 96 to 98% (Linnehan and MacMillan, 1991). Mean arterial blood pressure (MAP) has been measured in bottlenose dolphins and Pacific white-sided dolphins (Lagenorhynchus obliquidens) (Ridgway and McCormick, 1971; Ridgway et al., 1974). MAP varied from 120 to 130 mmHg in resting bottlenose dolphins and dropped to an average of 115 mmHg under halothane anesthesia when measured from the tail stock. These values were slightly higher in Pacific white-sided dolphins, 145 and 130 mmHg before and during anesthesia, respectively. Kidney perfusion has been assessed by monitoring urine output (Ridgway et al., 1974).
Support Temperature regulation is an important aspect of cetacean sedation, and can be accomplished by regulating the water temperature immediately surrounding an anesthetized animal in response to its core temperature (Ridgway et al., 1974). Some tranquilizing agents such as the phenothiazines (acepromazine and trifluromeprazine) may cause peripheral vasodilation and subsequent hypothermia (Ridgway and McCormick, 1971). In one of these cases, low internal body temperature may have resulted in apnea and the animal had to be placed on a respirator and kept moist with warm water until core temperature increased (Ridgway and McCormick, 1971). Assisted ventilation must be used to supplement respiration in anesthetized cetaceans that appear to become apneic as soon as consciousness is lost (Ridgway and McCormick, 1967; 1971; Rieu and Gautheron, 1968). Ventilators should have an apneustic plateau control unit to mimic the normal respiratory pattern of cetaceans (McCormick, 1969; Ridgway and McCormick, 1971) to allow for sufficient inflation time for oxygenation of blood (Ridgway et al., 1974). Respiratory rate has been set at approximately three breaths/min (Ridgway and McCormick, 1967).
0839_frame_C29 Page 662 Tuesday, May 22, 2001 11:57 AM
662
CRC Handbook of Marine Mammal Medicine
Bottlenose dolphins have a tidal volume which is usually 5 to 10 l, an average repiratory rate of two to three breaths/min, and an apneustic plateau that is usually held for 20 to 30 s after inspiration, followed by rapid exhalation and inhalation with flow rates through the air passages of between 30 and 70 l/s (Ridgway and McCormick, 1971). Williams et al. (1990) recommend a 30-s plateau and ventilating at 80% of the tidal volume for bottlenose dolphins. Since cetaceans live entirely in an aquatic environment where their buoyancy is near neutral, their body structure was not designed to support weight for any period of time. Therefore, restraint and positioning for anesthetic procedures out of the water may interfere with adequate circulation as well as impede thoracic movements, compromising cardiac and respiratory function (Ridgway and McCormick, 1971; Ridgway et al., 1974). This necessitates attempting to distribute weight as evenly as possible out of the water and choosing anesthetic agents that allow for rapid recovery and early return to the water (Ridgway and McCormick, 1971). The use of a specially designed surgery table that is partially filled with water, if the desired procedure allows for it, reduces weight bearing (Ridgway et al., 1974). Fluid volume has been maintained by the use of IV lactated Ringer’s solution via slow drip into a fluke vein (Linnehan and MacMillan, 1991).
Emergencies It is critical to monitor for return of normal respiratory and blowhole reflexes prior to extubation. If apnea continues for more than 3 min, or heart rate drops below 60 bpm, the animal should be reintubated and assisted ventilation provided (Ridgway and McCormick, 1971). Severe blood loss may occur with certain surgical procedures, and blood transfusions from conspecifics can be performed to maintain adequate blood volume (Ridgway et al., 1974).
Otariids Otariids are among the most popular marine mammals kept in public display facilities, are seen in many rehabilitation facilities, and are the subject of several field research programs. They are, therefore, some of the more commonly anesthetized marine mammals. Sedative agents can be used in conjunction with physical restraint to immobilize animals adequately for many procedures (Gales, 1989). Recently, the benzodiazepine midazolam has been reported for use as an IM agent to augment physical restraint in fur seals at 0.25 to 0.35 mg/kg (Lynch et al., 1999b) and at 0.15 to 0.2 mg/kg in California sea lions (Zalophus californianus) (Haulena, unpubl. data). Table 2 is a summary of recent anesthetic agents used in various otariids since 1989.
Induction Atropine IM at 0.02 mg/kg has been shown to prevent some of the detrimental aspects of the development of the dive reflex during anesthesia, such as bradycardia (Gage, 1993; Lynch et al., 1999b), and has also been used to decrease respiratory secretions (Sepulveda et al., 1994). Atropine should be delivered IM at least 10 min prior to the introduction of the anesthetic agent in otariids (Gage, 1993; Haulena et al., 2000). The action of atropine in otariids is not completely understood, and some anesthetists do not recommend its use (Woods, pers. comm.). Glycopyrrolate has been suggested as an alternative due to its longer activity in terrestrial species, but its use has not been reported in marine mammals. Because of poorly accessible peripheral vasculature, most induction in otariids has been accomplished by IM injection (Bester, 1988; Loughlin and Spraker, 1989; Heard and Beusse, 1993;
Tiletamine and zolazepam Ketamine Ketamine, diazepam Ketamine, xylazine Ketamine, xylazine Ketamine, xylazine Ketamine, diazepam
172
45
Arctocephalus phillipi (Juan Fernández fur seal)
Arctocephalus forsteri (New Zealand fur seal) Arctocephalus gazella (Antarctic fur seal)
12
7
14
30 23
5
1
8
4
Tiletamine and zolazepam Tiletamine and zolazepam, ketamine Tiletamine and zolazepam, ketamine Ketamine, midazolam Isoflurane
32
Arctocephalus australis (South American fur seal)
Agent
n
Species
IM
7
IM dart IM dart IM IM IM dart 17
0
14
0 4
IM dart IM dart IM dart
6.9 ± 0.1 mg/kg 6.4 ± 0.1 mg/kg 0.006 ± 0.001 mg/kg 7.3 ± 0.3 mg/kg 0.6 ± 0.02 mg/kg 3.8–10.8 mg/kg 0.7–2.0 mg/kg 5.6–7.8 mg/kg 0.5–1.3 mg/kg 2.16–6.76 mg/kg 0.04–0.28 mg/kg
3
IM dart
0
0
0
0
0
Mortality, %
1.2–1.7 mg/kg
IH
IM dart
IM IM dart
0.81 mg/kg 1.15 mg/kg 0.27 mg/kg 1 mg/kg 0.1 mg/kg 1.2–4.0%
IM dart
IM dart
Route
1.43 mg/kg
1.43 mg/kg
Dosage
TABLE 2 Some Recently Published Chemical Immobilizing Agents Used in Otariids
Increased induction and recovery times than when used IV; variable plane of anesthesia
Poor sedation with ketamine less than 5.6 mg/kg
Muscle tremors
Respiratory depression
Supplemental ketamine given due to insufficient sedation; partial reversal with flumazenil All chemical agents administered at once; partial reversal with flumazenil
Partial reversal with flumazenil
Comments
(Continued)
Ferreira and Bester, 1999 Sepulveda et al., 1994
Bester, 1988
Boyd et al., 1990
Boyd et al., 1990 Boyd et al., 1990
Gales and Mattlin, 1998 Boyd et al., 1990
Karesh et al., 1997
Karesh et al., 1997
Karesh et al., 1997
Karesh et al., 1997
Reference
0839_frame_C29 Page 663 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia 663
Ketamine, diazepam Ketamine
Ketamine, xylazine Carfentanil, xylazine
Carfentanil, xylazine, azaperone Carfentanil, xylazine, azaperone, ketamine Carfentanil, xylazine, ketamine Xylazine, azaperone Droperidol Ketamine, xylazine Tiletamine and zolazepam
10
7
7
2
15
29
2 32
2
5
27
Agent
n
IM dart IM dart IM dart IM dart IM dart IM dart IM dart
6–18 µg/kg na na na 6–18 µg/kg na na
IM dart
IM dart IM dart IM IM dart
IM dart IM dart IM dart
6–18 µg/kg na na
0.57–2.0 mg/kg 0.57–2.0 mg/kg na 3.1–11.4 mg/kg 0.3–1.7 mg/kg 1.8–8.1 mg/kg
IM dart IM dart IM dart IM dart
IM dart
IV
Route
4.2–5.2 mg/kg 0.6–0.9 mg/kg 6–18 µg/kg na
2.16–6.76 mg/kg 0.04–0.28 mg/kg 4.3–7.8 mg/kg
Dosage
21
50 13
7
na
na
Best results were found when using 1.8–2.5 mg/kg
Variable plane of anesthesia
20% of animals that were given some combination with carfentanil died; apnea, muscle convulsions; variable plane of anesthesia 20% of animals that were given some combination with carfentanil died; apnea, muscle convulsions; variable plane of anesthesia 20% of animals that were given some combination with carfentanil died; apnea, muscle convulsions; variable plane of anesthesia 20% of animals that were given some combination with carfentanil died; apnea, muscle convulsions; variable plane of anesthesia Sufficient sedation for branding; short immobilization time
na
na
Xylazine dosage estimated
Deeper immobilization compared with IM administration; variable plane of anesthesia
Comments
29
19
0
Mortality, %
David et al., 1988 Ferreira and Bester, 1999 Loughlin and Spraker, 1989
David et al., 1988
David et al., 1988
David et al., 1988
David et al., 1988
David et al., 1988
David et al., 1988
Sepulveda et al., 1994 David et al., 1988
Reference
664
Arctocephalus tropicalis (subantarctic fur seal) Eumetopias jubatus (Steller’s northern sea lion)
Arctocephalus pusillus pusillus (South African fur seal)
Species
TABLE 2 Some Recently Published Chemical Immobilizing Agents Used in Otariids (continued)
0839_frame_C29 Page 664 Tuesday, May 22, 2001 11:57 AM
CRC Handbook of Marine Mammal Medicine
Phocarctos hookeri (Hooker’s (New Zealand) sea lion) Zalophus californianus (California sea lion)
Otaria byronia (South American sea lion)
Detomidine, ketamine Isoflurane Tiletamine and zolazepam Isoflurane Halothane Medetomidine, ketamine Medetomidine, ketamine, isoflurane Medetomidine, tiletamine and zolazepam Medetomidine, tiletamine and zolazepam, 665isoflurane
4
16
115 30 35
60
Isoflurane Isoflurane
Tiletamine and zolazepam, isoflurane Tiletamine and zolazepam
7 29
13
51
IH IH IM IM IM IM IH IM IM IM IM IH
70 µg/kg 1 mg/kg 1–5%
IM IM IH IM
IH IH
IM dart IM dart IH IM
0.75–3% 0.75–5% 140 µg/kg 2.5 mg/kg 140 µg/kg 2.5 mg/kg 1–5% 70 µg/kg 1 mg/kg
40–55 µg/kg 2.0–4.3 mg/kg 1–5% in oxygen 1.7 mg/kg
na 0.8–4.0%
2.75 mg/kg
1.6–3.3 mg/kg
0
6
0
0 3 0
0
0
0 0
0
10
More consistent plane of anesthesia than when ketamine was used but recovery not as smooth
Variable plane of anesthesia, reversal with atipamezole Reversal with atipamezole
Apnea
Flumazenil given at 1 mg for every 20–25 mg of tiletamine and zolazepam for reversal
Haulena and Gulland, in press
Haulena and Gulland, in press
Haulena et al., 2000
Heath et al., 1997 Work et al., 1993 Haulena et al., 2000
Gage, 1993
Heard and Beusse, 1993
Karesh et al., 1997 Gales and Mattlin, 1998
Karesh et al., 1997
Heath et al., 1996
0839_frame_C29 Page 665 Wednesday, May 23, 2001 10:54 AM
Marine Mammal Anesthesia 665
0839_frame_C29 Page 666 Tuesday, May 22, 2001 11:57 AM
666
CRC Handbook of Marine Mammal Medicine
Heath et al., 1996) and inhalation (Work et al., 1993; Heath et al., 1997). Some authors have used IV induction in animals that have been physically restrained with the aid of restraint boards. Sepulveda et al. (1994) injected ketamine and diazepam into the cephalic vein and also into the epidural vein at the level of the front flippers in Juan Fernández fur seals (Arctocephalus philippii). Caution should be used when attempting injections into the epidural vein of otariids to prevent damage to the spinal cord. The caudal gluteal vein can be accessed (see Chapter 19, Clinical Pathology). However, it often requires maximal physical restraint not to dislodge the needle and cause perivascular damage. Interdigital veins may be used to inject chemical agents in some otariids, but are difficult to catheterize. Induction by inhalation is possible in situations that allow for restraint of the patient, adequate equipment, space for equipment, and personnel who are experienced with the procedure. Physical restraint may not be possible in many field situations or when dealing with large animals (Work et al., 1993). Sites chosen for IM injection have the largest available muscle mass with the least amount of overlying blubber, and can be accessed by a dart or by hand injection. IM injection sites that are most favored include the muscle overlying the hips and tibia (Loughlin and Spraker, 1989; Heard and Beusse, 1993; Sepulveda et al., 1994; Haulena et al., 2000), lumbar muscle (Bester, 1988; Loughlin and Spraker, 1989), and muscle over the shoulder (Loughlin and Spraker, 1989). Darting has been used in field (Loughlin and Spraker, 1989; Heath et al., 1996) and captive (Haulena et al., 2000) situations. Because of poor penetration to muscle layers due to thick blubber layers (Geraci, 1973) or inaccurate darting, results are more variable when darting is employed (in comparison to hand injection) to deliver anesthetic agents (Haulena et al., 2000). Free-ranging animals that are to be darted must be chosen carefully to prevent escape and potential drowning and must be in areas that can be easily, quickly, and safely approached once the dart has been delivered (Heath et al., 1996). Additional doses of zolazepam-tiletamine (ZT) (Loughlin and Spraker, 1989; Haulena, unpubl. data) and xylazine-ketamine (Bester, 1988) given to animals that have not been induced adequately, or to prolong immobilization, have been associated with mortalities. In some cases, anesthetists have chosen to abandon attempts at immobilization if adequate sedation is not accomplished after the first injection (Heath et al., 1996). However, additional ketamine after initial injection of medetomidine-ketamine in a small number of California sea lions did not result in any mortalities (Haulena et al., 2000), and additional ketamine has been used after initial ZT in South American fur seals (Arctocephalus australis) (Karesh et al., 1997). Induction by inhalation has been accomplished by placing either commercially available masks (Haulena et al., 2000) or customized masks, made from such items as soft polyurethane traffic cones, over the muzzle of the animal, thereby creating a seal that allows the animal to breathe the anesthetic agent (Work et al., 1993; Heath et al., 1996; 1997). Induction by inhalation has also been used in conjunction with injectable agents if adequate sedation is not achieved with the injectable agent alone (Heard and Beusse, 1993; Heath et al., 1996; Haulena et al., 2000).
Intubation Intubation of anesthetized animals is strongly recommended to maintain an airway to ensure adequate oxygenation, especially in an emergency situation and to prevent aspiration secondary to regurgitation or vomition (Work et al., 1993). Intubation of marine mammals may be especially prudent, as apnea may occur in these carbon dioxide–tolerant species (Sedgwick, 1999), and the anesthetist should be ready to provide ventilatory support. Many of the mechanical restraint devices used on marine mammals may cause undesirable pressure to the thorax
0839_frame_C29 Page 667 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
667
that decreases ventilation or partially obstructs the airway. The anesthetist must be aware of potential complications from the use of these mechanisms and the increased need for intubation to maintain adequate gas exchange. Training and experience are necessary to accomplish intubation quickly and easily. Gentle technique minimizes trauma to the larynx and associated tissues. Mouths can be kept open with woven nylon straps approximately 2 cm wide and of sufficient length to prevent placing hands between the jaws. Laryngoscopes facilitate passing tubes in smaller individuals (15 to 150 kg); 140- to 150-mm McIntosh (Rusch, Inc., Duluth, GA) (Work et al. 1993, Haulena et al., 2000) and 110- to 260-mm Miller laryngoscope blades have been used successfully in California sea lions (Work et al., 1993; Heard and Beusse, 1993; Heath et al., 1996). Large individuals over 200 kg can be intubated by the manual technique of passing one’s forearm into the oral cavity, palming the endotracheal tube, and placing it gently into the trachea after palpating the glottis (Heath et al., 1996). This necessitates fairly deep anesthesia during induction and speed. With experience and training, intubation may also be accomplished by blind techniques while listening for inhalation by the patient. Of vital importance are listening, timing, and gentle redirection. Care should be taken to guard against tracheal compression by netting, restraint devices, or table edges. The tracheal rings of otariids are incomplete and the trachea is easily collapsed, thereby preventing successful intubation (Lynch et al., 1999b). Endotracheal tube diameter can be expected to be of the same general size as a terrestrial mammalian carnivore of the same size. Cuffed endotracheal tubes are recommended to prevent aspiration; using excessively large tubes may cause laryngeal trauma. Positioning of the animal is extremely important. A straight neck with a slight opisthotonic position and extension of the head of the patient is recommended. The length of the tube is extremely important because of the early, prethoracic bifurcation of the trachea into primary bronchi in otariids (McGrath et al., 1981). Care must be taken to prevent unilateral lung intubation and the associated potential for ventilation/ perfusion mismatch. After intubation, the cuff can be inflated and pulled back to ensure an adequate seal (Woods, pers. comm.). Tubes should be secured using rolled gauze or tape by tying a knot around the tube and then around the maxilla of the animal, caudal to the canine teeth.
Inhalation Anesthesia Inhalation agents including isoflurane (Heard and Beusse, 1993; Heath et al., 1996; 1997; Gales and Mattlin, 1998; Haulena et al., 2000), sevoflurane, and halothane (Work et al. 1993) have all been used, with the best recovery characteristics obtained from isoflurane and sevoflurane. Diving animals have tremendous respiratory reserve and efficiency. In diving animals, 46% of the tidal volume enters alveolar exchange, compared with 15% in humans and horses (Williams et al., 1991; Reed et al., 1994). This directly relates to inhalant anesthetic uptake and distribution. Marine mammals are thus able to uptake anesthetic gases very efficiently and are masked to sleep readily, necessitating vigilance on the anesthetist’s part. Controlled studies on the efficacy of masking and maintainence of anesthesia with an inhalant are few for marine mammals. However, California sea lions, New Zealand fur seal (Arctocephalus forsteri) bulls, and adult female New Zealand sea lions (Phocarctos hookeri) appear to be quickly masked to anesthetic depths in comparison with terrestrial species (Heath et al., 1997; Gales and Mattlin, 1998). The use of inhalant anesthetic agents alone appears to be a reliable and safe method of anesthesia in otariids if it is possible to accomplish restraint and masking (see Table 2).
0839_frame_C29 Page 668 Tuesday, May 22, 2001 11:57 AM
668
CRC Handbook of Marine Mammal Medicine
Monitoring Respiratory rate in anesthetized otariids has been monitored using a stethoscope or by observing thoracic movements. Otariids typically display an apneustic breathing pattern while awake, with an exhalation first, then a pause and hold on inspiration. This reverses to a more typical terrestrial mammalian pattern once intubated, with an inspiration, exhalation, and then a pause in exhalation (Heath et al., 1996; 1997). Heart rate has also been monitored via chest auscultation using a stethoscope or by palpation of the intercostal space over the region of the heart. Intrathoracic noises are often dull compared with those of terrestrial mammals (Lynch et al., 1999b) most likely due to thick blubber layers. Perfusion of the peripheral vasculature can be assessed by noting mucous membrane color and capillary refill time (Work et al., 1993; Heath et al., 1997). Depth of anesthesia has been ascertained by responses to various stimuli, including noise, deep pain (interdigital web pinch, ear pinch, surgical stimulation), presence of reflexes, including palpebral and pupillary, respiratory character, and the degree of jaw tone (Work et al., 1993; Heath et al., 1996; 1997). Noninvasive monitoring in otariids has included the use of pulse oximeters with clip probes attached to the distal one third of the tongue (Heard and Beusse, 1993; Heath et al., 1996; 1997; Haulena et al., 2000) or nasal septum. Reflectance probes have been placed rectally; however, fecal matter tends to interfere with adequate readouts. Alternative sites have included buccal (Heath et al., 1996), vaginal, or esophageal mucosa. Pulse oximetry probes must be shielded from direct sunlight because they are dependent upon infrared light for accuracy. Flexible temperature probes have been used in the rectum (Bester, 1988; Loughlin and Spraker, 1989; Ferreira and Bester, 1999) or in the esophagus at the level of the heart (Heath et al., 1997) to record core body temperature. Capnometers have been attached to the endotracheal tube or “Y” piece via a filter line as is used in humans and domestic species (Heard et al., 1993; Haulena and Gulland, 2001) to monitor end-tidal carbon dioxide (EtCO2) and respiratory rate. Doppler flow probes have been used to detect arterial flow between the digits of the front flipper and cardiac flow (Heard and Beusse, 1993). ECG sensors have been attached in appropriate locations via adhesive pads that require shaving, alligator clips attached to a skin fold directly, or by use of a 20-gauge needle placed through a skin fold and then grasped with alligator clips. Heard and Beusse (1993) placed left and right forelimb leads on the left and right front flippers and the left hind limb lead onto the prepuce of male California sea lions. Small amounts of alcohol placed at the points of contact between the clips and skin have improved conductance. Recently, esophageal ECG probes (Vet/Sensor ECG Plus®, Heska Corporation, Fort Collins, CO) have been used successfully in California sea lions to monitor heart rate and developing arrhythmias. Advantages include increased ease of application, decreased artifact due to movement during the procedure, and fewer cables leading from the patient that may interfere with a sterile field during surgery. However, the probes are very dependent upon their position in the esophagus relative to the heart for accurate production of ECG complexes and only report from three leads. Therefore, the probes do not allow for easy interpretation of complexes and for in-depth investigation of cardiac electrical abnormalities. Indirect, oscillometric measurement of the blood pressure has been accomplished in sea lions by placing an appropriately sized cuff over the radial artery at the most proximal portion of the front flipper.
Support Monitoring core temperature changes is important, especially in free-ranging situations. Body temperatures of all anesthetized animals tend to go toward the ambient temperature; thus both hypothermia and hyperthermia are possible. Hyperthermia may be more important in larger animals (Work et al., 1993). Increasing temperature was suspected in contributing to death
0839_frame_C29 Page 669 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
669
associated with sedation of Antarctic fur seals (Arctocephalus gazella) (Bester, 1988). Stress of capture may exacerbate overheating, so animals should be allowed to stabilize after capture prior to attempting anesthesia. Certain agents, such as ketamine, may cause hyperthermia. IV injection of ketamine and diazepam in fur seals resulted in higher internal temperatures than when these drugs were given IM (Sepulveda et al., 1994). Hyperthermia was also noted in South American fur seals after administration of ketamine/xylazine and with combinations containing carfentanil (David et al., 1988). Hypothermia has been noted in adult Steller sea lions (Eumetopias jubatus) anesthetized in Alaska (Loughlin and Spraker, 1989) and in young California sea lions in California anesthetized with halothane (Work et al., 1993). Tents to protect from sunlight, along with cold water applied to extremities (Work et al., 1993; Sepulveda et al., 1994, Heath et al., 1997) help prevent hyperthermia. Wraps, insulating pads, and hot water bottles can warm exposed flippers and aid in decreasing the potential for hypothermia. Timing of procedures to coincide with the least extreme temperatures of the day will help decrease temperature irregularities (Sepulveda et al., 1994; Heath et al., 1997). Bradycardia has been associated with anesthetic agents in otariids (Heath et al., 1996). It is desirable to maintain a heart rate of over 60 bpm in anesthetized California sea lions (Haulena, unpubl. data). Administration of additional atropine at half the full amount of the original dose is performed for longer procedures or if a bradycardia (<60 bpm) occurs. However, both heart and respiratory rates have been shown to be greater in anesthetized fur seals than in those at rest; heart rate was greater in those animals given ketamine and diazepam IV than those that were given the agents IM (Sepulveda et al., 1994). IV catheters are difficult to place in many anesthetized otariids unless cut downs are employed to access vessels. Veins that may have potential for catheterization include the jugular, subclavian, and vessels running along the digits of the hind flipper. Low SpO2 values (<85%) have been reported from sea lions immobilized with ZT (Heath et al., 1996) and with medetomidine-ketamine (Haulena et al., 2000) especially in those animals that were not intubated and supported with oxygen. Conversely, sea lion pups given isoflurane with oxygen maintained higher SpO2 values (Heath et al., 1997). This may be due to the agents that were used, depth of anesthesia, or to the physiology of the animals but, nevertheless, suggests that the anesthetist should be prepared to intubate, provide oxygen therapy, or supplemental ventilation. High EtCO2 values have been reported in anesthetized sea lions (Heard and Beusse, 1993; Haulena and Gulland, 2001). They probably are a reflection of hypoventilation in anesthetized marine mammals, either due to positioning during the procedure or to an effect caused by the immobilizing agents. High EtCO2 levels support the need for assisted mechanical ventilation. However, normal EtCO2 levels have not been evaluated in marine mammals; high levels may represent a high CO2 tolerance that is an adaptation to diving. In this case, mechanical ventilation may overcompensate and result in an undesirable respiratory alkalosis. The effect of assisted ventilation on EtCO2 and blood gas parameters is an area that requires further investigation. To prevent corneal irritation during procedures, eyes of anesthetized otariids are often lubricated with an ophthalmic ointment (Work et al., 1993).
Emergencies Doxapram has been used in otariids to stimulate respiration during prolonged apnea and has been administered successfully by either the IV route or by injection into the tongue (Work et al., 1993; Heath et al., 1996). Dosages usually approximate those used in terrestrial carnivores of the same size (5 mg/kg). If not already intubated, animals with prolonged apnea should be
0839_frame_C29 Page 670 Tuesday, May 22, 2001 11:57 AM
670
CRC Handbook of Marine Mammal Medicine
intubated and ventilatory support provided. Epinephrine has also been administered to animals with severe bradycardia or with cardiac arrest (Work et al., 1993; Heath et al., 1996). The usual route of administration is IV or intratracheally at a dosage of 0.05 to 0.5 mg/kg. Intracardiac injection has been performed in anesthetic emergencies, but successful resumption of a normal ECG is rare. Rapidly dilating pupils is a very alarming sign in anesthetized otariids and is indicative of a hypoxic episode. Pupil dilation should be addressed quickly by reducing inhalant anesthetic levels, reversal of anesthetic agents if possible, and ventilatory support. Some anesthetic agents may be more advantageous than others due to their reversibilty. The α2-agonists such as xylazine, detomidine, and medetomidine have been used successfully to anesthetize otariids (see Table 1) and are reversed by the agonists yohimbine and, more specifically, atipamezole. Benzodiazepine sedatives such as diazepam, midazolam, and zolazepam can be reversed by flumazenil (Karesh et al., 1997). If these anesthetic agents are used, the reversal agents should be kept on hand in the event of an anesthetic emergency. Cardiac pulmonary resuscitation (CPR) has been performed by the authors and others during anesthetic emergencies (Work et al., 1993; Heath et al., 1996). It has been minimally successful. Compression of the thorax seemed to result in movement of both air and blood, and reperfusion of oral membranes seemed to occur, based on color changes of the mucous membranes. However, the underlying problem often cannot be reversed. Prevention and planning helps in all emergencies.
Phocids In general, diving adaptations are more developed in phocids than otariids, resulting in increased diving performance and, potentially, more anesthetic complications for the veterinarian (Lynch et al., 1999b). As with otariids, a number of sedative agents can be used to facilitate physical restraint (Gales, 1989; Lynch et al., 1999b). Some recently used agents include diazepam at 0.12 to 0.2 mg/kg IV in Hawaiian monk seals (Monachus schauinslandi) (Braun, Ryon, and Antonelis, pers. comm.). Midazolam has been administered at 0.25 to 0.35 mg/kg IM in crabeater seals (Lobodon carcinophagus) (Lynch et al., 1999b), and at 0.15 to 0.3 mg/ kg IM in Hawaiian monk seals (Ryon et al., 1999). Butorphanol has been used at 0.05 to 0.1 mg/ kg IM in harbor seals (Phoca vitulina) as an aid during restraint (Haulena and Gulland, unpubl. data) and also at 0.4 mg/kg alone or in combination with diazepam (0.2 mg/kg) prior to endoscopic examination and muscle biopsies (Tuomi et al., 2000). A summary of recent phocid immobilization studies is included in Table 3. Because many of the aspects of otariid anesthesia also apply to phocids, this section will focus mainly on differences between these two pinniped groups.
Induction Atropine has been recommended to prevent bradycardia associated with the dive reflex, excessive salivation, and excessive upper respiratory tract secretion in anesthetized seals (Gales, 1989; Woods et al., 1989; 1994b; 1996a; Mitchell and Burton, 1991; Phelan and Green, 1992; Lynch et al., 1999b). Although many studies report administering atropine in the same syringe as the immobilizing agents under field conditions, the authors recommend 0.02 mg/kg atropine IM approximately 10 min prior to the administration of the anesthetic agent (Gulland et al., 1999). Some researchers do not recommend the administration of atropine to phocids prior to anesthesia (Woods, pers. comm.). Terbutaline has been reported in a single harbor seal to prevent bronchial spasm while not increasing heart rate (Moesker, 1989).
26
Mirounga leonina (southern elephant seal) 15
4
13
17
30
5
Mirounga angustirostris (northern elephant seal)
Lobodon carcinophagus (crabeater seal)
Leptonychotes weddellii (Weddell seal)
34
1
Ketamine, diazepam Ketamine, diazepam
Medetomidine, ketamine
Ketamine, halothane Tiletamine and zolazepam Ketamine, diazepam Midazolam, pethidine
Ketamine, xylazine Tiletamine and zolazepam Xylazine
Tiletamine and zolazepam
7
3
Tiletamine and zolazepam
44
Hydrurga leptonyx (leopard seal)
Ketamine, diazepam
271
Halichoerus grypus (gray seal)
Agent
n
Species
IM dart IM dart IM IM
IM IM
70–140 µg/kg 2.5 mg/kg 6 mg/kg 0.30 mg/kg 2.3–3.9 mg/kg 0.02–0.35 mg/kg
IM IM IM dart IM dart
IV IH IM
IM
IM IM IM
IM
IM dart
IM dart IM dart
Route
2.9–7.7 mg/kg 0.11–0.25 mg/kg 0.29–0.37 mg/kg 1.3–2.2 mg/kg
0.15–0.25 mg/kg 1–4% 0.3–1.1 mg/kg
0.8–2.8 mg/kg
2.2–5.9 mg/kg 0.2–0.5 mg/kg 2.0 mg/kg
0.5 mg/kg
1 mg/kg
6 mg/kg 0.3 mg/kg
Dosage
TABLE 3 Some Recently Published Chemical Immobilizing Agents Used in Phocids
0
0
50
8
6
10
0
12
100
67
0
0
2
Mortality, %
Additional midazolam and/or ketamine given, reversed with naloxone and flumazenil Bradycardia, prolonged recovery, poor reversibility, variable plane of anesthesia Additional ketamine required in some animals, long recovery Apnea, poor muscle relaxation
Plane of anesthesia varied with dosage
Variable plane of anesthesia with lower dose
Apnea, bradycardia
Additional ketamine and diazepam required in some animals, long recovery Some animals became apneic requiring artificial respiration, tremors noted Additional doses given to maintain sedation; palpebral reflex present at all times Muscle tremors, apnea, bradycardia
Comments
Marine Mammal Anesthesia
(Continued)
Woods et al., 1994b
Baker et al., 1988
Haulena and Gulland, unpubl. data
Lynch et al., 1999a
Phelan and Green, 1992 Shaughnessy, 1991
Mitchell and Burton, 1991 Mitchell and Burton, 1991 Mitchell and Burton, 1991 Hurford et al., 1996
Lawson et al., 1996
Baker et al., 1990
Baker et al., 1988
Reference
0839_frame_C29 Page 671 Tuesday, May 22, 2001 11:57 AM
671
Species Ketamine, diazepam Ketamine, diazepam
Ketamine, diazepam Ketamine, xylazine Ketamine, xylazine Ketamine, xylazine Ketamine, xylazine Ketamine, xylazine Medetomidine
15
27
Midazolam, pethidine
Midazolam, pethidine, thiopentone Midazolam, pethidine, ketamine
13
10
6
Medetomidine, ketamine
10
2
0.02–0.07 mg/kg 2.7–6.7 mg/kg 2.2–5.9 mg/kg 0.02–0.07 mg/kg 2.7–6.7 mg/kg 1.0–3.5 mg/kg
IM IM IV IM IM IV
IM IM
IM IM
12–27 µg/kg 1.4–2.2 mg/kg 0.02–0.07 mg/kg 1.2–6.7 mg/kg
IM IM IM IM IM IM IM IM IM IM IM IM IM
IM IM IV IV
Route
4.4–8.6 mg/kg 0.04–0.13 mg/kg 3.6–6.4 mg/kg 0.6–1.2 mg/kg 1.6–7.5 mg/kg 0.25–1.2 mg/kg 2.1–11.4 mg/kg 0.2–0.5 mg/kg 2.1–3.1 mg/kg 0.2–0.5 mg/kg 2.5–3.4 mg/kg 0.5–0.6 mg/kg 13–27 µg/kg
2.0–4.8 mg/kg 0.05–0.12 mg/kg 1.8–3.4 mg/kg 0.04–0.18 mg/kg
Dosage
0
0
0
0
0
0
0
Apnea; responsive to doxapram (2 mg/kg) Vomiting, hyperthermia, bradycardia Hyperthermia, variable plane of anesthesia, poor reversibility, bradycardia Deeper sedation with higher doses of pethidine (2.7–6.7 mg/kg), faster recovery after naloxone or naltrexone Good immobilization allowing intubation of 5 min duration after thiopentone Good immobilization allowing intubation of 5 min duration after ketamine
Apnea, tremors
Prolonged sedation in postlactation and postpartum females Prolonged apnea
4 4
Apnea noted
Apnea, faster induction, more predictable, and shorter immobilization with IV compared with IM Apnea
Apnea, poor muscle relaxation
Comments
0
0
0
0
Mortality, %
Woods et al., 1994a
Woods et al., 1994a
Woods et al., 1994a
Woods et al., 1996a
Woods et al., 1996a
Woods et al., 1996b
Mitchell and Burton, 1991 Woods et al., 1994b
Woods et al., 1989
Bester, 1988
Slip and Woods, 1996
Slip and Woods, 1996
Woods et al., 1994b
Reference
672
25
15
55
194
5
32
Agent
n
TABLE 3 Some Recently Published Chemical Immobilizing Agents Used in Phocids (continued)
0839_frame_C29 Page 672 Tuesday, May 22, 2001 11:57 AM
CRC Handbook of Marine Mammal Medicine
Phoca vitulina (harbor seal)
Tiletamine and zolazepam Tiletamine and zolazepam Tiletamine and zolazepam Tiletamine and zolazepam Ketamine, diazepam, nitrous oxide, ethrane Propofol Propofol, isoflurane
90
12
9
1
4
15
5
15
Midazolam, pethidine, ketamine Ketamine, midazolam
3
3–5 mg/kg 2–5%
IV IH
0
Optimum short-acting anesthesia at 5 mg/kg, apnea Easily intubated, apnea
Gulland et al., 1999
Gulland et al., 1999
Moesker, 1989
0
IM IM IH IH IV
6 mg/kg 0.2 mg/kg 66% 1–2% 2–6 mg/kg 0
Karesh et al., 1997
0
Mitchell and Burton, 1991 Woods et al., 1994b
Baker et al., 1990
Woods et al., 1994b
Woods et al., 1994a
IM
Apnea, tremors, possible hallucinations, prolonged recovery
Incremental doses (every 3–24 min) of ketamine to maintain immobilization for 1 h Deep sedation, apnea, prolonged recovery, hyperthermia, bradycardia Some animals became apneic requiring artificial respiration Prolonged apnea, muscle tremors
0.6–1.7 mg/kg
0
40
0
0
0
IM
IM
IM dart
IM IM IV IM IM
0.7–1.2 mg/kg
1.6–2.4 mg/kg
1 mg/kg
0.02–0.07 mg/kg 2.7–6.7 mg/kg 4.4–10.0 mg/kg/h 2.1–3.7 mg/kg 0.02–0.03 mg/kg
0839_frame_C29 Page 673 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia 673
0839_frame_C29 Page 674 Tuesday, May 22, 2001 11:57 AM
674
CRC Handbook of Marine Mammal Medicine
Sites for IM injection of anesthetic agents include muscle overlying the hips (Bester, 1988; Woods et al., 1989) and the posterior lumbar muscles (Baker et al., 1988; 1990; Phelan and Green, 1992; Woods et al., 1994a). Darting has been used in field studies (Baker et al., 1988; 1990; McCann et al., 1989). As with otariids, injection into blubber layers may result in variable induction, plane of anesthesia, and recovery (Baker and Gatesman, 1985; Gales, 1989). Baker et al. (1990) and Woods et al. (1994b) suggest that IM ZT is a superior anesthetic combination in terms of rapid induction, animal safety, rapid recovery, and injection volume in comparison to ketamine-xylazine and ketamine-diazepam when used in gray seals (Halichoerus grypus) and southern elephant seals (Mirounga leonina). However, Mitchell and Burton (1991) had much more variable results, including several mortalities when ZT was used in southern elephant seals and leopard seals (Hydrurga leptonyx). The higher dosages used in the latter study may indicate that ZT has a narrow margin of safety in some phocids. When insufficient levels of sedation have been achieved, some authors have given additional doses of the agents. Additional ketamine and diazepam have been given after initial ketamine-diazepam (Baker et al., 1988). Additional ketamine alone has been given after initial ketamine-diazepam (Baker et al., 1988) and after ketamine-xylazine (Mitchell and Burton, 1991). After initial IM injection of ZT, additional ZT IV or IM has been given (Baker et al., 1990; Mitchell and Burton, 1991; Lawson et al., 1996). Supplementary doses of midazolam and/or pethidine IM or IV have been administered after initial pethidine-midazolam IM (Woods et al., 1994a; Lynch et al., 1999a). In addition, incremental ketamine IV has been been given after initial pethidinemidazolam IM (Woods et al., 1994a). Mortalities have been associated with the administration of additional ZT (Mitchell and Burton, 1991) and pethidine-midazolam (Lynch et al., 1999a). The use of strong narcotic agents such as etorphine or carfentanil may cause prolonged apnea resulting in mortalities; thus, these agents must be used with caution (Baker and Gatesman, 1985; Gales, 1989). In contrast to otariids, phocids have a readily accessible intravertebral, epidural vein that can be accessed in the caudal lumbar region with minimal potential for damaging the spinal cord (see Chapter 19, Clinical Pathology). This large vessel is used to pool shunted blood during dives in phocids and can be used to administer anesthetic agents and emergency drugs by the IV route (Woods et al., 1989; 1994a; Phelan and Green, 1992; Slip and Woods, 1996; Hurford et al., 1996; Gulland et al., 1999). An alternative site is the metatarsal vascular plexus. However, due to the close approximation of veins and arteries, agents may be mistakenly introduced into an artery. Comparing IM and IV ketamine-diazepam, Slip and Woods (1996) found that the IV route resulted in faster induction, decreased length of immobilization, more predictable plane of anesthesia, and required less ketamine. The IV route may not be possible in all instances because of the requirement for increased restraint (Gales, 1989). Induction by mask inhalation has been accomplished in young animals such as harbor seal pups and northern elephant seal (Mirounga angustirostris) pups in a similar manner to that already discussed for otariids. However, older phocids are more difficult to induce than otariids because of their propensity to breath-hold, which prolongs the necessary restraint time, and thus older phocids usually require some injectable method of induction (Gales, 1989).
Intubation Many of the indications and methods for intubating immobilized phocids are similar to those already mentioned for otariids. Important differences include a longer trachea prior to bifurcation in phocids, allowing for placement of a longer and more secure endotracheal tube. Many phocids, especially larger or more mature animals, have a large amount of peripharyngeal tissue and a flaccid soft palate that tend to obscure the glottis (Phelan and Green, 1992) and occlude
0839_frame_C29 Page 675 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
675
the airway in immobilized seals (Lynch et al., 1999b). Manual insertion of the tube in smaller animals, such as recently weaned northern elephant seals, is difficult for most personnel unless they possess small hands. Laryngoscopes of regular blade width do not displace a sufficient amount of the tissue to allow visualization of the glottis. Placing the animal in dorsal recumbency may improve visualization, as does the use of a frosting spatula of approximately 5 to 8 cm in width placed under the larngyscope blade. The authors have also intubated juvenile elephant seals by using a laparoscope as a stylette inside the endotracheal tube during placement. Blind techniques for inserting endotracheal tubes have also been used (Woods et al., 1994a). Several studies have immobilized phocids to conduct stomach lavage to investigate forage items (Antonelis et al., 1987; Mitchell and Burton, 1991; Slip and Woods, 1996). Mortalities in some of these studies have been associated with aspiration of stomach contents (Mitchell and Burton, 1991); thus, endotracheal intubation is especially recommended for these studies.
Inhalation Anesthesia Although sedation and some degree of immobilization have been successfully accomplished using the IM and IV routes (see Table 3), most of those studies did not require a surgical plane of anesthesia. For procedures of a longer duration, or for those requiring a surgical plane of anesthesia, animals usually require additional anesthesia that may be most safely provided by inhalation route. Recently reported inhalation anesthetics used in phocids have included ethrane (Moesker, 1989), halothane (Hurford et al., 1996), and isoflurane (Gulland et al., 1999).
Monitoring Respiratory rate has been monitored by observing thoracic movements (Mitchell and Burton, 1991; Woods et al., 1994b). Heart rate has been monitored by recording observable cardiac movements through the body wall and by direct stethoscopic auscultation (Gulland et al., 1999). Depth of anesthesia has been assessed in much the same way as for otariids (Mitchell and Burton, 1991; Woods et al., 1994b; 1996a; Slip and Woods, 1996). Pulse oximetry has been used by placing clip probes on the tongue and reflectance probes rectally (Gulland et al., 1999). Alternative sites have included nasal septum, vaginal mucosa, and buccal mucosa. Temperature in phocids has been monitored rectally (Mitchell and Burton, 1991; Phelan and Green, 1992; Woods et al., 1994b) and esophageally. Capnometers and blood pressure monitors have been used in harbor seals and elephant seals, as has already been described for otariids (Haulena, unpubl. data). Venous blood gases have been measured from the epidural vein (Woods et al., 1994a,b; 1996a). Arteries have been cannulated in the front flipper for possible arterial blood gas monitoring (Hurford et al., 1996).
Support Individual animals require different amounts of support as dictated by monitoring. Preanesthetic condition can have significant effects on duration, depth, and recovery from anesthesia (Woods et al., 1989). This may be especially important in phocids, which have a large seasonal change in body fat composition with breeding, pupping, and molting cycles. Unlike otariids, the epidural vein can be catheterized to provide fluids or other agents during anesthesia to maintain hydration and adequate circulation (Hurford et al., 1996). Hypothermia was associated with ZT administration in a Weddell seal (Leptonychotes weddellii) and was corrected by covering the animal with windproof material (Phelan and Green, 1992). Hypothermia is also commonly encountered when using isoflurane alone in recently
0839_frame_C29 Page 676 Tuesday, May 22, 2001 11:57 AM
676
CRC Handbook of Marine Mammal Medicine
weaned harbor seal pups and may prolong recovery from anesthesia (Haulena, unpubl. data). Hyperthermia has been reported during the use of ketamine-xylazine (Woods et al., 1994b) and medetomidine-ketamine (Woods et al., 1996a) in southern elephant seals. Muscular tremors are often observed in phocids given cyclohexamine drug combinations (see Table 3). Bradycardia (<50 bpm) was noted in elephant seals following ketamine-xylazine as well as medetomidine-ketamine, and in leopard seals given ZT and ketamine-xylazine (Mitchell and Burton, 1991; Woods et al., 1996a). Apnea and hypoventilation are very common in anesthetized phocids (see Table 3). Some authors report a transient period of apnea occurring shortly after administration of an anesthetic agent (Woods et al., 1994b; Gulland et al., 1999). Slip and Woods (1996) calculated the theoretical aerobic dive limit according to the formula by Kooyman (1989) for southern elephant seals that they had anesthetized. They found that, in most cases, duration of apnea did not exceed the aerobic dive limit. However, other studies have reported mortalities associated with prolonged apnea (see Table 3). Slip and Woods (1996) suggest that apnea approaching the aerobic dive limit be treated with emergency procedures (see below). Inherent ventilation effort seems to be less in phocids compared with otariids during inhalation anesthesia, and the need for ventilatory assistance for phocids must be planned in advance. Mechanical ventilators may be necessary, especially if longer periods of anesthesia are needed. Simple periodic hand assistance is usually not enough to overcome the hypoventilation. Hypoventilation may not be as important during short, strictly injectable regimes such as with diazepam-ketamine (Spraker, pers. comm.). During anesthetic management of all marine mammals, it is important to monitor respiratory effort of the patient. Phelan and Green (1992) noted that respiratory effort was greater in Weddell seals in sternal recumbency compared with those placed in lateral recumbency that breathed more easily. Those authors found it easier to artificially ventilate an animal in lateral rather than sternal recumbency. The authors also attributed a mortality to respiratory obstruction predisposed to by upper respiratory anatomy and the potential for tracheal collapse. Respiratory obstruction has also been noted during the use of ketamine-diazepam in elephant seals (Woods et al., 1994b).
Emergencies In general, anesthetic emergencies in phocids are handled in much the same way as those in otariids. Clinical signs noted in animals that have died as a result of immobilization have included tachycardia, bradycardia, cyanosis, hypoventilation, decreased peripheral perfusion, and hyperthermia (Woods et al., 1989; Mitchell and Burton, 1991; Phelan and Green, 1992). Intravascular doxapram has been shown to help in cases of immobilization-related apnea (Baker et al., 1988; Phelan and Green, 1992). It also acts as a general stimulant and dosedependent antagonist for several anesthetic agents (Woods et al., 1995; 1996b). Woods et al. (1996b) found that doxapram was most beneficial if introduced via an endotracheal tube directly into the lungs. Those authors recommended a dosage of 2 mg/kg to stimulate respiration and 4 mg/kg as a general stimulant and antagonist. The underside of the tongue is also a useful site for injection (Baker and Gatesman, 1985; Bester, 1988; Woods et al., 1996b). As with otariids, the anesthetist should be prepared to intubate and provide ventilatory assistance to animals with prolonged apnea. In some animals with prolonged apnea, the larynx is tightly closed requiring forced opening prior to insertion of the endotracheal tube (Baker et al., 1990). Some authors have breathed directly into the endotracheal tube to inflate the lungs of an apneic seal (Baker et al., 1990). However, this method may not be ideal as a result of the potential presence of zoonotic pathogens, and the use of an alternative method such as a mechanical ventilator should be considered, if possible. In some instances, intubation itself resulted in
0839_frame_C29 Page 677 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
677
apnea, and the presence of an endotracheal tube may hinder an anesthetized southern elephant seal from breathing (Woods et al., 1996b). Prolonged apnea often precedes development of bradycardia prior to death in immobilized phocids, as was noted in animals given ZT that were unresponsive to IV doxapram (Phelan and Green, 1992). The authors of that study suggest that doxapram be administered prior to the beginning of cardiovascular compromise to be effective. In cases where cardiac function has already decreased, the authors recommended intracardiac injection of doxapram to ensure adequate concentrations at receptor sites, although intracardiac injections in large animals under field conditions may be difficult. Thoracic compressions to facilitate cardiac contractions have been performed in phocids but, as with otariids, have not often been successful (Mitchell and Burton, 1991). The use of reversible agents may be advantageous in phocids, but cannot be relied upon in emergency situations. Recent reversible agents that have been evaluated in phocids include naloxone and naltrexone to reverse pethidine (Woods et al., 1994a; Lynch et al., 1999a), 4aminopyridine to reverse (partially) ketamine-diazepam (Woods et al., 1995), sarmazenil to reverse (partially) ketamine-diazepam and ZT (Woods et al., 1995), yohimbine to reverse (partially) ketamine-xylazine (Woods et al., 1995), and flumazenil to reverse midazolam (Lynch et al., 1999a; Ryon et al., 1999). Medetomidine-ketamine in elephant seals was poorly reversed using atipamezole, resulting in prolonged recovery, bradycardia, and mortality (Woods et al., 1996a; Haulena and Gulland, unpubl. data).
Odobenids In comparison with phocids and otariids, relatively little information is available on anesthesia of walruses (Odobenus rosmarus) (Gales, 1989). The most common sedative agents include meperidine with diazepam and meperidine with midazolam delivered by the IM route (Walsh, pers. comm.). Table 4 summarizes the more recent anesthesia literature.
Induction Like phocids, odobenids have a large intravertebral epidural sinus that can be safely accessed for venipuncture via the intervertebral spaces of L4 to L7 (Sweeney, 1993). However, because of their large size, the use of IV induction in walruses is usually limited to well-trained individuals in display facilities, or very young animals that can be easily restrained. Intravenous thiopental has been used in juvenile and subadult walruses after premedication with midazolam and meperidine (Gage, pers. comm.; Walsh, pers. comm.). The IM route, however, is necessary in many situations. Free-ranging walruses have been immobilized with a number of chemical agents delivered by either capture rifle (DeMaster et al., 1981; Griffiths et al., 1993; Tuomi et al., 1996; Lanthier et al., 1999) or crossbow (DeMaster et al., 1981). Jabsticks and modified jabsticks that allow for injection of agents from several meters away have also been employed (Stirling and Sjare, 1988; Griffiths et al., 1993). Sites for IM injection have included the tongue, neck, paralumbar muscles, shoulder (Griffiths et al., 1993; Lanthier et al., 1999), and the muscle overlying the hip, where blubber layers are relatively thin but still necessitate the use of needle lengths of 8 to 11 cm (Stirling and Sjare, 1988; Griffiths et al., 1993; Tuomi et al., 1996; Lanthier et al., 1999) in adult animals. If possible, hand injection of IM agents is a more reliable method of administration (Walsh, pers. comm.). DeMaster et al. (1981) investigated, and Gales (1989) summarized, many of the phencyclidine, ketamine, and acepromazine combinations that have been used. Immobilization with ZT
Meperidine
Meperidine, thiopental Etorphine, medetomidine, ketamine
23
na
1
38
3
Tiletamine and zolazepam Tiletamine and zolazepam Etorphine
10
Agent
0.22–0.45 mg/kg 0.74 mg/kg 5.2 µg/kg 83 µg/kg 1 mg/kg
IM IV IM dart IM IM
IM
IM dart
3.3–8 µg/kg
0.23–0.45 mg/kg
IM
IM
Route
1.4–2.2 mg/kg
0.6–2.25 mg/kg
Dosage
0
na
0
3
33
10
Mortality, %
Best results were found when using 2.0–2.25 mg/kg; prolonged recovery Smooth induction and recovery; animal given the highest dosage died Reversed with diprenorphine (15–18 mg for 10 mg of etorphine); convulsions and apnea noted Moderate sedation with moderate respiratory depression; several hours duration; reversed with naloxone (3.9–8.8 µg/kg) IV Maintained surgical plane with thiopental administered as required Etorphine reversed with diprenorphine prior to medetomidine/ketamine; medetomidine reversed with yohimbine (156 µg/kg IM); sedated for up to 5 h
Comments
Lydersen et al., 1992
Cornell and Antrim, 1987
Joseph and Cornell, 1988
Griffiths et al., 1993
Griffiths et al., 1993
Stirling and Sjare, 1988
Reference
678
n
TABLE 4 Some Recently Published Chemical Immobilizing Agents Used in Odobenids
0839_frame_C29 Page 678 Tuesday, May 22, 2001 11:57 AM
CRC Handbook of Marine Mammal Medicine
3
4
Meperidine, midazolam, thiopental, isoflurane Meperidine, midazolam, thiopental, isoflurane
Carfentanil, isoflurane Carfentanil
4
6
Carfentanil
13
IM IH IM dart
IM IM IV IH IM IM IV IH
2 mg/kg 0.03–0.05 mg/kg To affect 1–2% 1.1–2.3 mg/kg 0.11 mg/kg 2.2–4.0 mg/kg 0–3%
IM
2.4–2.7 mg/animal 3–5% 3.4–5.4 µg/kg
2.4–2.7 mg/animal
0
0
0
0
0
Flumazenil used at end of procedure
Adult male; muscular rigidity made intubation difficult; given 175–350 mg naltrexone IM as soon as could be approached and additional 175–350 mg upon completion of procedure As above; intubated after initial naltrexone given to relax jaw muscles Muscle spasms; apnea within 6–13 min; respiration within 3–7 min after administration of naltrexone (150–250 mg/mg carfentanil) Thiopental given 20–30 min after meperidine/midazolam; naloxone administered to speed recovery Gage, pers. comm., 2000
Walsh et al., 1988
Lanthier et al., 1999
Tuomi et al., 1996
Tuomi et al., 1996
0839_frame_C29 Page 679 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia 679
0839_frame_C29 Page 680 Tuesday, May 22, 2001 11:57 AM
680
CRC Handbook of Marine Mammal Medicine
seemed to result in fewer side effects than those combinations and, in contrast to the use of ZT in otariids, additional amounts of ZT have been given to walrus to prolong anesthesia or increase depth of anesthesia without detrimental effects (Stirling and Sjare, 1988).
Intubation and Inhalation Anesthesia Intubation of juvenile walruses has been accomplished by manual insertion of an endotracheal tube, much as has been described for large otariids. Animals weighing between 400 and 800 kg have been intubated using 16- to 22-mm-diameter tubes (Walsh et al., 1988; Gage, pers. comm.). Isoflurane has been used in display animals to maintain anesthesia after induction and intubation (Walsh et al., 1988; Gage, pers. comm.).
Monitoring As with otariids, respiratory and cardiac rates have been monitored by noting thoracic movements (Griffiths et al., 1993; Tuomi et al., 1996) and heart rate has also been monitored via ECG (Walsh et al., 1988). Resting values in unanesthetized male walruses were approximately four to eight breaths/min (Stirling and Sjare, 1988) while heart rate was found to be 52 to 66 bpm (Griffiths et al., 1993). Temperature has been recorded by placing a probe approximately 35 to 60 cm into the rectum (Walsh et al., 1988; Griffiths et al., 1993; Lanthier et al., 1999). Recorded temperatures have varied from 34.8 to 37.9°C in walruses immobilized with carfentanil (Lanthier et al., 1999). Attempts have been made to use pulse oximetry in walruses. Probes attached to nasal septum, tongue, prepuce, anal mucosa, and cheek were ineffective at recording saturation reliably (Lanthier et al., 1999). Corneal transducers have been most effective (Walsh, pers. comm.). Blood gases and EtCO2 have been monitored in anesthetized walruses (Walsh, pers. comm.).
Support Hyperthermia was noted during the use of phencyclidine and was controlled by pouring cold water over immobilized animals (DeMaster et al., 1981). No adverse effects on core temperatures were seen with ZT (Stirling and Sjare, 1988), carfentanil (Lanthier et al., 1999), or etorphine (Griffiths et al., 1993). Cardiac arrhythmias have been reported after multiple dosages of sedative agents (Walsh, pers. comm.). Apnea was a consistent finding when etorphine was used to immobilize walruses (Griffiths et al., 1993). Most of the animals in that study would breathe only after the injection of the reversal agent. Respiratory depression and hypoxia may occur when meperidine is used, especially in combination with barbiturates (Walsh, pers. comm.). Doxapram has been used to stimulate respiration in apneic walruses (Cornell and Antrim, 1987) and a dosage of 0.5 mg/kg IV has been recommended for barbirurate-induced apnea (Walsh, pers. comm.). Large animal ventilators have been used in walruses (Walsh et al., 1988). Poor positioning of immobilized animals may result in problems during anesthesia. DeMaster et al. (1981) recommended that animals not be immobilized on a slope while their heads are lower than their bodies. This may cause excessive compression of the thorax by the weight of the abdominal organs and interfere with adequate pulmonary expansion. Occlusion of the airway has occurred when a nearby animal blocked the nasal openings of an immobilized walrus that could not reposition itself (Stirling and Sjare, 1988). It was also found that the tongue would fall onto the palate and block the airway if animals were placed supine (Griffiths et al., 1993). If possible, animals should be intubated to maintain an adequate airway and tubes should remain in place for as long as possible after recovery has begun (Walsh, pers. comm.).
0839_frame_C29 Page 681 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
681
Darted animals may end up entering the water and drowning if they are not selected carefully (Lanthier et al., 1999).
Emergencies Doxapram has been injected IM into the neck muscles to prevent apnea but may not be effective (Griffiths et al., 1993; Lanthier et al., 1999). Doxapram is best used IV, and can be administered into the intravertebral sinus. Etorphine is reversible by the antagonist diprenorphine (Lydersen et al., 1992), which can be administered IV but has been injected IM because of poor accessibilty to the venous sinus and tongue under field conditions (Griffiths et al., 1993). Reversal appears slower when diprenorphine is given IM. Diprenorphine did not result in complete reversal, and immobilized animals appeared lethargic for up to 1 day after the immobilization. Moderate sedation by meperidine has been reversed within 2 min of the administration of IV naloxone (Joseph and Cornell, 1988). Yohimbine has been used to reverse medetomidine (Lydersen et al., 1992). Apnea that was noted after injection of carfentanil was postulated to be the result of muscular spasms of the upper airway and resolved after administration of the antagonist naltrexone given either IM into tongue, lips, or shoulder or IV (Tuomi et al., 1996; Lanthier et al., 1999).
Sirenians Relatively little is known about the anesthesia of sirenians. A recent publication (Walsh and Bossart, 1999) describes chemical restraint of the Florida manatee (Trichechus manatus latirostris). Midazolam (0.045 to 0.08 mg/kg IM) and diazepam (0.066 mg/kg) have been used to facilitate restraint in manatees. Meperidine has been used at up to 1 mg/kg along with midazolam and diazepam to increase sedation. Intubation has been accomplished transnasally by visualizing the larynx endoscopically through one nares while introducing the endotracheal tube through the other nasal opening. Isoflurane has been used to maintain anesthesia. Mechanical ventilation and the use of a large reservoir system is recommended. Antagonism of meperidine with naloxone and diazepam or midazolam with flumazenil (IM) has been perfomed. Doxapram has been used to stimulate respiration in manatees. The reader is referred to Chapter 43, Manatees, for a more complete discussion of manatee restraint and immobilization.
Sea Otters There are relatively few anesthetic agents routinely used in sea otters (Enhydra lutris) (Williams et al., 1990). Several agents that were tried during the Exxon Valdez oil spill in Alaska in 1989 have been recently reviewed. Best results were obtained with combinations of fentanyl-diazepam with either azaperone or acepromazine (Sawyer and Williams, 1996). Table 5 is a summary of some anesthetic agents that have been used in sea otters.
Induction Since sea otters do not have a blubber layer, vessels such as the jugular vein can be palpated for IV access, especially after the coat overlying it has been wetted down with alcohol. Sufficient restraint may be difficult to attain, especially in larger animals. The femoral and popliteal veins have been used for venipuncture and may be used for administering anesthetic agents (see Chapter 19, Clinical Pathology).
Fentanyl, acepromazine, diazepam Fentanyl, diazepam Fentanyl, azaperone, diazepam Fentanyl, acepromazine, diazepam Meperidine, diazepam
na
57
32
0.2 mg/kg 0.05 mg/kg 0.5 mg/kg 0.1 ± 0.003 mg/kg 0.1 ± 0.006 mg/kg 0.1 ± 0.02 mg/kg 0.5 ± 0.02 mg/kg 0.3 ± 0.01 mg/kg 0.1 ± 0.006 mg/kg 0.14 ± 0.01 mg/kg 0.2 ± 0.01 mg/kg 13 ± 0.5 mg/kg 0.2 ± 0.01 mg/kg
0.05 mg/kg 0.2 mg/kg 11–13.2 mg/kg 0.22–0.55 mg/kg
Dosage
IM IM IM IM IM IM IM IM
IM IM IM IM
IM IM IM IM
Route
na
na
na
na
na
na
na
Mortality
Numerous mortalities but many attributed to other factors
Deeper sedation than fentanyl/diazepam only and duration of up to 2.5 h
Lighter sedation and shorter acting than when either acepromazine or azaperone was added Deeper sedation than fentanyl/diazepam only and duration of up to 2.5 h
Best combination of several agents that were tested including meperidine, diazepam, nalbuphine, azaperone, fentanyl, and ketamine Recommended during Exxon Valdez oil spill (1989)
Recommended for field use
Comments
Sawyer and Williams, 1996
Sawyer and Williams, 1996
Sawyer and Williams, 1996
Sawyer and Williams, 1996
Williams and Sawyer, 1995
Joseph et al., 1987
Williams, 1986
Reference
682
61
294
na
Fentanyl, azaperone Meperidine, diazepam
Agent
na
n
TABLE 5 Some Recently Published Chemical Immobilizing Agents Used in Sea Otters (Enhydra lutris)
0839_frame_C29 Page 682 Tuesday, May 22, 2001 11:57 AM
CRC Handbook of Marine Mammal Medicine
0839_frame_C29 Page 683 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
683
Because of the difficulty associated with adequately restraining larger animals for IV injection, the IM route is more often employed, especially under field conditions (Sawyer and Williams, 1996). Sites for IM injection of anesthetic agents include the quadriceps, semimembrinosus-semitendinosus (Joseph et al., 1987), and lumbar muscles. The most common agents currently used for free-ranging sea otters include a combination of fentanyl (0.22 to 0.4 mg/kg IM) and diazepam (0.07 to 0.1 mg/kg IM) that is partially reversible using naltrexone at twice the fentanyl dose (Monson, pers. comm.). Ketamine was thought to have a narrow margin of safety in sea otters in comparison with its use in other mustelids (Williams and Kocher, 1978). Meperidine alone may cause diaphragmatic spasm, convulsions, and respiratory depression, side effects that were not seen when diazepam was added (Joseph et al., 1987). The use of atropine prior to anesthesia has not been reported in sea otters. Physical restraint is possible, especially in smaller individuals, to allow for induction by inhalation (Williams et al., 1990). Oxymorphone (0.3 mg/kg) has been combined with diazepam (0.5 mg/kg) IM (Huff, pers. comm.) and medetomidine with butorphanol IM (Murray, pers. comm.) to aid in restraint for induction using isoflurane. Isoflurane alone has been used as an induction agent in young sea otters.
Intubation The glottis and laryngeal folds are easily visualized in sea otters with the aid of a standard laryngoscope. However, the endotracheal tube diameter that can be passed into the trachea is often smaller than that for terrestrial carnivores of a similar mass.
Inhalation Anesthesia Isoflurane appears to be well tolerated by sea otters of varying age class and disease status. However, the use of inhalant anesthetics was contraindicated during cleaning of oiled sea otters due to potential volatilization of the petroleum products and exacerbation of pulmonary lesions (Williams et al., 1995; Sawyer and Williams, 1996). Halothane and nitrous oxide have also been used (Williams, 1986).
Monitoring Stethoscopic monitoring of respiratory and cardiac rate and character has been extensively used during anesthesia of sea otters. Pulse oximetry has been employed in sea otters by attaching clip probes to the tongue or genital mucosa. Reflectance probes can be used rectally. Anesthetized sea otters appear to have difficulty thermoregulating. Preexisting conditions that have affected the hair coat, body condition, or metabolic rate can quickly exacerbate thermoregulatory difficulties, and monitoring temperature is vital. Flexible temperature probes can be placed rectally. Doppler flow probes have been placed on the forelimbs to evaluate pulse strength. The use of capnography, ECG recording, and blood pressure monitoring have not been well described in sea otters.
Support Responding to changes in the temperature of anesthetized sea otters is extremely important. Dealing with hyper- or hypothermia is much the same as has already been described for pinnipeds. An easily palpable jugular vein in anesthetized sea otters can be catheterized for continuous intravenous fluid administration.
0839_frame_C29 Page 684 Tuesday, May 22, 2001 11:57 AM
684
CRC Handbook of Marine Mammal Medicine
Emergencies Although the use of emergency drugs has not been well described in sea otters, many of the agents used in terrestrial carnivores can be used in sea otters. The anesthetist should be well versed in the use of emergency anesthetic agents. Reversal agents including naloxone (Joseph et al., 1987) and naltrexone have been used to antagonize the effects of various narcotic agents.
Ursids The anesthesia of polar bears (Ursus maritimus) is more typical of terrestrial carnivores than of other marine mammals. Polar bears are highly intelligent, agile, and dangerous animals and pose a serious risk to personnel required to work with them. Specialized training and an experienced team approach are essential. Chapter 45, Polar Bears, discusses polar bear immobilization in detail.
Conclusion When anesthetizing marine mammals, planning is vital and training is more important. With new drugs and technologies come new responsibilities for knowledge and experience. This chapter has attempted to review the more recent literature pertaining to anesthesia of marine mammals. Consultation with experts may be necessary to understand the details and to develop an adequate protocol for each specific situation. New techniques and equipment are available for making complex procedures possible even in remote field situations. However, it may take more planning, personnel, and funding to take advantage of these methods.
Acknowledgments Many people offered their knowledge, experience, and insight during the completion of this chapter. Special thanks go to Rebecca Duerr, Laurie Gage, Frances Gulland, Bill Horne, Jim McBain, Mike Murray, Aleksija Neimanis, Sam Ridgway, Pam Tuomi, Mike Walsh, Rupert Woods, and Skip Young for their comments and review.
References Antonelis, G.A., Jr., Lowry, M.S., DeMaster, D.P., and Fiscus, C.H., 1987, Assessing northern elephant seal feeding habits by stomach lavage, Mar. Mammal Sci., 3: 308–322. Baker, J.R., and Gatesman, T.J., 1985, Use of carfentanil and a ketamine-xylazine mixture to immobilise wild grey seals Halichoerus grypus, Vet. Rec., 116: 208–210. Baker, J.R., Anderson, S.S., and Fedak, M.A., 1988, The use of a ketamine-diazepam mixture to immobilise wild grey seals (Halichoerus grypus) and southern elephant seals (Mirounga leonina), Vet. Rec., 123: 287–289. Baker, J.R., Fedak, M.A., Anderson, S.S., Arnbom, T., and Baker, R., 1990, Use of a tiletamine-zolazepam mixture to immobilise wild grey seals and southern elephant seals, Vet. Rec., 126: 75–77. Bester, M.N., 1988, Chemical restraint of Antarctic fur seals and southern elephant seals, S. Afr. J. Wildl. Res., 18: 57–60. Boyd, I.L., Lunn, N.J., Duck, C.D., and Barton, T., 1990, Response of Antarctic fur seals to immobilization with ketamine, a ketamine-diazepam or ketamine-xylazine mixture, and Zoletil, Mar. Mammal Sci., 6: 135–145. Cornell, L.H., and Antrim, J.E., 1987, Anesthesia and tusk extraction in walrus, J. Zoo Anim. Med., 18: 3–6.
0839_frame_C29 Page 685 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
685
David, J.H.M., Hofmeyr, J.M., Best, P.B., Meyer, M.A., and Shaughnessy, P.D., 1988, Chemical immobilization of free-ranging South African (Cape) fur seals, S. Afr. J. Wildl. Res., 18: 154–156. DeMaster, D.P., Faro, J.B., Estes, J.A., Taggart, J., and Zabel, C., 1981, Drug immobilization of walrus (Odobenus rosmarus), Can. J. Fish. Aquat. Sci., 38: 365–367. Dover, S.R., Beusse, D., Walsh, M.T., McBain, J.F., and Ridgway, S.H., 1999, Laparoscopic techniques for the bottlenose dolphin (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, Boston, 128–129. Elsner, R., 1999, Living in water: solutions to physiological problems, in Biology of Marine Mammals, Reynolds III, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 73–116. Ferreira, S.M., and Bester, M.N., 1999, Chemical immobilization, physical restraint and stomach lavaging of fur seals at Marion Island, S. Afr. J. Wildl. Res., 29: 55–61. Gage, L.J., 1993, Pinniped anesthesia, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 412–413. Gales, N.J., 1989, Chemical restraint and anesthesia of pinnipeds: A review, Mar. Mammal Sci., 5: 228–256. Gales, N.J., and Burton, H.R., 1988, Use of emetics and anaesthesia for assessment of Weddell seals, Aust. Wildl. Res., 15: 423–433. Gales, N.J., and Mattlin, R.H., 1998, Fast, safe, field-portable gas anesthesia for otariids, Mar. Mammal Sci., 14: 355–361. Geraci, J.R, 1973, An appraisal of ketamine as an immobilizing agent in wild and captive pinnipeds, J. Am. Vet. Med. Assoc., 163: 574–577. Greenwood, A.G., Taylor, D.C., and Wild, D., 1978, Fibreoptic gastroscopy in dolphins, Vet. Rec., 102: 495–497. Griffiths, D., Wiig, Ø., and Gjertz, I., 1993, Immobilization of walrus with etorphine hydrochloride and Zoletil®, Mar. Mammal Sci., 9: 250–257. Gulland, F.M.D., Haulena, M., Elliott, S., Thornton, S., and Gage, L., 1999, Anesthesia of juvenile Pacific harbor seals using propofol alone and in combination with isoflurane, Mar. Mammal Sci., 15: 234–238. Hammond, D., and Elsner, R., 1977, Anaesthesia in phocid seals, J. Zoo Anim. Med., 8: 7–13. Haulena, M., and Gulland, F.M.D., 2001, Use of medetomidine-zolazepam-tiletamine with and without atipamezole reversal to immobilize captive California sea lions, J. Wildl. Dis., 37: 566–573. Haulena, M., Gulland, F.M.D., Calkins, D.G., and Spraker, T.R., 2000, Immobilization of California sea lions using medetomidine plus ketamine with and without isoflurane and reversal with atipamezole, J. Wildl. Dis., 36: 124–130. Hawkins, E.C., Townsend, F.I., Lewbart, G.A., Stamper, M.A., Thayer, V.G., and Rhinehart, H.L., 1997, Bronchoalveolar lavage in a dolphin, J. Am. Vet. Med. Assoc., 211: 901–904. Heard, D.J., and Beusse, D.O., 1993, Combination detomidine, ketamine, and isoflurane anesthesia in California sea lions (Zalophus californianus), J. Zoo Wildl. Med., 24: 168–170. Heath, R.B., Calkins, D., McAllister, D., Taylor, W., and Spraker, T., 1996, Telazol and isoflurane field anesthesia in free-ranging Steller’s sea lions (Eumetopias jubatus), J. Zoo Wildl. Med., 27: 35–43. Heath, R.B., DeLong, R., Jameson, V., Bradley, D., and Spraker, T., 1997, Isoflurane anesthesia in free ranging sea lion pups, J. Wildl. Dis., 33: 206–210. Hurford, W.E., Hochachka, P.W., Schneider, R.C., Guyton, G.P., Stanek, K.S., Zapol, D.G., Liggins, G.C., and Zapol, W.M., 1996, Splenic contraction, catecholamine release, and blood volume redistribution during diving in the Weddell seal, J. Appl. Physiol., 80: 298–306. Joseph, B.E., and Cornell, L.H., 1988, The use of meperidine hydrochloride for chemical restraint in certain cetaceans and pinnipeds, J. Wildl. Dis., 24: 691–694. Joseph, B.E., Cornell, L.H., and Williams, T., 1987, Chemical sedation of sea otters, Enhydra lutris, J. Zoo Anim. Med., 18: 7–13. Karesh, W.B., Cook, R.A., Stetter, M., Uhart, M.M., Hoogesteijn, A., Lewis, M.N., Campagna, C., Mailuf, P., Torres, A., House, C., Thomas, L., Braselton, W.E., Dierenfield, E.S., McNamara, T.S., Duignan, P., Raverty, S., and Linn, M., 1997, South American pinnipeds: Immobilization, telemetry, and health evaluations, in Proceedings of the American Association of Zoo Veterinarians, Houston, TX, 291–295.
0839_frame_C29 Page 686 Tuesday, May 22, 2001 11:57 AM
686
CRC Handbook of Marine Mammal Medicine
Kooyman, G.L., 1981, Weddell Seal Consummate Diver, Cambridge University Press, Cambridge, 135 pp. Kooyman, G.L., 1989, Diverse Divers: Physiology and Behavior, Springer-Verlag, Berlin, 200 pp. Kooyman, G.L., Castellini, M.A., and Davis, R.W., 1981, Physiology of diving in marine mammals, Annu. Rev. Physiol., 43: 343–356. Lanthier, C., Stewart, R.E.A., and Born, E.W., 1999, Reversible anesthesia of Atlantic walruses (Odobenus rosmarus rosmarus) with carfentanil antagonized with naltrexone, Mar. Mammal Sci., 15: 241–249. Lawson, J.W., Parsons, J.L., Craig, S.J., Eddington, J.D., and Kimmins, W.C., 1996, Use of electroejaculation to collect semen samples from wild seals, J. Am. Vet. Med. Assoc., 209: 1615–1617. Linnehan, R.M., and MacMillan, A.D., 1991, Propofol/isoflurane anesthesia and debridement of a corneal ulcer in an Atlantic bottlenosed dolphin (Tursiops truncatus), in Proceedings of the American Association of Zoo Veterinarians, Calgary, 290–291. Loughlin, T.R., and Spraker, T., 1989, Use of Telazol to immobilize female northern sea lions (Eumetopias jubatus) in Alaska, J. Wildl. Dis., 25: 353–358. Lydersen, C., Griffiths, D., Gjertz, I., and Wiig, Ø., 1992, A tritiated water experiment on a male Atlantic walrus (Odobenus rosmarus rosmarus), Mar. Mammal Sci., 8: 418–420. Lynch, M.J., Oosterhaus, A.D.M.E., Cousins, D.V., Selleck, P., and Williams, P., 1999a, Anesthesia, hematology and disease investigation of free-ranging crabeater seals (Lobodon carcinophagus), in Proceedings of the American Association of Zoo Veterinarians, 41–42. Lynch, M.J., Tahmindjis, M.A., and Gardner, H., 1999b, Immobilisation of pinniped species, Aust. Vet. J., 77: 181–185. McCann, T.S., Fedak, M.A., and Harwood, J., 1989, Parental investment in southern elephant seals, Mirounga leonina, Behav. Ecol. Sociobiol., 25: 81–87. McCormick, J.G., 1969, Relationship of sleep, respiration, and anesthesia in the porpoise: A preliminary report, Proc. Nat. Acad. Sci. U.S.A., 62: 697–703. McGrath, C.J., Feeney, D., Crimi, A.J., and Ruff, J., 1981, Upper airway of the California sea lion: An anesthetist’s perspective, Vet. Med. Small Anim. Clin., 76: 548–549. Medway, W., McCormick, J.G., Ridgway, S.H., and Crump, J.F., 1970, Effects of prolonged halothane anesthesia on some cetaceans, J. Am. Vet. Med. Assoc., 157: 576–582. Mitchell, P.J., and Burton, H.R., 1991, Immobilisation of southern elephant seals and leopard seals with cyclohexamine anaesthetics and xylazine, Vet. Rec., 129: 332–336. Moesker, A., 1989, General anesthesia in a case of laparotomy on a harbour seal (Phoca vitulina), Aquat. Mammals, 15: 46–48. Nagel, E.L., Morgane, P.J., and McFarland, W.L., 1964, Anesthesia for the bottlenose dolphin, Tursiops truncatus, Science, 146: 1591–1593. Phelan, J.R., and Green, K., 1992, Chemical restraint of Weddell seals (Leptonychotes weddellii) with a combination of tiletamine and zolazepam, J. Wildl. Dis., 28: 230–235. Reed, J.Z., Chambers, C., and Fedak, M.A., 1994, Gas exchange of captive freely diving grey seals, Halichoerus grypus, J. Exp. Biol., 191: 1–18. Reidarson, T.H., Harrell, J.H., Rinaldi, M.G., and McBain, J., 1998, Bronchoscopic and serologic diagnosis of Aspergillus fumigatus pulmonary infection in a bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 451–455. Ridgway, S.H., and McCormick, J.G., 1967, Anesthetization of porpoises for major surgery, Science, 158: 510–512. Ridgway, S.H., and McCormick, J.G., 1971, Anesthesia of the porpoise, in Textbook of Veterinary Anesthesia, Soma, L.A. (Ed.), Williams & Wilkins, Baltimore, MD, 394–403. Ridgway, S.H., McCormick, J.G., and Wever, E.G., 1974, Surgical approach to the dolphin’s ear, J. Exp. Zool., 188: 265–276. Ridgway, S.H., Green, R.F., and Sweeney, J.C., 1975, Mandibular anesthesia and tooth extraction in the bottlenosed dolphin, J. Wildl. Dis., 11: 415–418. Rieu, M., and Gautheron, B., 1968, Preliminary observations concerning a method for introduction of a tube for anesthesia in small delphinids, Life Sci., 7: 141–1146.
0839_frame_C29 Page 687 Tuesday, May 22, 2001 11:57 AM
Marine Mammal Anesthesia
687
Rommel, S.A., Pabst, D.A., McLellan, W.A., Williams, T.M., and Friedl, W.A., 1994, Temperature regulation of the testes of the bottlenose dolphin (Tursiops truncatus): Evidence from colonic temperatures, J. Comp. Physiol. B, 164: 130–134. Ryon, B., Braun, R., and Dalton, L., 1999, Sedation effects of midazolam and its antagonist flumazenil in the Hawaiian monk seal (Monachus schauinslandi), in Proceedings of the Society for Marine Mammalogy, Wailea, Maui, HI, 164. Sawyer, D.C., and Williams, T.D., 1996, Chemical restraint and anesthesia of sea otters affected by the oil spill in Prince William Sound, Alaska, J. Am. Vet. Med. Assoc., 208: 1831–1834. Sedgwick, C.J., 1999, Anesthesia for small to medium sized exotic mammals, birds, and reptiles, in Manual of Small Animal Anesthesia, Paddleford, R.R. (Ed.), W.B. Saunders, Philadelphia, 318–356. Sepulveda, M.S., Ochoa-Acuña, H., and McLaughlin, G.S., 1994, Immobilization of Juan Fernández fur seals, Arctocephalus phillipi, with ketamine hydrochloride and diazepam, J. Wildl. Dis., 30: 536–540. Shaughnessy, P.D., 1991, Immobilisation of crabeater seals, Lobodon carcinophagus, with ketamine and diazepam, Wildl. Res., 18: 165–168. Slip, D.J., and Woods, R., 1996, Intramuscular and intravenous immobilization of juvenile southern elephant seals, J. Wildl. Manage., 60: 802–807. Stirling, I., and Sjare, B., 1988, Preliminary observations on the immobilization of male Atlantic walruses (Odobenus rosmarus rosmarus) with Telazol®, Mar. Mammal Sci., 4: 163–168. Sweeney, J.C., 1993, Blood sampling and other collection techniques in marine mammals, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 425–428. Sweeney, J.C., and Ridgway, S.H., 1975, Procedures for the clinical management of small cetaceans, J. Am. Vet. Med. Assoc., 167: 540–545. Tuomi, P.A., Mulcahy, D.M., and Garner, G.W., 1996, Immobilization of Pacific walrus (Odobenus rosmarus divergens) with carfentanil, naltrexone reversal and isoflurane anesthesia, in Proceedings of the International Association for Aquatic Animal Medicine, Chattanooga, TN, 121–123. Tuomi, P., Grey, M., and Christen, D., 2000, Butorphanol and butorphanol/diazepam administration for analgesia and sedation of harbor seals (Phoca vitulina), in Proceedings of the American Association of Zoo Veterinarians and International Association for Aquatic Animal Medicine, New Orleans, LA, 382–383. Walsh, M.T., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., Webb, A.I., Beusse, D.O., Brock, K.A., Robertson, S.A., Abou-Madi, N., Cook, R.A., and Klein, L., 1988, Sedation and general anesthesia of four Artic [sic] walrus (Odobenus rosmarus), in Proceedings of the International Association for Aquatic Animal Medicine, 161. Williams, T.D., 1986, Mustelidae (sea otter), in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 820–822. Williams, T.D., and Kocher, F.H., 1978, Comparison of anesthetic agents in the sea otter, J. Am. Vet. Med. Assoc., 173: 127–130. Williams, T.D., and Sawyer, D.C., 1995, Physical and chemical restraint, in Emergency Care and Rehabilitation of Oiled Sea Otters, Williams, T.M., and Davis, R.D. (Eds.), University of Alaska Press, Fairbanks, 39–43. Williams, T.D., Williams, A.L., and Stoskopf, M.K., 1990, Marine mammal anesthesia, in CRC Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 175–192. Williams, T.M., Kooyman, G.L., and Croll, D.A., 1991, The effect of submergence on heart rate and oxygen consumption of swimming seals and sea lions, J. Comp. Physiol. B, 160: 637–644. Williams, T.M., O’Connor, D.J., and Nielsen, S.W., 1995, The effects of oil on sea otters: Histopathology, toxicology, and clinical history, in Emergency Care and Rehabilitation of Oiled Sea Otters, Williams, T.M., and Davis, R.D. (Eds.), University of Alaska Press, Fairbanks, 3–22. Woods, R., Hindell, M., and Slip, D.J., 1989, Effects of physiological state on duration of sedation in southern elephant seals, J. Wildl. Dis., 25: 586–590.
0839_frame_C29 Page 688 Tuesday, May 22, 2001 11:57 AM
688
CRC Handbook of Marine Mammal Medicine
Woods, R., McLean, S., Nicol, S., and Burton, H., 1994a, Use of midazolam, pethidine, ketamine and thiopentone for the restraint of southern elephant seals (Mirounga leonina), Vet. Rec., 135: 572–577. Woods, R., McLean, S., Nicol, S., and Burton, H., 1994b, A comparison of some cyclohexamine-based drug combinations for chemical restraint of southern elephant seals (Mirounga leonina), Mar. Mammal Sci., 10: 412–429. Woods, R., McLean, S., Nicol, S., and Burton, H., 1995, Antagonism of some cyclohexamine-based drug combinations used for chemical restraint of southern elephant seals (Mirounga leonina), Aust. Vet. J., 72: 165–171. Woods, R., McLean, S., Nicol, S., and Burton, H., 1996a, Chemical restraint of southern elephant seals (Mirounga leonina); use of medetomidine, ketamine and atipamezole and comparison with other cyclohexamine-based combinations, Br. Vet. J., 152: 231–224. Woods, R., McLean, S., Nicol, S., Slip, D.J., and Burton, H., 1996b, Use of the respiratory stimulant doxapram in southern elephant seals (Mirounga leonina), Vet. Rec., 138: 514–517. Work, T.M., DeLong, R.L., Spraker, T.R., and Melin, S.R., 1993, Halothane anesthesia as a method of immobilizing free-ranging California sea lions (Zalophus californianus), J. Zoo Wildl. Med., 24: 482–487.
0839_frame_C30.fm Page 689 Wednesday, May 23, 2001 8:06 AM
30
Intensive Care Michael T. Walsh and Scott Gearhart
Introduction Principles of intensive care for marine mammals do not differ significantly from those for other species, although the adaptations of marine mammals for an aquatic existence result in some peculiarities. Relatively little is known about this subject, as there are few arenas to experiment with the principles, and few animals have been involved. As a result, there are very few published articles on intensive care. However, it is a rapidly developing field, as veterinarians become more involved in the care of marine mammals. Clinicians are therefore encouraged to share their experiences at scientific meetings such as the International Association for Aquatic Animal Medicine (IAAAM) and the American Association of Zoo Veterinarians (AAZV). Current technology used to document meetings with searchable CD-ROMs (IAAAM) allows the interested clinician better access to otherwise obscure information.
Records and Instructions Two of the most important areas of critical care are record keeping and communication. Unfortunately, these are also the least favorite tasks for many clinicians. All treatments and their responses should be recorded in the patient’s record. In addition, it is essential that there be a set of written orders including treatments to be administered, how much, how often, and when, pertinent environmental settings where possible, diagnostic techniques to be performed, physiological parameters to monitor, behavior to note, and who to call and when. Check boxes placed next to each drug, with times of administration, are helpful to increase treatment compliance in complicated cases. Orders should be updated daily, and the clinician should observe the animal at least twice daily. Veterinary instructions should include directions for hydration, as well as nutritional support. Intensive care without concern for these aspects is often less successful.
Patient Evaluation Initial evaluation of the marine mammal patient involves a visual inspection of the animal, including observations on position in the water, respiratory rate and depth, attitude, reaction to stimuli, body condition (skin, fat, and muscle mass), movement, eye position, as well as characteristics of stool and urine, if present. These basic parameters are important for correlations with clinical laboratory data (see Chapter 19, Clinical Pathology). For animals with known histories,
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
689
0839_frame_C30.fm Page 690 Wednesday, May 23, 2001 8:06 AM
690
CRC Handbook of Marine Mammal Medicine
their appetite over the previous week may be important, as well as abnormal behavior, coughing, and any discharges. Interactions with other animals can also be important, since ill individuals often isolate themselves or may be harassed by other more dominant animals. The next level of evaluation should include blood work to determine other system involvement, as well as the severity of the illness (see Chapter 19, Clinical Pathology). All individuals should have blood samples taken for hematology and serum chemistries. Additional serum needs to be collected for further tests such as protein electrophoresis, serology, or to bank. This is also the appropriate time to attempt blood cultures, before antibiotic therapy begins. Presentation of an ill individual may be after commercial laboratory hours, not allowing completion of pertinent data, but the clinician can perform simple tests until laboratory analysis is completed. Hypoglycemia can be detected using commercially available glucose reagent strips used by diabetics. Many normal animals can show elevated glucose levels because of stress. A hypoglycemic animal may exhibit a partial glucose stress response, leading the clinician to believe that glucose levels are not dangerously low at that moment. Yet, a short time later, the animal may succumb to hypoglycemia. This is most commonly seen with very thin or young animals. Blood glucose must be tested first, if clinically indicated, and may require immediate therapy. A packed cell volume (HCT) and total solids can prompt the clinician to institute therapy for dehydration more quickly. Blood samples should be repeated often in the critically ill patient. If the animal is not stabilized, blood samples every 48 hours may be beneficial until blood parameters respond. In some cases, daily hematology may help guide initial therapy. In-house testing can be dramatically expanded with the use of handheld chemistry and blood gas devices, allowing patient-side diagnosis and rapid onset of guided therapy (see Chapter 29, Anesthesia). Cardiac evaluation, although standard in most species, is often underutilized in cetaceans, partially as a result of animal size and difficulty with auscultation. Electrocardiograms should be considered as a basic portion of a diagnostic workup in the critically ill or emergency patient (Van Bonn et al., 1996). Electrodes are available that are adhesive or resemble small suction cups that are useful for cetaceans (see Chapter 29, Anesthesia).
Rehydration Assessing total-body water status in the critically ill marine mammal patient may be a challenge. While the traditional method of measuring skin turgor (tenting the skin and assessing how quickly it returns to its original position) may work in theory on furred species, such as the polar bear (Ursus maritimus), sea otter (Enhydra lutris), and smaller pinnipeds, this may be impractical, and even dangerous, in the nonanesthetized patient. It is of little or no value in the totally aquatic species or in severely emaciated animals. Clinical signs such as sunken globes, lack of tearing, thin body condition, curled whiskers (in harbor seals, Phoca vitulina), dry feces or constipation may alert the veterinarian to suspect dehydration. Diagnostic tests yield perhaps the most important clues to the marine mammal’s hydration. Generally, a dehydrated animal will have elevated HCT (>55% in cetaceans) and total protein, as well as increased levels of sodium, chloride, BUN, and serum creatinine (see Chapter 19, Clinical Pathology). However, HCT and total protein may not be dependable indicators if the animal is anemic. Although elevated sodium and chloride levels in dehydrated marine mammals may be the result of intracellular fluid loss, the elevations may also be indicators of increased consumption of seawater.
0839_frame_C30.fm Page 691 Wednesday, May 23, 2001 8:06 AM
Intensive Care
691
Choice of method for rehydrating the critical patient is usually determined by the severity of dehydration, the animal’s temperament, and its physiology. The maintenance fluid requirement for marine mammals has been estimated to be 40 ml/kg/day, and replacement volumes at 80 ml/kg/day (Sweeney, 1990). The preferred method of rehydration is oral supplementation. If gastric motility is assumed to be normal, giving fresh water or half-strength electrolyte solutions via stomach tube is an excellent method utilized in cetaceans, manatees ( Trichechus manatus), and some smaller pinnipeds. Adequate gastric motility may be indicated by normal defecation and lack of gastric reflux upon the passage of the stomach tube. The choice of fluid is determined by the abnormalies in the blood work (e.g., electrolyte imbalances or elevated packed cell volume). A solution such as lactated Ringer’s or normal saline may be chosen and diluted to the strength desired, usually at a ratio of 50 : 50 with water. Often, plain water rehydrates the critical marine mammal, and may stimulate the animal’s appetite, sometimes as soon as the procedure is completed. Volumes of up to 2 to 4 l of fluids have been given to adult bottlenose dolphins (Tursiops truncatus) up to four times daily, average of 2 l twice a day per 200-kg (440-lb) animal, and pinnipeds between 0.5 and 1 l, two to four times a day, depending upon size of the animal. Manatees can accept anywhere between 1 and 4 l of oral fluids twice daily, depending on their size, and possibly more often if HCT and electrolytes are closely monitored and remain within normal limits (Walsh and Bossart, 1999). Severely debilitated marine mammals will not usually be able to handle large quantities of fluid at first, and thus must be given lower volumes initially. Always monitor for reflux and stop immediately if this occurs; it may be a result of too frequent tubing and/or too large a volume being given to the animal, or it could be an indicator of other complications, such as ileus. Should gastric motility be compromised, parenteral administration of fluids may be attempted. The subcutaneous route of administration is commonly used to administer fluids such as lactated Ringer’s or D5W to California sea lions (Zalophus californianus), northern elephant seals (Mirounga angustirostris), sea otters, polar bears, and other semiterrestrial marine mammals. Any area of loose skin that can accommodate adequate fluid volumes may be used (interscapular, axillary, inguinal). McBain and Reidarson (1995) attempted subcutaneous fluid administration at 50 ml/kg/day in a pilot whale (Globicephalus sp.) diagnosed with renal failure. The intraperitoneal route has also been used in the sea otter (Williams, 1990), small California sea lions, and cetaceans (Sweeney, 1990; Townsend, 1999), using large-bore catheters designed for intravenous use. Volumes of up to 3 l may be infused over a 1-hour period up to three times daily in bottlenose dolphins (Sweeney, 1990). The intraosseous route has been successfully used in debilitated sea otters, with insertion of the needle at the trochanteric fossa of the femur (Black and Williams, 1993). A similar site has been used in fur seals and California sea lions (Haulena, pers. comm.). It is likely that this route of parenteral support may prove to play a greater role in the future, particularly in young marine animals. It is possible to place intravenous catheters in critically ill marine mammals, although this is, at least in species other than phocids, often used as a last resort, because of the relative difficulty in placing and maintaining access. Intravenous catheters placed in the extradural veins of phocids and walruses work well to administer relatively large volumes of fluids in a short time (Ellsworth et al., 1997). Catheters can sometimes be maintained for several days by suturing a butterfly tape collar to the skin of the animal, and conducting periodic flushing with heparinized saline. A larger Tuohy-like catheter has been used by SeaWorld Orlando as
0839_frame_C30.fm Page 692 Wednesday, May 23, 2001 8:06 AM
692
CRC Handbook of Marine Mammal Medicine
a means of intravenous injection and fluid administration in walruses. The silastic tube is introduced through an angled needle inserted into the extradural vein through a caudal lumbar intervertebral space (see Chapter 19, Clinical Pathology). The advantage of this catheter is that the larger-bore tubing allows the greater flow rate needed for larger species. The angled end allows a reliable placement through thick tissues over the dorsum. Jugular catheters have been used in sea lions (MacDonald and Dierauf, 1981) and sea otters. Some pinnipeds may require surgical cut-downs to access either caudal gluteal or jugular veins (Gage, 1990). Despite a usually blind approach, it is possible to access the spinal venous sinus in bottlenose dolphins. This has been performed by infusion of 1% lidocaine with an 18-gauge, 1.5-in. needle at a caudal intervertebral space, followed by insertion of a 14-gauge, 5.5-in. Teflon over-theneedle catheter into the venous sinus (Van Bonn et al., 1996). Intravenous fluids have also been administered via 19-gauge butterfly catheters inserted into the central vein of the dorsal fins of bottlenose dolphins (Jensen, pers. comm.) and into the tail veins. Thrombosis has, however, been observed by the authors. McBain and Reidarson passed a 12-gauge, 24-in. needle through the abdominal body wall and into the caudal vena cava of an azotemic pilot whale, and administered fluids at a rate of 30 ml/kg/day.
Blood Transfusion Blood transfusions have rarely been given to marine mammals; the few cases reported are described below. A severe laceration of the uterus of a bottlenose dolphin occurred during extraction of a dead fetus, resulting in massive hemorrhage. The hemorraging was initially controlled with oxytocin given intravenously, intramuscularly, and into the vaginal wall. In addition, pressure was applied with towels and gauze placed in the vagina for 20 min. This animal did not require a transfusion, although another dolphin with similar uterine lacerations did. For the transfusion of this second dolphin, killer whale (Orcinus orca) blood was used, because no donor of the same species was immediately available. The dolphin’s serum was cross-matched with the killer whale’s cells before proceeding; then the female was given 2 l of blood, and did survive the procedure. Two other bottlenose dolphins have been transfused with up to 2 l of blood from other bottlenose dolphins without complication. Blood was removed from the ventral vertebral vessels where the main fluke vessels converge at the peduncle, using an 18-gauge needle (see Chapter 19, Clinical Pathology). Since venous access is not dependable, a blood collection bag was not used. In the procedure, 60-ml syringes were prefilled with 6 ml ACD anticoagulant, then gently rocked after collection. The blood was administered through the same anatomical site, after the recipient animal received 100 mg prednisolone sodium succinate or 30 mg of dexamethasone intravenously. While a major cross-match between donor and recipient would be preferred, a minor cross-match, at the least, should be attempted. A 13-kg Guadalupe fur seal (Arctocephalus townsendi) was successfully transfused with 300 ml of whole blood from a California sea lion (Gage et al., 1996). The blood was administered to the seal by the intraosseous route through the trochanteric fossa of the femur. The animal recovered and was released. Control of hemorrhage has also been aided by administration of vitamin K at 0.5 to 1 mg/kg orally twice a day. This may be especially important in animals on extended courses of antibiotics. Another, less standard, approach has been the use of yunan paiyio, a Chinese herb that contains a glycoside known to enhance clotting (McBain, pers. comm.).
0839_frame_C30.fm Page 693 Wednesday, May 23, 2001 8:06 AM
Intensive Care
693
Nutritional Therapy Hypoglycemia Emergency administration of nutrition is not common, unless the animal is hypoglycemic. Oral supplementation of glucose in a severely hypoglycemic animal is a questionable practice, and should not be relied upon as the primary method of support. If it is the only option, then a 10 to 12% glucose solution can be administered, but the animal should be reevaluated within 30 min for signs of clinical improvement, and by blood sampling to determine whether the glucose has been absorbed. The amount of glucose can also be determined by giving 1 ml of 50% glucose/kg. Both approaches are rescue amounts, and will not maintain adequate levels for an extended period, so the patient should be rechecked frequently. The dose is repeated as needed, and a nutritional formula is then assembled to augment glucose and begin supplying other forms of energy such as protein and fats. In baby manatees, in which hypoglycemia is common, glucose therapy will typically only maintain the animal for 3 to 6 days unless adequate calories, including proteins and appropriate fats, are quickly included. Parenteral supplementation of glucose is usually reliant on access to dependable veins. In cetaceans, otariids, and manatees, this approach is complicated by poor venous access, but is still worth a try if the animal is not responding to other approaches. It should be noted that if access is questionable and extravascular deposition likely, then the concentration should not exceed 5% glucose to avoid hyperosmotic damage. With a reliable catheter, a 7 to 10% glucose solution can be used. Another potential route is intraperitoneal, using 5% dextrose that is near normal osmolarity. In all routes of administration, it is important to monitor serum response, and to wean the individual off the glucose when it is apparent that caloric needs are being adequately met. Premature removal from glucose supplementation and/or recurrence of severe hypoglycemia can often be fatal.
Emaciation A critical-care approach to nutrition may be needed in cases of extreme emaciation or when use of normal food results in vomiting, regurgitation, or complications such as intestinal abnormalities. The degree of nutritional support needed is directly related to the condition of the animal at the time of examination or intervention. If the animal is thin, as determined both by weight and by visual observation, then supplementation for fish eaters can be achieved with a gruel made of ground whole fish. This diet is considered a partial solution, since it is difficult to maintain an animal based on tube-feeding a fish gruel, especially a juvenile or neonate (see Chapter 37, Hand-Rearing, for details on feeding young animals). Tube feeding is a common technique for feeding emaciated or wounded manatees and pinnipeds. When making fish gruel for force-feeding, it is best to use the fish with the highest fat content. Herring may range from 890 calories/kg (April catch date) to over 2300 calories/kg (October catch date). An adult dolphin eating 8 kg of high-calorie herring will take in over 18,000 calories, while the same animal on 8 kg of low-calorie herring will take in only 7250 calories. Lower-caloric-density fish diets can be fortified by a number of techniques. Once the animal is rehydrated, the gruel can be made thicker by adding less water. Initially, gravity feeding is the usual approach taken until the correct volume is reached. Once the fear of overfeeding, or regurgitation, is lessened, then the thicker gruel can be pumped in, more closely approximating whole fish. Fish oils, such as menhaden or salmon, can be added to the gruel to increase the caloric density. Manatees can be supplemented with a gruel consisting
0839_frame_C30.fm Page 694 Wednesday, May 23, 2001 8:06 AM
694
CRC Handbook of Marine Mammal Medicine
of monkey chow biscuits, lettuce, and other vegetables. Common lettuce types fed range from 15 to 20 calorie/100 g. There are numerous diets available for humans formulated for different ailments. Unfortunately, many depend on carbohydrates as an important energy source. This can be a problem in species not used to large amounts of sugars; their use results in gas formation and diarrhea. In addition, some fats and proteins can cause similar effects. Most of the use of human intensive-care formulas by the authors has been as a supplement with gruel to bolster the caloric content rather than as a sole source. TwoCal HN® (Ross Products, Columbus, OH), a high-nitrogen formula, has been used in both manatees and dolphins, but to reduce complications, it is incorporated as an additive. The amount is added over a few days, time to allow for adaptation to the new nutrients. An elemental diet has been used in neonatal manatees with intestinal disease, such as pneumatosis intestinalis. It is designed to avoid complications by simplifying absorption of basic nutritional components in the small intestine, and allowing the large intestine to rest. When combined with medium-chain triglyceride oil, it can provide 2 calories/ml formula. It has not been used in any marine mammal other than the manatee, but may definitely have application in other species. Force-feeding whole fish is another option for dietary supplementation of cetaceans and pinnipeds. There are a number of complications that can develop with what would seem the most natural approach. Force-feeding an animal that is dehydrated can result in regurgitation or accumulation of dehydrated fish debris in the stomach, followed by vomiting. Before beginning force-feeding, it is best first to tube-feed the animal with water to help rehydration. This is advisable even if the blood work does not indicate dehydration, since the intestinal tract will be relatively dehydrated and motility may be impaired. This is especially important in beached animals with no history, which may be force-fed before being properly rehydrated.
Appetite Stimulants Marine mammals in a debilitated state usually find themselves in a negative energy balance and thus require vastly increased caloric intakes (see Chapter 36, Nutrition). The most common compounding problem is that the animal often suffers from partial to complete anorexia at the same time. Appetite stimulants often prove beneficial in the critical animal that is actively eating, but still not meeting its nutritional needs. Medications such as diazepam can serve dual purposes. Frequently, it will improve the patient’s feeding response (as is commonly seen in cats), while at the same time may act as an anxiolytic, thereby reducing what may be elevated levels of mental stress resulting from repeated handling. Many clinicians successfully utilize glucocorticoids to stimulate appetite in critically ill marine mammals. These drugs may also act as anti-inflammatory agents, although this can be detrimental, as they may further suppress what is most likely an already compromised immune system. Patients must be monitored for any of the side effects that may result from prolonged use of steroids, including the propensity for development of hypocalcemia (see Chapter 31, Pharmaceuticals). This is a condition that often needs to be corrected in the stranded or emaciated animal through the administration of calcium gluconate. Megestrol acetate is another drug that appears to stimulate appetite in cetaceans. It must be noted, however, that this compound takes roughly 10 days for effect in these animals. This may limit its usage in the critically ill patient. In addition, prolonged use in females may lead to adverse effects on reproduction.
0839_frame_C30.fm Page 695 Wednesday, May 23, 2001 8:06 AM
Intensive Care
695
Respiratory Emergencies Supplemental oxygen has been used for smaller animals that can be placed within oxygen cages. Larger pinnipeds requiring oxygen have been placed in plastic sealed transport boxes. A cetacean that is immobile can have an oxygen hose placed above its external nares (blowhole). Cetaceans that are still mobile are more difficult to supplement, although use of a long oxygen hose on an extendable rod has been attempted. Where mechanical oxygen support is not feasible, some respiratory relief may be provided with drug therapy such as steroids, aminophylline, and appropriate antibiotics (see Chapter 31, Pharmaceuticals). Emergency tracheotomy is not a common procedure; it has been performed on a young walrus (Odobenus rosmarus) with severe dyspnea (Dalton, pers. comm.). Chest tubes have been placed in manatees with pneumothorax by the authors. These animals usually present with asymmetric buoyancy, dyspnea, and severe distress. They are sometimes in danger of aspiration and cannot dive to eat. To keep water from entering the Heimlich valve of the chest tube, two one-way valves are added distally. In one case, the desired results, however, were not achieved. Removal of the trapped air in this breath-holding species appeared to have prolonged the healing of torn pulmonary tissue. During constant positive pressure, the air escaping from the lung kept the wound open for a long period, delaying closure. This gave the body time to respond to the presence of the tube, resulting in a granulomatous response, and secondary pleuritis. Treatment was changed to draining the chest periodically, and maintaining the animal in a flotation jacket to correct asymmetrical buoyancy, avoid inspiration of water, and allow a normal position in the water column for eating. The manatee’s diaphragm divides the chest into hemithorax (see Chapter 9, Anatomy), allowing some variability in treatment options. Depending on the severity of the original injury, resolution may take a few weeks to 4 months. Pneumothorax in cetaceans results in a higher degree of discomfort and dyspnea. Air leakage can be rapid, causing the animal to roll 90°, making each respiration a difficult task. One beached whale developed a severe pneumothorax after attempts to access the caudal vena cava for administration of fluids. Insertion of three 16-gauge needles could not keep up with the pressure, and the animal expired. Constant air release with a valve system should be a higher priority, but may still require a flotation jacket to maintain sternal buoyancy.
Trauma Surgical management of traumatic injuries follows principles of wound repair in terrestrial species. Few reports of surgical management of trauma cases in marine mammals exist, despite the frequency with which marine mammals strand with traumatic lesions (see Chapter 23, Noninfectious Diseases). The authors, in partnership with a dentist, repaired a fracture of the mandible of a juvenile California sea lion by wiring an indwelling piece of molded silicone between the mandibles and maxilla. It contained a central hole for tube-feeding that allowed handling of the mouth area without manipulating the jaw. Kirschner–Ehmer apparatuses have been used on cetaceans with success. Elastic Velcro strips have been used to splint a fractured maxilla to the mandible in a spotted dolphin (Stenella attenuata) (Townsend, 1996). An elastic band resulted in pressure necrosis in one animal from improper use by the authors. The authors have successfully used sections of neoprene from the arms of wet suits as splints for fractured mandibles in bottlenose dolphins. These can be removed for short periods of time to feed
0839_frame_C30.fm Page 696 Wednesday, May 23, 2001 10:58 AM
696
CRC Handbook of Marine Mammal Medicine
the animal, then reapplied. There is less chance of pressure necrosis, but they can work loose, and slip down the tapered rostrum. Making relief cuts in the middle of the material can stabilize the position. It is also important that the splint not be removed for any length of time until the jaw is healed. Some animals will resist reapplication and learn to take these devices off. Fracture fixation in polar bears is a challenge because of the inability to handle the individuals without general anesthesia. A juvenile polar bear with multiple hind-limb fractures was treated using standard orthopedic techniques combined with good clinical management (Cook et al., 1994).
Wound Management Management of open wounds in species that are resistant to handling is a challenge to the clinician. While bandaging is usually out of the question, the wounds can often still benefit from the application of disinfectant solutions such as betadine applied while the animal is eating or otherwise distracted. Most species can be trained fairly quickly to accept this approach. For cetaceans with open wounds, some ointments have been used but with little real success. Antibiotics, regular debridement and cleaning, and good water quality are often the best therapy. Hosing wounds from a distance is a useful method for cleaning pinniped wounds. In manatees, some wounds might benefit from neoprene body wraps, whereas others need to be treated like cetaceans. The clinician can also take advantage of changing water salinity, to undermine a wound infection that began in salt water, by switching the animal to fresh water, or vice versa.
Central Nervous System Neurological conditions requiring emergency care include metabolic disturbances, toxicoses, trauma, and infectious diseases. Hyponatremic pinnipeds may present seizuring or comatose (see Chapter 41, Seals and Sea Lions). Distinction from hypoglycemia and diagnosis of hyponatremia are based on decreased sodium and chloride levels in serum. Adult sea lions may show symptoms of varying severity with serum sodium levels below 147 mEq/l (Geraci, 1972). An important predisposing factor is housing in fresh water. Treatment with saline intravenously, subcutaneously, or even intraperitoneally can result in therapeutic response. A 40-kg (88-lb) juvenile California sea lion that presented comatose, with serum sodium of 116 mEq/l, was given 3.5 l of saline subcutaneously. This animal began to show a return of some reflexes within 1 hour. Over the next 48 hours, the sea lion progressed to sitting up and being attentive to its surroundings. Supplementation with subcutaneous saline was continued until the serum sodium levels were greater than 140 mEq/l, and the animal was eating fish with 2 to 3 g of sodium chloride/kg of food. Nonsteroidal anti-inflammatory drugs may be useful in trauma cases associated with neurological signs. Control of seizure activity in marine mammals is similar to that in other species. Seizures in California sea lions exposed to domoic acid can be treated with diazepam, midazolam, or lorazepam initially (Gulland, 2000), and phenobarbitol at 4 mg/kg for longer-term control (Haulena, pers. comm.). For idiopathic seizures in captive California sea lions, the dose may be 2 mg/kg twice daily, with adjustments based on clinical response and blood levels in the 20 µg/dl range. Larger species, such as a killer whale, would receive 1 mg/kg (McBain, pers. comm.).
0839_frame_C30.fm Page 697 Wednesday, May 23, 2001 8:06 AM
Intensive Care
697
Reproductive Emergencies Dystocia The most common form of reproductive emergency is a dystocia or difficult labor. This can occur as a result of problems related to the fetus, such as a large baby, malposition, or in utero death, or from problems with the mother, such as uterine inertia. It is helpful to have an idea of the normal steps in the process and when to intervene. In cases where the clinician has less exposure to a particular species, he or she must call on colleagues who can assist. In cetaceans and manatees, fluke presentations are preferred, since the presence of the fluke first allows it to unfold and solidify, making swimming much more efficient after birth. Headfirst presentation can still result in a live calf, but it will initially be more prone to complications, such as difficulty in swimming and surfacing. With close observation, the rupture of the placental membranes may be observed followed by the exteriorization of the tail. There have been a few cases of prolonged births that resulted in stillborn fetuses, in which the tail of the fetus was flexed laterally, almost dislocating the vertebrae. It has been suggested that the tail of the fetus did not enter the cervix properly in these cases, resulting in attempts to force the peduncle through the canal, damaging the vertebrae. Whether these were acute problems, or chronic malpositions, is unknown. There have been bottlenose dolphin calves born with tail malformations, but it is not known if these were birth-related trauma or not. The time from tail presentation to birth in most cetaceans may range from 1 to 4 hours, with new mothers generally taking longer. A few normal live births have occurred as late as 6 hours after fluke presentation, but the clinician needs to become concerned if there is no progress after 5 hours, and prepare for possible intervention. On-site clinical judgment should take precedence over set time intervals. Signs of difficulty to watch for include loss of calf tail movement, cessation of progress in calf expulsion or active labor, and expulsion of placental tissue during the birth. Sometimes the new mother is fatigued and will rest between contraction periods, so it is important to keep track of this activity. If it appears that the mother is not able to finish the delivery, the decision must be made to evaluate and possibly assist. Once a decision has been made to restrain the mother, the animal should be examined using ultrasonography to determine whether the fetus is still alive. If alive, the choices are to administer oxytocin (20 units intramuscularly) and sometimes injectable calcium, or to pull the calf. In six different dystocia cases in cetaceans, including a killer whale and a beluga (Delphinapterus leucas), the calf was already dead, with two showing early fetal decomposition. In five of these cases, ultrasound was helpful in determining calf viability before deciding on a course of action (see Chapter 26, Ultrasonography). The use of oxytocin appears to be of minimal help and should always be used in conjunction with calcium (Robeck, pers. comm.). In the six cases above, oxytocin did not result in expulsion of any of the calves, and appeared to make extraction more difficult, because the uterus contracted down on the fetus. Intervention in the beluga case did not utilize oxytocin preextraction, and proceeded more smoothly. When handling a dystocia, the female may require sedation. Four of the animals above received injectable midazolam to decrease stress and improve muscle relaxation. Copious amounts of lubricant were injected via a small tube around the fetus, since there is a tendency for the fetus to adhere to the placenta and uterus. In some cases, placing a fluid pressure bag in the canal and inflating it enhanced dilatation of the vaginal opening. When a fetus presents tail first, there is an obvious site for traction. With a head-first presentation, the clinician must gain access to the mouth, eyes, and blowhole for placement of hooks and cables or chains for extraction. With very large calves and a tail-first delivery, the major areas of detainment include the chest, dorsal fin (which is fairly pliable), and the pectoral flippers. Initial attempts at removal
0839_frame_C30.fm Page 698 Wednesday, May 23, 2001 8:06 AM
698
CRC Handbook of Marine Mammal Medicine
may result in lacerations of the mother’s reproductive tract, sometimes with severe consequences. It is imperative that there be one clinician directing the procedure, since there is a tendency to hurry once there is progress, thereby resulting in more tears. If at all possible, the position of the flippers must be determined manually, at various intervals in the procedure, to avoid the fetus taking on a knifelike posture. Also, avoid the temptation to try to spin the calf once tension is placed on the fetus. This may promote a change in the position of the pectoral flippers, and wind the uterine or placental tissue on the fetus. Pulling against a large fetus results in complications that can be avoided by using a modified fetotomy. Once the abdomen of the fetus is accessible, an incision is made through the wall, and the abdominal and thoracic contents removed. This allows the chest to collapse, facilitating removal and avoiding additional tears to the female. After removal, the uterus can be flushed with a weak betadine/saline solution followed by sterile saline. Next, oxytocin is administered to shrink the uterus. Uterine boluses have also been instilled and the last portion of the saline flush can incorporate antibiotics. In some cases, additional flushes may be needed to clean the reproductive tract. Aerobic and anaerobic bacterial cultures should be done on the fetus to determine which bacteria may be involved. Other complications related to dystocia and calf extraction have included a severe cervical laceration resulting in chronic hemorrhage and septicemia, severe metritis secondary to bacterial infection, and a severe laceration of the uterus and vaginal area resulting in a chronic abdominal abscess. Broad-spectrum antibiotics (including one specific for anaerobes) should be included in the therapy. In stranded manatees, dystocias are more complicated because of the extended delays in finding and rescuing sick individuals. These animals are sometimes compromised by watercraft trauma that can result in abortion. Of four cases in which females were rescued with dead fetuses, only one female survived. Each of the other three had necrotic fetuses, were very toxic with severe necrosis of the uterus, and lived only a few days after removal of the fetus.
Other Reproductive Emergencies The authors have seen pyometra that was detected by ultrasound in a bottlenose dolphin 1 week after her calf had died. The uterus was grossly distended but did not respond to medical therapy including attempts to open and drain the uterus through the cervix, or to initiate hormonal manipulation. The animal became very toxic and was taken to surgery, but expired the day after recovery from anesthesia.
Antibiotics Broad-spectrum, bactericidal, injectable antibiotics are usually the best choices for treatment of a critically ill marine mammal. Injectable agents are preferred as they reach therapeutic levels more quickly and reliably. In addition, they are not affected by vomiting, regurgitation, or diarrhea, and do not have as many interactions with other oral therapeutics. Oral antibiotics may be used in combination with parenteral drugs, as part of therapy for intestinal diseases, or as follow-up once the animal is stabilized. In some species, such as the manatee, oral medications routinely cause diarrhea. Third-generation cephalosporins, such as ceftriaxone and ceftiofur, are particularly useful drugs, not only because of their spectrum of activity, but also because they are administered only once daily (see Chapter 31, Pharmaceuticals). Should the veterinarian suspect a resistant or overwhelming infection, a combination of antimicrobials that act synergistically may be indicated. The cephalosporins, given
0839_frame_C30.fm Page 699 Wednesday, May 23, 2001 8:06 AM
Intensive Care
699
in tandem with an aminoglycoside such as amikacin, have been effective treatment regimens that can cover many severe bacterial infections, regardless of tissue site within the patient. While amikacin is less nephrotoxic than some of the other aminoglycosides, it is important to ensure proper hydration status of the animal through the serial monitoring of such blood parameters as HCT, total protein, blood urea nitrogen, and creatinine. Supplemental fluid therapy may be considered a necessary part of the critical care support approach when using these agents.
Analgesics Pain relief must also be a consideration in the intensive care of marine mammals. The greatest challenge is knowing when pain is present, since pain may be a major factor in treatment failure. Marine mammal patients that present with obvious traumatic lesions or exhibit symptoms such as abdominal cramping, bloating, or dyspnea suggesting internal pain may benefit from the administration of analgesics. Flunixin meglumine has been given to a variety of marine species for its potent analgesic properties. It may also be valuable for its ability to lessen the effects of endotoxins. Originally, its application was based on equine use, with beached cetaceans given dosages of 0.5 mg/kg as often as twice a day. Clinically, it appeared to provide notable relief, but a small percentage of animals that died showed histological signs of renal papillary necrosis. McBain and Reidarson (1995) caution against its use in pilot whales, as it was suspected to cause renal papillary necrosis. It may also contribute to ulcer development or recurrence. The dosage used at SeaWorld for larger animals has been reduced to 0.25 mg/kg once a day, and the length of therapy is limited. The authors have employed the use of opioids and their derivatives in attempts to alleviate pain in marine mammals. Meperidine has been used orally and parenterally in some cetaceans and the walrus, with some perceived positive effects. Butorphanol, a narcotic agonist–antagonist, has been used in manatees, California sea lions, and northern elephant seals (Haulena, pers. comm.). One of the major disadvantages to the use of butorpha-nol is the presumed relatively short half-life, necessitating a frequency of treatment that would prove to be fairly impractical in aquatic animals. Narcotics are also known for their capacity to decrease gut transit time, which is an undesirable effect in animals that may have some degree of decreased gastrointestinal motility.
Miscellaneous Therapeutic Agents Stranded marine animals often suffer from a variety of illnesses, and may benefit from therapy for hypocalcemia, myositis, gastric ulcers, hemorrhage, acidosis, and septicemia. As a matter of protocol, the clinician may elect to administer compounds such as vitamin E and selenium, B complex, or others, i.e., vitamin K, especially if the patient appears to have suffered from severe hemorrhage or perhaps internal blood loss due to gastric ulcers.
Support Equipment Many cetaceans, when seriously ill, succumb to drowning, so emergency care includes physical support. One method is to use personnel who support the animal by walking adjacent to it in the water. The disadvantage of this method is that it can exhaust personnel, and the animal may not cooperate when it gains strength, yet is still unable to support itself. Another method is to suspend an animal in a stretcher that is held in the water column. The disadvantages to this method include pressure sores from stretcher contact, poor application with movement
0839_frame_C30.fm Page 700 Wednesday, May 23, 2001 8:06 AM
700
CRC Handbook of Marine Mammal Medicine
of the animal, and the need for constant surveillance. A third method is the use of a flotation device, a jacket constructed of wet suit material that is designed to provide buoyancy for the animal without restricting its activity, but still avoiding drowning (Walsh et al., 1995).
Conclusion Intensive care in marine mammals parallels procedures for similar situations in terrestrial domestic and wild animals. It is important for the clinician to have emergency equipment and pharmaceuticals at the ready and easily accessible. It is only through careful planning, meticulous record keeping, constant communication with colleagues, and response efficiency that intensive care procedures are now saving the lives of many marine mammals in our care.
References Black, M., and Williams, T.E., 1993, Intraosseous infusion in the sea otter, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. 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. Castellini, J.M., Meiselman, H.J., and Castellini, M.A., 1996, Understanding and interpreting hematocrit measurements in pinnipeds, Mar. Mammal Sci., 12: 251–264. Cook, R.A., Calle, P.P., and Wood, R.L., 1991, Successful medical management of an Erysipelothrix rhusiopathiae infection in a beluga whale (Delphinapterus leucas), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Cook, R., Thatcher, C., Calle, P., Raphael, B., Kapatkin, A., and Stetter, M., 1994, Multiple hindlimb fracture repair in an adolescent polar bear (Ursus maritimus), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Cooperrider, D.E., 1968, Skin lesions of Erysipelothrix insidiosa, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Dalton, L., and McBain, J., 1993, Mucormycosis in three cetaceans, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Dierauf, L.A., 1990, Pinniped husbandry, in Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 553–590. Ellsworth, L.B., St. Aubin, D.J., and Dunn, J.L., 1997, Effects of saline infusion on circulating atrial natriuretic peptid in harbor seals (Phoca vitulina), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Gage, L.J., 1990, Rescue and rehabilitation of cetaceans, in Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 685–692. Gage, L.J., Beckman, K., Wickman, D., and Smith, D.M., 1996, Transfusion of a Guadalupe fur seal (Arctocephalus townsendi) with California sea lion (Zalophus californianus) blood, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Geraci, J.R., 1968, Diet-induced thiamine deficiency in captive marine mammals, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Geraci, J.R., 1972, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618–623. Gulland, F.M.D., 2000, Domoic acid toxicity in California sea lions (Zalophus californianus) stranded along the central California Coast, May–October 1998, NOAA Technical Memorandum, NMFS-OPR, 44 pp. Hammond, D., 1977, Epidemic of Pseudomonas pseudomallei in an aquatic park, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive.
0839_frame_C30.fm Page 701 Wednesday, May 23, 2001 8:06 AM
Intensive Care
701
Lowenstine, L.J., Groff, J., Rideout, B., Wong, A., and Gage, L., 1990, Necropsy findings in two juvenile beaked whales (Mesoplodon sp.) maintained in captivity for rehabilitation and stranding, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. MacDonald, M.K., and Dierauf, L.A., 1981, Surgical treatment of a compound, comminuted fracture of a radius in a beached California sea lion, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Martensson, G., 1996, Seasonal changes in energy density of prey in northeast Atlantic seals and whales, Mar. Mammal Sci., 12: 635–640. McBain, J., and Reidarson, T.H., 1995, A case of renal failure in a Pacific pilot whale (Globicephala macrorhynchus) and attempted therapy, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Reynolds III, J.E., and Odell, D.K., 1991, The Florida manatee, Manatees and Dugongs, Facts on File, New York. Seibold, H.R., and Neal, J.E., 1956, Erysipelthrix septicemia in the porpoise, J. Am. Vet. Med. Assoc., 128: 537–539. Stetter, M.D., Calle, P.P., McClave, C., and Cook, R.A., 1997, Marine mammal intravenous catheterization techniques, in Proceedings of the American Association of Zoo Veterinarians, Houston, TX. Sweeney, J.C., 1990, Surgery, in Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 215–234. Townsend, F.L., 1996, Medical management of a maxillary fracture in a Stenella attenuata, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Townsend, F.I., 1999, Medical management of stranded small cetaceans, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 485–493. Van Bonn, W.G., Jensen, E.D., Miller, W.G., and Ridgway, S.H., 1996, Contemporary diagnostics and treatment of bottlenose dolphins: A case report, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., and Dover, S.R., 1997a, Dietary factors II: Caloric seasonality and management implications, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., and Dover, S.R., 1997b, Dermatitis associated with hypothermia in juvenile Florida manatees (Trichechus manatus), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., Bossart, G.D., and Campbell, T.W., 1991, Traumatic injuries in manatees (Trichechus manatus), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CDROM Archive. Walsh, M.T., Thomas, L.A., Songer, G., Campbell, T.W., Schroeder, P., and Tucker, L., 1994, Clostridial perfringens isolates from cetaceans, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., Wagoner, B.C., Campbell, T.W., and Rodríguez, A., 1995, Use of flotation devices in marine mammals: Treatment and physical therapy, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., Friday, R.B., Johnson, A.B., and Messinger, D., 1996, Regurgitation in cetaceans: Medical implications, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Williams, T.D., 1990, Sea otter biology and medicine, in Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 625–648.
0839_frame_C30.fm Page 702 Wednesday, May 23, 2001 8:06 AM
702
CRC Handbook of Marine Mammal Medicine
Young, S.J.F., Huff, D.G., Ford, K.B., Anthony, J.M.G., Ellis, G., and Lewis, R.L., 1997, First case report—Mortality of wild resident killer whale (Orcinus orca) from Erysipelothrix rhusiopathiae, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive.
0839_frame_C31 Page 703 Tuesday, May 22, 2001 1:37 PM
31 Pharmaceuticals and Formularies Michael K. Stoskopf, Scott Willens, and James F. McBain
Introduction Over the decade since the first edition of this book, dedicated veterinarians, biologists, and other scientists have added to the knowledge of the pharmacological requirements and tolerances of marine mammals. This chapter provides a compilation of the available pharmacological information on cetaceans, pinnipeds, sirenians, sea otters, and polar bears to provide clinicians and scientists working with marine mammals a convenient and rapidly accessible single source on the subject. Readers must be aware at all times that drugs discussed in this chapter have only been used on a very limited number of individual animals from a narrow range of species, so all information must be interpreted with caution. None of the drugs in common usage today has been licensed for use in marine mammals. The authors have relied heavily on published documentation, which is included relatively uncritically, but they have also included unpublished information from clinicians with experience with some of the less frequently encountered species. Even so, numerous gaps remain. Most of the drug regimens included have been supported by either pharmacokinetic studies or documented clinical response, although only rarely have detailed studies been performed on serum levels of pharmaceutical agents (e.g., Tyczkowska et al., 1992). Furthermore, these observations have been made on extremely few individuals, so undocumented effects may still occur when these drugs are used on larger numbers of individuals. The tabular format was selected for the convenience of clinicians needing information quickly. There are advantages and limitations to this presentation. The primary advantage is accessibility. The primary weakness is the limited amount of information presented with each entry. The tables are not intended to replace a strong background in clinical veterinary medicine and pharmacology. Readers are directed to individual references for further information, and are cautioned to read, understand, and discuss with colleagues the pharmacological properties of the drugs they intend to administer to a marine mammal, even though dose regimens appear in the tables (Benet et al., 1996; Riviere, 1999). Dosages and adverse effects of anesthetic agents are discussed in Chapter 29, Anesthesia. Always be cautious when administering drugs or drug combinations for the first time in a marine mammal. If the decision is made to adopt a novel treatment regime, only one animal should be treated and then observed for an appropriate time to ascertain whether adverse reactions occur. Finally, always check with others working
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
703
0839_frame_C31 Page 704 Tuesday, May 22, 2001 1:37 PM
704
CRC Handbook of Marine Mammal Medicine
with marine mammals before using an unfamiliar drug, as they may be aware of adverse reactions that have not been reported.
Routes for Administering Drugs to Marine Mammals Essentially all of the routes used to deliver drugs to domestic animals are available for delivering drugs to marine mammals. Practical considerations, however, frequently limit the choice of routes in a clinical situation, and anatomical adaptations can make application of some routes particularly challenging. Details of choice of route are discussed in each species medicine chapter (see Chapters 40 through 45), but precautions to take are given below. From a practical standpoint, oral (PO) administration of drugs is often the preferred route in an animal still taking feed regularly or being tube-fed routinely for nutritional support. Few special caveats for oral administration of drugs to marine mammals have been discovered, and a good knowledge of human or domestic animal pharmacology provides excellent guidance for the appropriate selection of this route. The major considerations are food interactions and factors that might affect absorption. These factors include specific physiochemical properties of the drug, stomach pH, gastrointestinal microflora, and anatomy (Riviere, 1999). For example, phosphorus binders are commonly used to treat California sea lions (Zalophus californianus) with leptospirosis, as is tetracycline, yet the absorption of this antibiotic will be reduced by chelating agents (Gulland, 1999). Subcutaneous (SC) administration may be problematic because of the blubber layer in most pinnipeds and cetaceans, but can be effective in sea otters (Enhydra lutris) and fur seals. Prior to the development of the thick blubber layer, young pinnipeds, especially neonates, can also adequately absorb even lipophilic medications delivered by this route, which avoids muscle trauma or necrosis. Although subcutaneous administration of fluids is often avoided in cetaceans, experienced clinicians have successfully administered higher volumes of fluids to these animals by this route by ensuring that the fluids are given between the blubber and muscle layers (see Chapter 40, Cetacean Medicine). Intramuscular (IM) administration is frequently applied in marine mammals that are difficult to restrain and are inappetent. Be cautious and avoid superficial injection into the extensive subcutaneous blubber, which has dramatically different drug-partitioning properties than does muscle. Accidental delivery into the blubber can result in failure to achieve any appreciable systemic distribution of highly lipid soluble medications. Another precaution applies to the volume of injection. The irritation caused by some injectable drugs is a local effect. The recommended total volume injected per site does not increase in scale with the mass of the animal. Very large marine mammals may require large volumes of drug. Care should be taken to use multiple injection sites and to keep the drug volume per injection site reasonable. Many venipuncture approaches (IV) are nearly perpendicular to the vessel, which can be quite deep, complicating catheterization. Needles with side ports for directing a catheter at right angles can help avoid perivascular leakage of irritating drugs (Sweeney, 1990). Nonirritating drugs can often be delivered intraperitoneally (IP). The difficulties of this route are generally related to the size of the patient, and the availability of needles suitable to penetrate the abdominal wall. Placing a flexible catheter through a cannula for intraperitoneal administration avoids the problem of accidental organ laceration with a rigid needle (Sweeney, 1990). Intratracheal (IT) and inhalation (IH) administration of drugs has been employed in marine mammals, primarily for induction of anesthesia and targeted delivery of medications to the lungs. Nebulization and aerosolization can be used very effectively, either by holding a mask on a restrained animal or by placing it in a nebulization chamber, if drug delivery times are adjusted to accommodate an animal’s tendency to hold its breath.
0839_frame_C31 Page 705 Tuesday, May 22, 2001 1:37 PM
Pharmaceuticals and Formularies
705
The major challenge with topical administration in marine mammals is to achieve appropriate contact time for drug efficacy in an aquatic environment. Baths and dips are possible for some of the smaller species, and behavioral modification can augment dipping body parts into smaller containers for large species. However, the most desirable application in some cases would be an ointment or salve that would remain in place; thus, ointments for marine mammals are often specially compounded. The use of human dental bases has had some success but can be prohibitively expensive. Less expensive lipid bases such as lanolin and petroleum gels have been used with varying success.
Dose Scaling Extrapolation of doses and pharmacokinetic parameters across species is often necessary as pharmacological data for drugs in most species of marine mammal do not exist (Riviere, 1999). Even for drugs that have been studied in marine mammals, sample sizes are often small, and consideration of covariates such as body weight, enzymatic composition, and genetics are necessary to give a full picture of allometric relationships (Riviere, 1999). Allometric equations usually compare parameters of interest (e.g., half-life, volume of distribution, clearance) to body weight (Riviere, 1999), and a dizzying array of exponential equations can be found in the literature. Choosing which equation to use can be daunting. The reader should be cautioned that not all drugs scale well by body mass, even when theoretical metabolic rates are figured into the equation. It is important to know the expected metabolism and excretion routes of the drug when making decisions on scaling a dose between species. Recent studies on species of phocids of different size suggest that effective doses based on body weight, rather than on a complex allometric equation, may not be that unreasonable for some drugs (Gulland et al., 2000).
Drug Interactions When new combinations are considered, it is best if there has been some experience with the drugs individually in the species. If not, the consideration should be made to administer one drug at a time in an individual animal. It is not feasible to present a comprehensive list of all possible drug interactions in marine mammals in this chapter. The reader should be prepared for the possibility of any drug interaction described in any species occurring. However, it is particularly important to be aware of some of the potential interactions of more commonly used medications. At present, little documentation exists concerning drug interactions in marine mammals, and the majority of these reports are intuitive from knowledge of terrestrial animal pharmacology. Texts on veterinary and human pharmacology should be consulted, and discussions with colleagues undertaken, when planning a mixed medication treatment for any marine mammal.
Cimetidine and Antacids The use of alumina gel–based antacids and histamine receptor (H2) blockers, such as cimetidine and ranitidine, in the treatment of gastric ulcers is relatively routine. Several important drug interactions can be expected with concurrent use of these compounds, based on knowledge gained in human medicine. Simultaneous administration of antacids with cimetidine will significantly decrease the absorption and effectiveness of the cimetidine. To avoid this complication, administering antacids or cimetidine at least 1 hour prior to the other will allow adequate absorption of cimetidine. Antacids with di- and trivalent cations also decrease the absorption of oral steroids, such as prednisolone and dexamethasone. Antacids also reduce tetracycline
0839_frame_C31 Page 706 Tuesday, May 22, 2001 1:37 PM
706
CRC Handbook of Marine Mammal Medicine
and imidazole absorption. In the case of the imidazole ketoconazole, the use of antacids may alter stomach pH, to the point that requirements for dissolution and subsequent absorption of drugs are not met. To maximize absorption, steroids, imidazoles, and tetracyclines should be administered at least 2 to 3 hours before or after administration of antacids. McBain (1985) has documented a negative impact of cimetidine on absorption of tetracycline from the stomach in killer whales (Orcinus orca). Either combinations of these two drugs should be administered concurrently or the tetracycline should be administered first. Cimetidine can also impair benzodiazepine metabolism in terrestrial mammals by inhibiting specific hepatic microsomal enzymes. This can result in increased sedation when diazepam is administered to animals being treated with cimetidine. In such cases, lorazepam, oxazepam, or temazepam might be valuable alternatives to diazepam, since they have been found to have different metabolic pathways in terrestrial mammals. Cimetidine will also increase gastric pH and thereby decrease the absorption of concomitantly administered ketoconazole. A common problem encountered in cetaceans administered antacids is vomiting due to overmedication (McBain, unpubl. data). The gastric pH becomes too high to demineralize fish bones, which can be seen in the vomitus or via endoscopy. Normally, the acidic pH of the stomach will demineralize and digest fish bones within 30 min.
Tetracyclines In killer whales, the interactions between tetracyclines and cimetidine as described previously are complicated by the relatively poor levels of tetracycline achieved by oral administration (McBain, 1985). Calcium in the ingesta chelates with tetracycline and reduces absorption. Other factors that can decrease tetracycline absorption from the gastrointestinal tract include the presence of other multivalent cations such as magnesium, iron, or zinc in the stomach. Absorption of some synthetic tetracyclines, such as doxycycline or minocycline, is not as adversely affected by cations in the food, and these drugs may offer some benefit over other tetracyclines when medically indicated in cetaceans.
Fluoroquinolones Fluoroquinolones have been reported to cause cartilage damage in weight-bearing joints of young, rapidly growing animals (Burkhardt et al., 1990; 1997; Burkhardt, 1996; Yoshida et al., 1998). A proposed mechanism for this chondrotoxicity is the chelation of magnesium during a phase when high circulating magnesium is critical for the growth and development of articular cartilage (Burkhardt, 1996). The clinical responses of marine mammals to fluoroquinolone treatments have often been favorable, so they are often the broad-spectrum antibiotic of choice for many experienced clinicians in treating neonatal marine mammals suffering from severe infections of unknown origin. However, it is still wise to use caution when administering fluoroquinolones to juvenile marine mammals, and these patients should be carefully monitored for any signs that might be attributable to joint pain. Fluoroquinolones offer a useful broad-spectrum antibacterial therapeutic option. Crossresistance to other classes of antibiotics has yet to be well demonstrated. Although plasmidmediated resistance to fluoroquinolones is relatively uncommon (Burkhardt, 1996), indiscriminant use could lead to development of bacterial resistance to the drugs. Public concerns about the development of fluoroquinolone-resistant organisms are strong. Clinicians selecting these antibiotics for treatment of marine mammals should therefore take special care to ensure that the drug is indicated on the basis of culture and sensitivity data, and that an effective dose is delivered over a sufficient duration to effect a cure (Johnsen, 1994; Johnson et al., 1998). As with the tetracyclines, divalent cations may decrease absorption and enhance elimination of fluoroquinolones.
0839_frame_C31 Page 707 Tuesday, May 22, 2001 1:37 PM
Pharmaceuticals and Formularies
707
Other Antibiotics Little has been documented about interactions or adverse effects of antibiotics in marine mammals, although there are some important precautions that can be deduced from experience with terrestrial mammals. Cephalosporins and aminoglycosides are frequently administered to marine mammals. Clinicians should be aware that concurrent administration of aminoglycosides and cephalosporins increases the risk of renal toxicity, because the renal effects of these drug groups are additive. Prolonged administration of cephalosporins can also result in vitamin K deficiency and subsequent blood-clotting defects. Concurrent administration of aminoglycosides and flunixin meglumide has also been linked to renal papillary necrosis in a pilot whale (Globicephala spp.) (McBain, unpubl. data). Aminoglycosides and flunixin meglumide may be contraindicated in cases of toxemia because they both have antiprostaglandin activity. Administration of aminoglycosides, such as amikacin, in a single daily dose increases bactericidal activity and postantibiotic effect, allows more rapid attainment of high serum concentrations, and decreases risk of nephrotoxicity compared with administering multiple lower doses each day (Townsend et al., 1996; Riviere, 1999). Adverse reactions to sulfonamides have been reported in cetaceans. A moderate reaction is characterized by neutrophilia (Cornell, 1978). A severe reaction in killer whales included both neutropenia and thrombocytopenia. The exceptionally long half-life of some sulfonamides in cetaceans is likely a contributing factor in producing adverse reactions. Sulfamethoxazole was found to have a half-life of 5.3 to 7.2 days in killer whales (McBain, 1984). This extremely long half-life has not been noted in other sulfa drugs but the possibility that other sulfas may also have prolonged excretion times must be considered. Most of the cases of adverse reactions to sulfas have occurred with sulfa, trimethoprim combination drugs. In spite of this observation, the evidence points to the sulfa drugs as the culprits when reactions occur. The concurrent administration of folic acid at the rate of 25 to 50 mg per os twice daily, to bottlenose dolphins receiving sulfadiazine and trimethoprim, seems to reduce the likelihood of neutropenia and thrombocytopenia (McBain, 2000). Rifampin is gaining more widespread use in domestic animal medicine and may play a useful role in the treatment of marine mammals. This drug has been documented to cause an idiopathic thrombocyte dysfunction that can result in prolonged bleeding times (Marcus, 1982; Stoskopf et al., 1987). Rifampin also stimulates microsomal enzymes that are involved in the metabolism of steroids. Therefore, administration with oral or parenteral steroids may prevent the effects of steroids. This inhibition of steroid action through increased metabolic inactivation can have long-lasting effects, even after discontinuation of rifampin therapy. Rifampin administration enhances the elimination of both exogenous and endogenous steroids, compromising the ability of an animal to maintain metabolic homeostasis. Regulatory agencies cannot rule out the possibility that chloramphenicol poses a risk of a potentially fatal aplastic anemia to humans handling it. Bone marrow complications have not been reported in marine mammals. Use of another amphenicol, florfenicol, does not pose a potential threat to human handlers. However, florfenicol has been shown to cause an increase in aspartate transaminase (AST) and lactate dehydrogenase (LDH) due to muscle trauma at injection sites in bottlenose dolphins and belugas (Delphinapterus leucas) (Sweeney, 1986). It has not been demonstrated to cause anorexia, which is a common side effect of chloramphenicol administration in marine mammals (McBain, unpubl. data). Azithromycin has been reported to cause inappetence and abdominal discomfort in bottlenose dolphins, belugas, and orcas (Dalton et al., 1995). An increase in triglycerides that was not related to clinical morbidity was also observed in two beluga calves (Dalton et al., 1995).
0839_frame_C31 Page 708 Tuesday, May 22, 2001 1:37 PM
708
CRC Handbook of Marine Mammal Medicine
Antifungals The azoles have received considerable attention in the last decade for the treatment of fungal infections in marine mammals. Use of itraconazole, ketoconazole, or fluconazole in bottlenose dolphins has led to mild and reversible liver pathology and 2- to 25-fold increases in AST, alanine transaminase (ALT), and LDH concentrations (Reidarson and McBain, 1994; Reidarson et al., 1998). Fluconazole is considered less likely to cause inappetence than ketoconazole or itraconazole when administered to bottlenose dolphins (Reidarson et al., 1999). If inappetence occurs as a result of itraconazole administration, appetite can return by reducing the dose of itraconazole. Some advocate concomitant administration of prednisolone with ketoconazole to reduce the impact of inappetence. Flucytosine should be administered in a combination therapy with an azole to prevent resistance to flucytosine (Reidarson et al., 1999). Premature cessation of the azole may lead to flucytosine resistance (Poelma et al., 1974). Both drugs should be administered beyond the elimination of infection, as determined by physical examination, cytology, cultures, radiology, and endoscopy (Reidarson et al., 1999).
Antiparasitic Drugs Ivermectin has caused transient central nervous system abnormalities in small odontocetes (Townsend, 1999). Two fatalities in belugas occurred after intramuscular administration of levamisole (Boehm, pers. comm.). Both whales were treated with levamisole; only one was treated concomitantly with ivermectin. Fatalities occurring in both animals strongly suggested that levamisole alone was the cause of the mortalities. Other antiparasitic medications, such as the organophosphate dichlorvos, can cause neurological signs in marine mammals.
Steroids Steroid administration can have complex metabolic effects on marine mammals, particularly on electrolyte balance. Dexamethasone, prednisolone, or florinef administration reduces calcium and phosphate absorption and increases the urine output of calcium and potassium in terrestrial animals. Prolonged therapy could predispose an animal to hypocalcemia. Steroids also increase circulating serum glucose, triglyceride, and cholesterol concentrations. Dexamethasone administration in bottlenose dolphins can cause neutrophilia, lymphopenia, eosinopenia, elevated insulin, depressed ACTH and cortisol concentrations, and enhanced appetite (Reidarson and McBain, 1999). These changes in hematology and serum chemistry may return to normal upon cessation of steroid therapy. Supplemental vitamin D, folate, ascorbic acid, and pyridoxine may be appropriate during prolonged steroid administrations, because serum content of these vitamins can be depleted. Although estrogen therapy is not common in marine mammals, the seasonal or iatrogenically induced cycling of females should be considered when evaluating steroid therapy (Kirby, 1990; Schroeder, 1990). High estrogen levels will increase the anti-inflammatory effects of steroids by approximately 20-fold (Hansten, 1985). Corticosteroids will not be metabolized properly in animals undergoing estrogen therapy (Hansten, 1985).
Diuretics The potassium depletion caused by furosemide administration may be exacerbated by concurrent steroid administration. This is particularly a problem if sodium intake is high, as is often the case in marine mammals being fed supplemental salt in the diet. If a diuretic is indicated in a marine mammal receiving steroid therapy, alternatives to furosemide should be considered.
0839_frame_C31 Page 709 Tuesday, May 22, 2001 1:37 PM
709
Pharmaceuticals and Formularies
Chronic stress, hyperadrenocorticism, and other conditions resulting in high levels of endogenous corticosteroids can cause similar problems. Furosemide may also increase the ototoxicity of gentamicin. The nephrotoxicity of gentamicin, as well as of cephalosporins, is also exacerbated with concurrent furosemide administration. Additionally, furosemide diuresis results in increased renal loss of thiamine and pyridoxine, which can be important in situations where nutrition or oral supplementation is marginal.
Drug Dosages Some published drug dosages, and dosages used by the authors, are listed in Tables 1 through 5. When reading these tables, it is important to remember that these drugs have only been used in a limited number of individuals, and have not been exhaustively tested for efficacy or potential side effects. The tables are merely compiled so that readers have easy access to existing information used by practitioners, and can know where to add information for future editions of this book. TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) Drug Acetylpromazine Altrenogest Amikacin
Dosage
Species/Comments
100 mg/m body length IV 0.044 mg/kg 20 d postpartum 15 mg/kg IM SID
Premed for euthanasia
Needham, 1993
Killer whale (Orcinus orca) Nocardiosis in beluga (Delphinapterus leucas) Small odontocetes Small odontocetes Killer whale Bottlenose dolphin (Tursiops truncatus) Bottlenose dolphin Short-finned pilot whale (Globicephala macrorhyncus) Beluga Beluga Killer whale Killer whale Bottlenose dolphin Small odontocetes Killer whale Bottlenose dolphin Beluga a Zygomycosis tx in bottlenose dolphin Bottlenose dolphin
Young and Huff, 1996
14 mg/kg IM SID 7 mg/kg IM BID 4.8 mg/kg BID 14 mg/kg SID 7 mg/kg BID 5.8 mg/kg BID
Amoxicillin
Amoxicillin/Clavulanic acid
Amphotericin-B
Ampicillin
16.4 mg/kg SID 7.7 mg/kg BID 5 mg/kg PO BID 2.5 mg/kg PO BID 5 mg/kg PO BID 5–10 mg/kg PO BID 7 mg/kg PO BID 5–10 mg/kg PO BID 22 mg/kg PO BID 1–2 mg/kg PO SID (liposomal) 1–2 mg/kg PO SID (microencapsulated) 2.5 g cumulative dose 2.25 mg/kg PO BID
Killer whale
Reference
Robeck et al., 1996
Townsend, 1999 Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs.
McBain, unpubl. obs. McBain, unpubl. obs. Dunn et al., 1982 McBain, unpubl. obs. McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Townsend et al., 1996 Reidarson et al., 1999
Sweeney, 1985 (Continued)
0839_frame_C31 Page 710 Wednesday, May 23, 2001 12:26 PM
710
CRC Handbook of Marine Mammal Medicine
TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) (continued) Drug
Dosage
Species/Comments
Ascorbic acid Atropine
10 mg/kg PO BID 8 mg/kg PO 0.2 mg/kg
Killer whale Bottlenose dolphin
Azithromycin
6.7 mg/kg loading
Killer whale, bottlenose dolphin, beluga Caused inappetence and abdominal discomfort in beluga calves
3.7 mg/kg maintenance
Reference McBain, unpubl. obs. Colgrove et al., 1975 Geraci and Sweeney, 1986 Dalton et al., 1995; McBain, unpubl. obs. Dalton et al., 1995
9.6 mg/kg loading PO
Small odontocetes, bottlenose dolphin
McBain, unpubl. obs. Townsend, 1999
5.3 mg/kg maintenance PO SID
Small odontocetes, bottlenose dolphin Killer whale Killer whale
Ceftriaxone
3.7 mg/kg loading PO 1.7 mg/kg maintenance PO BID 4 mg/kg PO q3 d × 5 doses 20 mg/kg SID IM
McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs.
Small odontocetes, bottlenose dolphin, beluga
Townsend, 1999
Cefuroxime
20 mg/kg BID PO
Small odontocetes, bottlenose dolphin, beluga Killer whale Commerson’s dolphin (Cephalorhyncus commersoni) Small odontocetes, bottlenose dolphin Killer whale Bottlenose dolphin Possible intestinal bleeding after 20 d Small odontocetes Killer whale Bottlenose dolphin Commerson’s dolphin Short-finned pilot whale and beluga
McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs.
Bithionol
10 mg/kg BID 25 mg/kg BID
Carbenicillin
Cephalexin monohydrate
22–44 mg/kg TID 11 mg/kg TID 24 mg/kg PO BID
22 mg/kg PO TID 11 mg/kg PO TID 22 mg/kg PO TID 33 mg/kg PO TID 15 mg/kg PO TID Cephloridine Chloramphenicol Chlordiazepoxide HCl
6.6 mg/kg IT 22 mg/kg PO BID 0.5 mg/kg IM
Cimetidine
6 mg/kg PO TID 2100 mg PO QID 4.5 mg/kg PO BID
Sweeney, 1986
Killer whale Small odontocetes
Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. Colgrove et al., 1975
Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Sweeney, 1977 Sweeney, 1986a Geraci and Sweeney, 1986 Sweeney, 1986b Hoey et al., 1982 Townsend, 1999
0839_frame_C31 Page 711 Tuesday, May 22, 2001 1:37 PM
711
Pharmaceuticals and Formularies
TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) (continued) Drug Ciprofloxacin
Clindamycin
Copper sulfate Dexamethasone Dichlorvos
Dosage 8–13 mg/kg PO BID 15–29 mg/kg PO BID 6–9 mg/kg PO BID 7.7–9.6 mg/kg PO BID
Killer whale Bottlenose dolphin Beluga whale Bottlenose dolphin
4.5–5.5 mg/kg PO BID 11 mg/kg PO BID 4.4–7.7 mg/kg PO BID
Killer whale Commerson’s dolphin Short-finned pilot whale Beluga whale
7.7 mg/kg PO BID 4 ppm bath immersion 0.11 mg/kg PO
Dihydrostreptomycin Dimercaptosuccinic acid Doxycycline
13.2–16.5 mg/kg PO, repeat in 7–10 d 11 mg/kg IM SID 11 mg/kg PO BID 5–7 d, 9 cycles 1.5 mg/kg PO BID
Enrofloxacin
5 mg/kg PO SID 5 mg/kg BID 2.5 mg/kg PO BID 4.5 mg/kg PO BID
Epinephrine Erythropoietin Fenbendazole Florfenicol
Fluconazole
Species/Comments
0.02 mg/kg IM 63 U/kg twice 48 h apart IM 11 mg/kg PO 20 mg/kg IM q48 h, <20ml per site 2.8 mg/kg PO SID 2 mg/kg PO BID 2 mg/kg PO BID
Flucytosine
20 mg/kg PO TID
Folic acid
10 mg BID 25–50 mg BID
Furosemide Gentamicin
2–4 mg/kg IM 4 mg/kg IM SID 1.1 mg/kg IT 5 mg/kg IM BID
Bottlenose dolphin
Lead chelation in bottlenose dolphin Small odontocetes, bottlenose dolphin Bottlenose dolphin Small odontocetes, Killer whale, beluga Short-finned pilot whale Small odontocetes Rough toothed dolphin (Steno bredanensis) Small odontocetes Bottlenose dolphin, beluga whales; elevated AST, LDH Histoplasmosis tx in bottlenose dolphin Bottlenose dolphin Small odontocetes Candidiasis tx in bottlenose dolphin; itraconazole adjunct, never use as monotherapy Dose with TMP-S Bottlenose dolphin, dose with TMP-S Small odontocetes Use with caution, nephrotoxicity common
Reference McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Needham, 1978 Reidarson and McBain, 1999 Sweeney, 1986b Needham, 1978 Stetter et al., 1999 Townsend, 1999 McBain, unpubl. obs. Linnehan et al., 1999 McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. Townsend, 1999 Manire and Rhinehart, 2000 Townsend, 1999 Dalton and Robeck, 1998 Jensen et al., 1998 McBain, unpubl. obs. Reidarson et al., 1999 Townsend, 1999 McBain, unpubl. obs. Reidarson and McBain, 1995 Reidarson et al., 1999 Townsend, 1999 McBain, unpubl. obs. Townsend, 1999 Colgrove et al., 1975 Sweeney, 1977 Needham, 1978 (Continued)
0839_frame_C31 Page 712 Wednesday, May 23, 2001 10:59 AM
712
CRC Handbook of Marine Mammal Medicine
TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) (continued) Drug
Dosage 2.5 mg/kg PO TID
Human chorionic gonadotropin Imipenem Itraconazole
1000–3000 IU IM SID for 5 d 7.7–11.6 mg/kg BID 2.5 mg/kg PO BID
4.6 mg/kg 5 mg/kg PO BID
2.5–5 mg/kg PO BID 5 mg/kg PO BID 2.5 mg/kg PO BID 2.5 mg/kg PO BID 1.25 mg/kg PO BID Ivermectin
200 µg/kg PO
Ketoconazole
6 mg/kg PO 18 mg/kg PO 4–7 mg/kg PO, gradual increase to 10– 16 mg/kg 5 mg/kg PO BID
5 mg/kg PO BID 1.9 mg/kg PO BID Leuprolide acetate
0.075 mg/kg IM q28 d
Lidocaine HCl
10–20 ml 2%
Megestrol acetate Meperidine
0.3–0.5 mg/kg PO BID 2 mg/kg IM 1 mg/kg IM 0.5–1 mg/kg PO 2 mg/kg IM
Metronidazole
7 mg/kg PO TID
Species/Comments Duodenitis tx in rough toothed dolphin Bottlenose dolphin, induce ovulation Beluga Candidiasis tx in bottlenose dolphin, flucytosine adjunct Lobomycosis tx in bottlenose dolphin Aspergillosis tx in bottlenose dolphin associated 2- to 25-fold increase in liver enzymes Bottlenose dolphin Commerson’s dolphin Beluga Small odontocetes Killer whale, shortfinned pilot whale Crassicauda tx in small odontocetes, associated transient CNS signs Lobomycosis tx Candidiasis tx
Reference Townsend and Petro, 1998 Schroeder, 1993 McBain, unpubl. obs. Reidarson and McBain, 1995 Dalton et al., 1992 Reidarson et al., 1998
Reidarson et al., 1999 Reidarson et al., 1999 Reidarson et al., 1999 Townsend, 1999 McBain, unpubl. obs. Reidarson et al., 1999 Townsend, 1999
Sweeney, 1986a Dudok van Heel, 1977 Nakeeb et al., 1980 Schroeder, 1983b
Bottlenose dolphin, adjunct prednisolone
McBain, unpubl. obs.
Small odontocetes Beluga, adjunct prednisolone Bottlenose dolphin
Reidarson et al., 1999 Townsend, 1999 Reidarson et al., 1999 Briggs, 1995
For infra-alveolar nerve block in bottlenose dolphins
Ridgway et al., 1975
Small odontocetes Killer whale Killer whale Bottlenose dolphin, beluga Small odontocetes, bottlenose dolphin
Sweeney, 1989 Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Townsend, 1999 McBain, unpubl. obs.
0839_frame_C31 Page 713 Tuesday, May 22, 2001 3:34 PM
713
Pharmaceuticals and Formularies
TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) (continued) Drug Minocycline
Nystatin
Dosage 4 mg/kg loading 2 mg/kg maintenance BID 600,000 IU PO TID 600,000 IU PO TID
Ofloxacin
5 mg/kg PO BID
Pepto-Bismol® Phenytoin Praziquantel
Scale to humans PO See diphenylhydantoin 10 mg/kg 3 mg/kg
Prednisolone Prednisolone sodium succinate Pregnant mare serum gonadotropin
Procaine/benzathine penicillin
0.01 mg/kg PO SID 1–10 mg/kg IM, IV 1200 IU on day 1 IM; day 3, 400 IU PMSG; day 7, 1000 IU HCG 1500 IU on day 1 IM, repeat 48 h 10–20,000 IU/kg IM
Procaine penicillin G
47,000 IU/kg IM
Ranitidine Rifampin
2 mg/kg PO BID 2.5 mg/kg PO BID
Streptomycin Sucralfate
2.2 mg/kg PO BID 11 mg/kg IM SID 1 g PO QID
Tetracycline
Thiamine
Trimethoprimsulfadiazine
1–2 g PO BID-QID 6.7 mg/kg PO BID 22–35 mg/kg PO BID 55–65 mg/kg PO BID 77 mg/kg PO BID 55 mg/kg PO BID 1 mg/kg IM SID 2–4 mg/Kcal feed PO SID 25–35 mg/kg fish PO SID 30 mg/kg PO SID 15.7 mg/kg q48 h
Species/Comments
Reference
Small odontocetes, bottlenose dolphin, and beluga
Townsend, 1999 McBain, unpubl. obs.
Duodenitis tx in rough toothed dolphin Small odontocetes, bottlenose dolphin
McBain, unpubl. obs. Dunn et al., 1982 Townsend and Petro, 1998 Townsend, 1999 Townsend, 1999
Nasitrema tx in small odontocetes Tapeworm tx in small odontocetes Adjunct ketoconazole For tx of shock in small odontocetes Bottlenose dolphin; see Chapter 11, Reproduction
Townsend, 1999 Townsend, 1999 Reidarson et al., 1999 Townsend, 1999 Schroeder, 1983a
Schroeder, 1993 Occasionally used every other day SID for leptospirosis in California sea lions For diarrhea Small odontocetes Small odontocetes, bottlenose dolphin, and beluga Killer whale
McBain, unpubl. obs. Gulland, pers. comm. Sweeney, 1986a Colgrove et al., 1975 Townsend, 1999 Townsend, 1999 McBain, unpubl. obs.
Killer whale Bottlenose dolphin Commerson’s dolphin Beluga Follow by oral 2 h before feeding
McBain, unpubl. obs. Needham, 1978 Townsend and Petro, 1998 Townsend, 1999 McBain, 1985 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Geraci, 1986 Geraci, 1986
At main feeding
Geraci, 1986
Bottlenose dolphin Beluga calf
Schroeder et al., 1984 Cook et al., 1992
Duodenitis tx in rough-tooth dolphin Small odontocetes
(Continued)
0839_frame_C31 Page 714 Tuesday, May 22, 2001 1:37 PM
714
CRC Handbook of Marine Mammal Medicine
TABLE 1 Drug Dosages Reported for Cetaceans (see text for precautions) (continued) Drug
Dosage 16–22 mg/kg PO SID
22 mg/kg PO SID 32 mg/kg IM SID
Bottlenose dolphin, small odontocetes; give with folic acid; may cause fatal pancytopenia Killer whale Commerson’s dolphin Short-finned pilot whale Beluga Myositis, but efficacious
50 mg/kg PO TID
May cause diarrhea
1–1.5 mg/kg TID PO 1.1 mg/kg BID PO 100 IU/kg fish PO SID
Small odontocetes Bottlenose dolphin Killer whale
7.7–11 mg/kg PO SID 16 mg/kg PO SID 7.7 mg/kg PO SID
Tylosin
Vancomycin Vitamin E
Species/Comments
Reference McBain, unpubl. obs.
Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Thurman and Windsor, 1984 Thurman and Windsor, 1984 Townsend, 1999 McBain, unpubl. obs. McBain, unpubl. obs. Geraci, 1986
Note: tx = treatment.
TABLE 2 Drug Dosages Reported for Pinnipeds (see text for precautions) Drug
Dosage
Acetylpromazine Acetylcysteine Acetylcysteine + isoproterenol
Contraindicated 20% solution nebulized BID-QID 400 mg IT BID-QID
Aluminum hydroxide
30–90 mg/kg PO
Amikacin
7 mg/kg BID 7.7 mg/kg BID
Aminophylline
5.5 mg/kg IV, IM, PO; BID, TID
Amoxicillin
22 mg/kg PO BID 20 mg/kg IV single dose
Amoxicillin/clavulanic acid Ampicillin
Species/Comments
22 mg/kg PO BID 22 mg/kg IM, IV, PO TID
Nebulized in 12–15 ml saline with 1:50,000 isoproterenol Leptospirosis tx in California sea lion (Zalophus californianus) to bind phosphorus California sea lion Walrus (Odobenus rosmarus) California sea lion, harbor seal (Phoca vitulina), northern elephant seal (Mirounga angustirostris) California sea lion, harbor seal, elephant seal Northern elephant seal, harbor seal California sea lion, harbor seal, elephant seal
Reference Dierauf, in Stoskopf 1990 Dierauf, in Stoskopf, 1990 Sweeney, 1986b
Gulland, 1999
McBain, unpubl. obs. McBain, unpubl. obs. Dierauf, in Stoskopf, 1990
Gulland, pers. comm. Gulland et al., 2000 Gulland, pers. comm. Haulena, pers. comm.
0839_frame_C31 Page 715 Tuesday, May 22, 2001 1:37 PM
715
Pharmaceuticals and Formularies
TABLE 2 Drug Dosages Reported for Pinnipeds (see text for precautions) (continued) Drug
Dosage Ophthalmic TID-QID
Bromsulphalein Castor oil/dry mustard Cefuroxime Cephalexin Cephaloridine Chloramphenicol
Contraindicated 1.2 ml/kg of 4:1 ratio PO 10–15 mg/kg PO BID 22 mg/kg PO TID 8.8 mg/kg IT BID 4.1 mg/kg IV, PO TID
Cimetidine
20–30 mg/kg PO BID, TID 5–15 mg/kg PO TID
Ciprofloxacin
Ophthalmic TID-QID
Clindamycin Dexamethasone
7.5 mg/kg PO BID 8–11 mg/kg PO BID 7.3 mg/kg PO BID 0.2–1.0 mg/kg IM, PO SID 2.2 mg/kg IV
Dextrose (5% in LRS)
100 ml/kg/d SQ or RO
Dichlorvos
9.7–11.5 mg/kg tablet PO
Disophenol
Enrofloxacin Erythromycin Fenbendazole Fluconazole
Fluoxitine HCl (5-HT reuptake inhibitor)
29.3–32.8 mg/kg capsule PO 9.9 mg/kg SC BID, SID 12.5 mg/kg SC 2.5–5 mg/kg PO BID 3.3 mg/kg PO BID 5.5 mg/kg PO BID 11 mg/kg PO SID × 2 doses 0.5 mg/kg PO BID Ophthalmic TID-QID
0.2–1.2 mg/kg PO SID
Species/Comments Keratitis tx in harbor seal, subpalpebral lavage with ciprofloxacin and fluconazole Unless IV catheter is used
Walrus California sea lion California sea lion Harbor seal California sea lion California sea lion with leptospirosis Keratitis tx harbor seal, subpalpebral lavage with fluconazole and atropine Walrus California sea lion Walrus California sea lion and elephant seal For tx of shock in California sea lion California sea lions undergoing leptospirosis tx Northern fur seal (Callorhinus ursinus); caution organophosphate toxicity Northern fur seal, caution organophosphate toxicity Northern fur seal Northern fur seal, associated diarrhea California sea lion Walrus Northern elephant seal, California sea lion Keratitis tx in harbor seal, subpalpebral lavage with ciprofloxacin and atropine Tx for regurgitation and flank sucking in California sea lion
Ref. Borkowski et al, 1999
Needham, 1978 Geraci and Sweeney, 1986 McBain, unpubl. obs. McBain, unpubl. obs. Sweeney, 1977 Koski and Vandenbroek, 1986 McBain, unpubl. obs. Gulland, pers. com. Borkowski et al., 1999
McBain, unpubl. obs. McBain, unpubl. obs. McBain, unpubl. obs. Gage et al., 1985 Dierauf, in Stoskopf, 1990 Gulland, 1999
Bigg and Lyons, 1981
Lyons et al., 1978 Lyons et al., 1978 Lyons et al., 1980 McBain, unpubl. obs. McBain, unpubl. obs. Sweeney, 1974b Gage et al., 1985 Reidarson et al., 1999 Borkowski et al., 1999
Dalton et al., 1997
(Continued)
0839_frame_C31 Page 716 Tuesday, May 22, 2001 1:37 PM
716
CRC Handbook of Marine Mammal Medicine
TABLE 2 Drug Dosages Reported for Pinnipeds (see text for precautions) (continued) Drug Furosemide Gentamicin Griseofulvin
Dosage
Species/Comments
Reference
2.5–5.0 mg/kg IM, IV, PO BID 0.75 mg/kg IT BID 2 d, then SID 15 mg/kg PO SID 45 d 5000 mg/d PO 4 w
Harbor seal, California sea lion
Haulena, pers. comm.
Haloperidol Hydrogen peroxide
0.11 mg/kg PO BID 5 ml/kg PO PRN
Indomethacin
Isoniazid
0.1–0.3 mg/kg PO initial dose 0.1–0.2 mg/kg 12–24 h 0.45 mg/kg 48 h 90 mg nebulized in 1% soln. PRN 2 mg/kg PO SID
Isoproterenol Isuprel
1:50,000 IT 0.4 mg/kg PO BID, TID
Isoetharine
Itraconazole
Ivermectin
Ketoconazole
Leuprolide acetate Levamisole
Megestrol acetate Metronidazole Neomycin Nystatin Oxytocin
0.5 ml nebulized BID-QID 1.5–2 mg/kg PO SID 0.5–1 mg/kg PO BID 1.5–2 mg/kg PO SID 100–200 µg/kg SC 200 µg/kg SC
10 mg/kg IM, PO SID 4.4 mg/kg PO BID 1 mg/kg PO BID 4.4 mg/kg PO BID 0.09–0.12 mg/kg IM q28 d 15 mg/kg SC 15 mg/kg SC 8 mg/kg PO 0.3–0.5 mg/kg PO BID 10 mg/kg PO TID 20 mg/kg PO TID 600,000 U PO TID 20–40 USP units IM
Sweeney, pers. comm.
Australian sea lion (Neophoca cinerea) California sea lion
California sea lion, northern elephant seal
For tx of bronchospasm Gray seal (Halichoerus grypus) California sea lion California sea lion Walrus Other pinnipeds Walrus Northern fur seal California sea lion, northern elephant seal, harbor seal; tremors observed in a Guadalupe fur seal (Arctocephalus townsendii) after administration Monitor liver enzymes Walrus Other pinnipeds Walrus California sea lions, may cause injection site pain California sea lion Northern elephant seal California sea lion
California sea lion California sea lion
Farnsworth et al., 1975 Phillips et al., 1986 Dalton et al., 1997 Geraci and Sweeney, 1986b Dierauf, in Stoskopf, 1990
Dierauf, in Stoskopf, 1990 Stoskopf, unpubl. obs. Sweeney, 1986b Dierauf, in Stoskopf, 1990 Dierauf, in Stoskopf, 1990 Reidarson et al., 1999 Reidarson et al., 1999 McBain, unpubl. obs. Beekman, 1984 Gage et al., 1985 Gulland and Haulena, pers. comm.
Dunn et al., 1984 Reidarson et al., 1999 Reidarson et al., 1999 McBain, unpubl. obs. Calle et al., 1997 Gage et al., 1985 Gage et al., 1985 Dalton and Robeck, 1998 Sweeney, 1989 McBain, unpubl. obs. McBain, unpubl. obs. Dunn et al., 1982 Schroeder, 1993
0839_frame_C31 Page 717 Wednesday, April 10, 2002 8:07 AM
717
Pharmaceuticals and Formularies
TABLE 2 Drug Dosages Reported for Pinnipeds (see text for precautions) (continued) Drug
Dosage
Penicillin-G
9090 IU/kg PO BID
Penicillin benzathine/procaine Penicillin (benzathine) Phenobarbital
4545–9090 IU/kg IM SID 30,000 IU/kg IM SID 1.0–1.5 mg/kg PO SID-BID See diphenylhydantoin 110 mg/kg PO 10 mg/kg PO SID
Phenytoin Piperazine Praziquantel Primidone Profenal
Propriopromazine Pyridoxine Rifampin Ronnel Sodium chloride Tetracycline
Theophylline
1–2.5 mg/kg PO SID 2.5 mg/kg PO TID Ophthalmic BID
1–2 mg/kg IM 0.25 mg/kg PO SID 5 mg/kg PO SID Manufacturer’s directions 3 g/kg of fish SID 100–200 mg/kg PO, IP 12.5 mg/kg PO SID 22 mg/kg PO SID 4.5 mg/kg PO TID 22 mg/kg PO BID 44 mg/kg PO BID 8.9 mg/kg loading
Species/Comments California sea lion
California sea lion Tx of idiopathic seizures in California sea lion
California sea lion, harbor seal, elephant seal Maintenance dose Initial control For tx of corneal opacities in Hawaiian monk seals (Monachus schauinslandi)
Thiamine HCl
Trimethoprimsulfadiazine
Vitamin A
0.44 mg/kg IV slow drip, twice 1 mg/kg IM SID 2–4 mg/Kcal feed PO SID 25–35 mg/kg fish PO SID 4.5 mg/kg PO BID
Vanderbroek et al., 1985 Vanderbroek et al., 1985 Gage et al., 1985 Gage, 1999
Sweeney, 1974 Gage et al., 1985; Gulland pers. comm. Needham, 1978 Needham, 1978 Braun et al., 1996
Geraci, 1986 Stoskopf et al., 1987 Stoskopf et al., 1987 Sweeney, 1974 For maintenance
California sea lion California sea lion Walrus
Geraci, 1986 Geraci, 1972 Farnsworth et al., 1975 Vanderbroek et al., 1985 Gage et al., 1985 McBain, unpubl. obs. McBain, unpubl. obs. MacDonald and Dougherty, 1983 MacDonald and Dougherty, 1983 Sweeney, 1986b
Follow with oral 2 h before feeding
Geraci, 1986 Geraci, 1986
At feeding
Geraci, 1986
Harbor seal
Koski and Vandenbroek, 1986 Vanderbroek et al., 1985 McBain, 2000 McBain, 2000 Mazzaro et al., 1995b
6.1 mg/kg maintenance Thiacetarsamide
Reference
3.6 mg/kg PO BID
California sea lion
22–30 mg/kg PO SID 10–13 mg/kg PO SID 300–600 IU/d PO
California sea lion Walrus Northern fur seal
(Continued)
0839_frame_C31 Page 718 Tuesday, May 22, 2001 1:37 PM
718
CRC Handbook of Marine Mammal Medicine
TABLE 2 Drug Dosages Reported in Pinnipeds (see text for precautions) (continued) Drug Vitamin E
Dosage 100 IU/kg PO in fish 100 mg/d PO
Species/Comments High levels (50,000 IU/D) may increase vit E req. In fish Harbor seals, adult males
Reference Mazzaro et al., 1995a Geraci, 1986 Mazzaro et al., 1995c
Note: tx = treatment.
TABLE 3 Drug Dosages Reported for Sirenians (see text for precautions) Drug
Dosage
Amikacin
7 mg/kg IM BID
Ampicillin
5.5 mg/kg PO SID
Ascorbic acid
1 mg/kg PO SID
Ceftriaxone
22 mg/kg IM SID
Cephalexin
40 mg/kg PO SID
Dexamethasone
2.2 mg/kg IM
Fenbendazole
10 mg/kg PO
Gentamicin
4.4 mg/kg IM SID 2.5 mg/kg PO TID
Itraconazole Ivermectin Metronidazole
2.5 mg/kg PO BID 200 µg/kg PO 7 mg/kg PO BID
Mineral oil Oxytetracycline
2–3 ml/kg up to 1.5 l 15 mg/kg IM BID 4.5 mg/kg IM BID
Species/Comments Manatee (Trichechus manatus)
At main feeding
Hemorrhagic colitis tx in manatees, adjunct to metronidazole Manatee Hemorrhagic colitis tx in manatees; adjunct to gentamycin For tx of constipation Manatee Dugong (Dugong dugon)
Penicillin G/benzathine Penicillin, benzathine Penicillin, procaine Penicillin/streptomycin
Cattle dosage 22,000 IU/kg 25,000 IU/kg IM SID 25,000 IU/kg SC 1000 IU/kg IM
Praziquantel
8–16 mg/kg PO
Manatee Dugong Dugong Manatee, adjunct to streptomycin For tx of trematodes
Sulfasalazine Tetracycline Thiamine
10 mg/kg IM BID 55 mg/kg IM BID 1 mg/kg IM SID
Manatee Manatee Follow by oral
Reference Walsh and Bossart et al., 1999 White, in Stoskopf, 1990 White, in Stoskopf, 1990 Walsh and Bossart et al., 1999 White, in Stoskopf, 1990 White, in Stoskopf, 1990 Walsh and Bossart et al., 1999 White, in Stoskopf, 1990 Walsh et al., 1999
Bossart, see Chapter 43 Walsh and Bossart, 1999 Walsh et al., 1999
Walsh and Bossart, 1999 White, in Stoskopf, 1990 Elliot et al., 1981 Walsh and Bossart, 1999 Walsh and Bossart, 1999 Cohen, 1993 Elliot et al., 1981 White, in Stoskopf, 1990 Walsh and Bossart et al., 1999 Bossart, see Chapter 43 Bossart, see Chapter 43 Geraci, 1986
0839_frame_C31 Page 719 Tuesday, May 22, 2001 1:37 PM
719
Pharmaceuticals and Formularies
TABLE 3 Drug Dosages Reported for Sirenians (see text for precautions) (continued) Drug
Vitamin E
Dosage 2–4 mg/Kcal feed PO SID 25–35 mg/kg fish PO SID 100 IU/kg fish PO SID
Species/Comments
Reference
2 h before feeding
Geraci, 1986
At main feeding
Geraci, 1986 Geraci, 1986
Note: tx = treatment.
TABLE 4 Drug Dosages Reported for Sea Otters (Enhydra lutris) (see text for precautions) Drug Acetylsalicylic acid Amikacin Aminopentamidine sulfate Aminophylline Aminopropazine fumarate Amoxicillin Ascorbic acid Atropine Carprofen Cefazolin Cephalexin Charcoal, activated Cimetidine
Cyproterone Dawn liquid detergent
Dosage 10 mg/kg PO q36 h 5 mg/kg IM BID, or 10 mg/kg SID 0.1–0.4 mg/kg IM BID 10 mg/kg PO BID 2 mg/kg IM, SC, PO BID 10–20 mg/kg PO QID 50–100 mg PO, IM, SC SID 0.02–0.04 mg/kg IM 1.5–2 mg/kg PO BID for 5–10 d 10–30 mg/kg IM QID 20 mg/kg PO BID 100 ml tube feed 5–10 mg/kg IV, IM, SC, PO BID 5 mg/kg IM TID
Deslorelin Dexamethasone
1.5–2.3 mg/kg PO SID 1:16 dilution, 4–8 l topically 0.18–0.23 mg/kg SC 2 mg/kg IV, IM
Dextrose 10%
To effect IV
Dextrose 5% in LRS
IP
Diphenhydramine Diphenoxylate Diphenylhydantoin
0.5–2 mg/kg PO BID 0.1–0.2 mg/kg PO BID 20–30 mg/kg PO BID
Enrofloxacin Folic acid Furosemide
5–20 mg/kg SID 2.5 mg PO 2 mg/kg IM
Species/Comments Nephrotoxicity reported
Reference Williams, 1990 Williams, 1990; Murray, pers. comm. Williams et al., 1995a Williams et al., 1995a Williams, 1990 Williams et al., 1995b Williams, 1990
For relief of injection site pain Pups Adults Reversible increase in liver enzymes Pups, hemorrhagic diarrhea To remove oil
Repeat doses with caution Hypoglycemic seizures tx Hypoglycemic seizures tx
Antiepileptic; phenobarb preferred
Murray, pers. comm. Calle et al., 1999 Murray, pers. comm. Williams, 1993 Williams, 1993 Murray, pers. comm. Williams, 1993 Calle et al., 1999 Williams, 1993 Calle et al., 1999 Williams et al., 1995b Williams, 1993 Williams, 1993 Williams, 1990 Williams et al., 1995a Williams, 1990 Murray, pers. comm. Williams, 1990 Williams et al., 1995a (Continued)
0839_frame_C31 Page 720 Tuesday, May 22, 2001 1:37 PM
720
CRC Handbook of Marine Mammal Medicine
TABLE 4 Drug Dosages Reported for Sea Otters (Enhydra lutris) (see text for precautions) (continued) Drug Gentamicin
Griseofulvin Hetacillin Hydrocortisone Insulin (NPH) Isoproterenol Ivermectin Kaopectate Ketoconazole Lactated Ringer’s
Leuprolide acetate
Levamisole Lidocaine Lincomycin Medroxyprogesterone acetate Methylprednisolone Metoclopramide HCl Metronidazole Neomycin Nitrous oxide Oxacillin Oxytocin Penicillin G
Phenobarbital Phenylephrine Phenytoin Pitressin Praziquantel Ranitidine Selenium (Seletoc) Sodium bicarbonate Sodium iodine Stanozolol
Dosage 4.4 mg/kg IM BID 2 mg/kg IM BID 5 d 2 mg/kg IM TID 30 mg/kg PO BID 45 d 20 mg/kg PO BID 50 mg/kg IV, 5–150 mg/kg 2 IU/kg (adjust) SC BID 0.1–0.2 mg IM, SC SID 50–300 mg/kg PO 2 ml/kg PO Q6 h 10 mg/kg PO TID 40–50 ml/kg/d IV, SC, IP 66 ml/kg SC 500 ml SC 0.11–0.19 mg/kg IM q28 d 0.9–1.1 mg/kg IM, SC q4 m 15 mg/kg 7–10 d PO SID 2 mg/kg bolus, repeat in 20 min 20 mg/kg IM BID 75 mg/kg IM q21 d × 3 doses 0.06 mg/kg/day IM or IV 0.2 mg/kg IM BID 25–30 mg/kg PO BID 5d 10–14 mg/kg PO SID 1 l/min IH 20 mg/kg IM TID 10–20 USP units IV, IM 22,000 IU/kg IM BID 20,000 IU/kg IM SID 1 mg/kg IV PRN 0.1 mg/kg IV, IM See diphenylhydantoin 2.5–5 IU IV, IM q2 d 6 mg/kg IM 1–4 mg/kg PO TID 0.1 mg/kg IV, IM 1 mEq/kg IV 0.2 ml/kg 20% soln. IV, PO BID 10–25 mg IM q7 d
Species/Comments Nephrotoxicity reported Pups Adults
For shock To effect Once
Pups Adults Injection site pain, lameness 4-month depot formulation
Reference Williams, 1990 Williams, 1993 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1990 Williams, 1993 Williams, 1993 Calle et al., 1997 Calle et al., 1999 Williams, 1990 Williams, 1990
In males to decrease sex drive Pups, hemorrhagic diarrhea
Williams, 1990 Williams, 1990 Williams et al., 1995a Williams, 1990 Williams, 1990
As presurgical GI prep
For milk letdown
For seizures
In aqueous solution
Infuse slowly
Contraindicated in gravid females
Williams et al., 1995a Williams, 1990 Williams, 1990 Williams, 1990 Williams and Siniff, 1983 Williams, 1993 Williams, 1990 Williams, 1990 Williams, 1990 Williams et al., 1995a Williams et al., 1995b Williams et al., 1995b Williams, 1990 Williams, 1990 Williams, 1990
0839_frame_C31 Page 721 Tuesday, May 22, 2001 1:37 PM
721
Pharmaceuticals and Formularies
TABLE 4 Drug Dosages Reported for Sea Otters (Enhydra lutris) (see text for precautions) (continued) Drug
Dosage
Tetracycline
20 mg/kg IM SID
Thiamine
1mg/kg IM SID 2–4 mg/Kcal feed PO SID 25–35 mg/kg fish PO SID 33.6 mg/kg PO BID
Trimethoprimsulfadiazine
20 mg/kg IM BID Vitamin B complex Vitamin E Zinc chlorhexidate gel
2 ml/l LRS SC 1 ml/10 kg SC 100 IU/kg fish SID 400 IU/day 0.5 ml/ice cube SID
Species/Comments Can cause muscle necrosis Follow by oral 2 h before feeding
Reference Williams, 1990 Geraci, 1986 Geraci, 1986
At main feeding
Geraci, 1986 Calle et al., 1999
Hemorrhagic diarrhea tx, pups Pups Adults
Williams, 1993
Dental prophylaxis
Williams, 1993 Geraci, 1986 Williams et al., 1995b Young et al., 1999
Note: tx = treatment.
TABLE 5 Drug Dosages Reported for Polar Bears (Ursus maritimus) (see text for precautions) Drug Ampicillin Ascorbic acid Carbenicillin Chloramphenicol Dexamethasone Dichlorvos Doxycycline Fenbendazole Flumethasone Ivermectin Mebendazole Milbemycin oxime Niclosamide Nitrofurantoin Oxytetracycline Penicillin, benzathine Penicillin, procaine Penicillin (K) Pepto-Bismol Piperazine Potassium Prednisolone
Tetracycline Thiamine
Dosage 4–5 g PO TID 2–4 g PO SID 2.5 g PO QID 5 d 3–4 g PO BID, TID 2.5 g/l (4 l) topical spray q7 d 0.1 mg/kg IM 20 mg/kg PO 1 mg/kg PO SID 12 d 10 mg/kg PO SID × 2 doses 0.6–1.25 mg IM, PO SID 0.1 mg/kg IM q7 d × 2 doses 20 mg/kg PO 1 mg/kg PO 80 mg/kg PO 4–6 mg/kg PO TID 10 mg/kg IM BID 30,000 IU/kg IM q3 d 15,000 IU/kg IM q3 d 15,000 IU/kg IM q3 d 1 ml/kg PO QID 100 mg/kg PO 2.5 mEq PO BID 20 mg PO SID 1–5 mg/kg IV, IM SID 5 mg/kg PO q7 d 1 mg/kg IM SID 2–4 mg/kcal feed PO SID 25–35 mg/kg fish PO
Species/Comments
In 86-kg bear Adult bear for 6–10 d
Bovine preparation Balisascaris transfuga tx
Procaine and potassium Penicillin doses combined
In 86-kg bear For tx of allergies For tx of respiratory distress For tx of skin conditions Follow with oral 2 h before feeding At feeding
Reference Kuntze, 1986 Kuntze, 1986 Crawshaw, 1980 Elze et al., 1986 Kitchen et al., 1977 Stoskopf, 1988 Stoskopf, 1990 Smith, 1973 Stoskopf, 1990 Hoff, 1979 Kohm, 1986 Stoskopf, 1990 Hedberg et al., 1995 Stoskopf, 1990 Kuntze, 1986 Stoskopf, 1990 Newman et al., 1975 Newman et al., 1975 Newman et al., 1975 Stoskopf, 1990 Stoskopf, 1990 Crawshaw, 1980 Elze et al., 1986 Elze et al., 1986 Kitchen et al., 1977 Geraci, 1986 Geraci, 1986 Geraci, 1986 (Continued)
0839_frame_C31 Page 722 Tuesday, May 22, 2001 1:37 PM
722
CRC Handbook of Marine Mammal Medicine
TABLE 5 Drug Dosages Reported for Polar Bears (Ursus maritimus) (see text for precautions) (continued) Drug Thyroxine Thyroid-stimulating hormone Vitamin A Vitamin E
Dosage 3 mg PO BID 30 IU IM 1 million IU/d PO SID 20,000 IU/kg PO SID 600–2400 IU PO q30 d
Species/Comments Thyroid stimulation test As dietary supplement
Reference Hoff, 1979 Hoff, 1979 Foster, 1981 Kock et al., 1986 Elze et al., 1986
Note: tx = treatment.
Acknowledgments The authors thank Jeff Boehm and Beth Chittick for reviewing this chapter, and Mike Murray for comments on the sea otter formulary.
References Beekman, G.K., 1984, Clinical use of ivermectin against the hookworm (Uncinaria lucasi) of northern fur seal pups (Callorhinus ursinus), in Proceedings International Association for Aquatic Animal Medicine, 15: 50. Benet, L.Z., Oie, S., and Schwartz, J., 1996, Appendix II, Design and optimization of dosage regimens; Pharmacokinetic data, in The Pharmacological Basis of Therapeutics, 9th ed., Goodman, L., and Gilman, A. (Eds.), Macmillan, New York, 1707–1792. Bigg, M.A., and Lyons, E.T., 1981, Clinical observations on three northern fur seal pups treated with dichlorvos, J. Am. Vet. Med. Assoc., 179: 1284–1286. Borkowski, R., Moore, P.A., Mumford, S., Patchett, K., Rice, J., and Rubinstein, B., 1999, Extended use of subpalpebral lavage systems for treatment of keratitis in a harbor seal (Phoca vitulina), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 39–40. Bossart, G.D., and Dierauf, L.A., 1990, Marine mammal clinical laboratory medicine, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 1–52. Braun, R., Paulson, M., Omphroy, C., Heath, B., Gilmartin, W., Ragen, T., Breese, M., and Driscoll, C., 1996, Corneal opacities in Hawaiian monk seals, in Proceedings of the International Association for Aquatic Animal Medicine, 27: 97–99. Briggs, M.B, 1995, Effects of leuprolide acetate in depot suspension on testosterone levels, testicular size and semen production in male Atlantic bottlenose dolphins (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 26: 112–114. Burkhardt, J.E., 1996, Review of quinolone arthropathy in the dog, Chemotherapeutics, 4: 14–18. Burkhardt, J.E., Hill, M.A., and Carlton, W.W., 1990, Histologic and histochemical changes in articular cartilages of immature beagle dogs dosed with difloxacin, a fluoroquinolone, Vet. Pathol., 27: 162–170. Burkhardt, J.E., Walterspiel, N.N., and Schaad, U.B., 1997, Quinolone arthropathy in animals versus children, Clini. Infect. Dis., 25: 1196–1294. Calle, P.P., Stetter, M.D., Raphael, B.L., and Cook, R.A., 1997, Use of depot leuprolide acetate to control undesirable male associated behaviors in the California sea lion (Zalophus californianus) and California sea otter (Enhydra lutris), in Proceedings of the International Association for Aquatic Animal Medicine, 28: 6–7.
0839_frame_C31 Page 723 Tuesday, May 22, 2001 1:37 PM
Pharmaceuticals and Formularies
723
Calle, P.P., Raphael, B.L., Cook, R.A., McClave, C., Basinger, J.A., and Walters, H., 1999, Use of depot leuprolide, cyproterone, and deslorelin to control aggression in an all male California sea otter (Enhydra lutris nereis) colony, in Proceedings of the International Association for Aquatic Animal Medicine, 30: 42–45. Cohen, M.A., 1993, Successful treatment and release of a stranded dugong (Dugong dugon), in Proceedings of the International Association for Aquatic Animal Medicine, 24: 58. Colgrove, G.S., Sawa, T.R., Brown, J.T., McDowell, P.F., and Nachtigall, P.E., 1975, Necrotic stomatitis in a dolphin, J. Wildl. Dis., 11: 460. Cook, R.A., Calle, P.P., McClave, C., and Palma, S., 1992, Health care and medical problems of a captive bred and mother reared beluga whale (Delphinapterus leucas), in Proceedings of the International Association for Aquatic Animal Medicine, 23: 107–110. Cornell, L.H., 1978, Drug induced granulocytic leukopenia in cetaceans, Proceedings of the American Association of Zoo Veterinarians. Crawshaw, G.J., 1980, Acute renal failure in a polar bear, Proc. Am. Assoc. Zoo Vet., 11: 135–137. Dalton, L.M., and Robeck, T.R., 1998, Florfenicol serum levels in beluga whales (Delphinapterus leucas) and bottlenose dolphins (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 29: 37–38. Dalton, L.M., Mathey, W.S., Crawley, R.R., and Young, W.G., 1992, Lobomycosis in a bottlenose dolphin (Tursiops truncatus): Diagnosis and treatment, in Proceedings of the International Association for Aquatic Animal Medicine, 23: 112. Dalton, L.M, Robeck, T.R., and Campbell, T.W., 1995, Azithromycin serum levels in cetaceans, in Proceedings of the International Association for Aquatic Animal Medicine, 26: 23. Dalton, L.M., Robeck, T.R., and Young, W.G., 1997, Aberrant behavior in a California sea lion (Zalophus californianus), in Proceedings of the International Association for Aquatic Animal Medicine, 28: 137. Dudok van Heel, W.H., 1977, Successful treatment in a case of lobomycosis (Lobo’s disease) in Tursiops truncatus (Mont) at the Dolphinarium Harderwijk, Aquat. Mammals, 5: 8. Dunn, J.L., Buck, J.D., and Spotte, S., 1982, Candidiasis in captive cetaceans, J. Am. Vet. Med. Assoc., 181: 1316. Dunn, J.L., Buck, J.D., and Spotte, S., 1984, Candidiasis in captive pinnipeds, J. Am. Vet. Med. Assoc., 185: 1328. Elliot, H., Thomas, A., Ladds, P.W., and Heinsohn, G.E., 1981, A fatal case of salmonellosis in a dugong, J. Wildl. Dis., 17: 203. Elze, V.K., Krische, G., Eulenberger, K., Seifert, S., and Selbitz, H.J., 1986, Erkrankungen und Todesfälle beim Eisbar (Thalarctos maritimus Phipps, 1774), Erkrankungen Zootiere, 28: 51. Farnsworth, R.J., McKeever, P.J., and Fletcher, J.A., 1975, Dermatomycosis in a harbor seal caused by Microsporum canis, J. Zoo Anim. Med., 6: 26. Foster, J.W., 1981, Dermatitis in polar bears: A nutritional approach to therapy, Proc. Am. Assoc. Zoo Vet., 12: 58. Gage, L.J., 1999, Geriatric medicine in aged captive pinnipeds, in Proceedings of the International Association for Aquatic Animal Medicine, 30: 130–131. Gage, L.J., Vandenbroek, D., and Amaya, L., 1985, Common medical problems encountered in pinnipeds at the California Marine Mammal Center, in Proceedings of the International Association for Aquatic Animal Medicine, 16: 42. Geraci, J.R., 1972, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618. Geraci, J.R., 1986, Nutrition and nutritional disorders, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M. (Ed.), W.B. Saunders, Philadelphia, 568. 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. Gulland, F.M., 1999, Leptospirosis in marine mammals, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 469–471.
0839_frame_C31 Page 724 Tuesday, May 22, 2001 1:37 PM
724
CRC Handbook of Marine Mammal Medicine
Gulland, F.M., Stoskopf, M.K., Johnson, S.P., Riviere, J., and Papich, M.G., 2000, Amoxicillin pharmacokinetics in harbor seals (Phoca vitulina) and northern elephant seals (Mirounga angustirostris) following single dose intravenous administration: Implications for interspecific dose scaling, J. Vet. Pharmacol. Ther., 23: 223–228. Hansten, P.D., 1985, Drug Interactions, 5th ed., Lea & Febiger, Philadelphia, 448 pp. Hedberg, G.E., Bennett, R.A., and Dunker, F.H., 1995, Preliminary studies on the use of milbemycin oxime for treatment of polar bears (Ursus maritimus) with chronic Baylisascaris transfuga infection, in Proceedings, Joint Conference of American Association of Zoo Veterinarians, Wildlife Disease Association, and American Association of Wildlife Veterinarians, 299–301. Hoey, A., McBain, J., and Green, J.D., 1982, Incidental post mortem findings in a juvenile female Orcinus orca following accidental death, in Proceedings of the International Association for Aquatic Animal Medicine, 13: 9. Hoff, S., 1979, Skin diseases in two polar bears (Thalarctos maritimus) at the Stanley Park Zoo, Proc. Am. Assoc. Zoo Vet., 10: 107. Jensen, E.D., Van Bonn, W., Lipscomb, T., and Ridgway, S.H., 1998, Disseminated histoplasmosis in Atlantic bottlenose dolphins, in Proceedings of the International Association for Aquatic Animal Medicine, 29: 64. Johnsen, R.I., 1994, Effects of antibacterial agents on transformation of fatty acids in sediment from a marine fish farm site, Sci. Total Environ., 152: 143–52. Johnson, S.P., Nolan, S., and Gulland, F.M.D., 1998, Antimicrobial susceptibility of bacteria isolated from pinniped stranded in central and northern California, J. Zoo Wildl. Med., 29: 288–294. Kirby, V.L., 1990, Endocrinology of marine mammals, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 303–351. Kitchen, H., Dayhuff, K., and Ricketts, C., 1977, Diseases of hair in polar bears and Bengal tigers, Proc. Am. Assoc. Zoo Vet., 8: 118–120. Kock, R.A., Thomsett, L.R., and Henderson, G.M., 1986, Alopecia in polar bears (Thalarctos maritimus), a report of two cases, Erkrankungen Zootiere, 28: 63–72. Kohm, V.A., 1986, Hauterkrankungen bei Eisbaren (Thalarctos maritimus) und ihre Behandlung mit Ivomec, Erkrankungen Zootiere, 28: 73–76. Koski, M., and Vandenbroek, D.J., 1986, Plesiomonas shigelloides-associated gastroenteritis in seals (Phoca vitulina richardsi), in Proceedings of the International Association for Aquatic Animal Medicine, 17: 125. Kuntze, V.A., 1986, Alterskrankheiten in einer Zirkus Eisbarengruppe, Erkrankungen Zootiere, 29: 265–270. Linnehan, R.M., Ulrich, R.W., and Ridgway, S., 1999, Enrofloxacin serum bioactivity in bottlenose dolphins, Tursiops truncatus, following oral administration of 5 mg/kg in whole fish, J. Vet. Pharmacol. Ther., 22: 170–173. Lyons, E.T., Keyes, M.C., and Conlogue, J., 1978, Activities of dichlorvos or disophenol against the hookworm (Uncinaria lucasi) and sucking lice of northern fur seal pups (Callorhinus ursinus) on St. Paul Island, Alaska, J. Wildl. Dis., 14: 455. Lyons, E.T., Kim, K.C., and Keyes, M.C., 1980, Variable activity of disophenol against hookworms and lice of northern fur seal pups on St. Paul Island, Alaska, J. Wildl. Dis., 16: 53–57. MacDonald, M.K., and Dougherty, S., 1983, Clinical pharmacokinetics in pinnipeds, in Proceedings of the International Association for Aquatic Animal Medicine, 14: 11. Manire, C., and Rhinehart, H.L., 2000, Use of human recombinant erythropoietin for the treatment of non-regenerative anemia in a rough-toothed dolphin (Steno bredanensis), J. Zoo Wildl. Med., 31: 157–163. Marcus, A.J., 1982, Hemorrhagic disorders: Abnormalities of platelet and vascular function, in Cecil Textbook of Medicine, Wyngaarden, J.B., and Smith, L.H. (Eds.), W.B. Saunders, Philadelphia, 979–992. Mazzaro, L.M., Dunn, J.L., Furr, H.C., and Clark, R.M., 1995a, Study of vitamin A supplementation in captive northern fur seals (Callorhinus ursinus) and its effect on serum vitamin E, Mar. Mammal Sci., 11: 545–553.
0839_frame_C31 Page 725 Tuesday, May 22, 2001 1:37 PM
Pharmaceuticals and Formularies
725
Mazzaro, L.M., Dunn, J.L., Furr, H.C., and Clark, R.M., 1995b, Vitamin A plasma kinetics in northern fur seals (Callorhinus ursinus), using vitamin A2 as a tracer, Can. J. Zool., 73: 10–14. Mazzaro, L.M., Helmick, K.E., Dunn, J.L., and Furr, H.C., 1995c, Vitamin E kinetics in harbor seals (Phoca vitulina), in Proceedings of the International Association for Aquatic Animal Medicine, 26: 118. McBain, J.F., 1984, Sulfamethoxazole toxicity in three killer whales, in Proceedings of the International Association for Aquatic Animal Medicine, 15: 38. McBain, J.F., 1985, Oral tetracycline absorption and excretion rates in killer whales (Orcinus orca), in Proceedings of the International Association for Aquatic Animal Medicine, 16: 79. Nakeeb, S., Babus, B., and Viele, D., 1980, Ketoconazole treatment for candidiasis, in Proceedings of the International Association for Aquatic Animal Medicine, 11: 109. Needham, D.J., 1978, Diseases of marine mammals, in Proceedings 36, Course for Veterinarians, University of Sydney, Australia, 36. Needham, D.J., 1993, Cetacean strandings, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 415–425. Newman, M.S., Cook, R.W., Appelhof, W.K., and Kitchen, H., 1975, Dermatophilosis in two polar bears, J. Am. Vet. Med. Assoc., 167: 561–564. Phillips, P.H., Davenport, P.K., and Schultz, D.J., 1986, Microsporum gypseum dermatophytosis in captive Australian sea lions, Neophoca cinerea, J. Zoo Anim. Med., 17: 136–138. Poelma, F.G., de Vries, G.A., Blythe-Russel, E.A., and Luykx, H.F., 1974, Lobomycosis in an Atlantic bottlenose dolphin in the dolphinarium Harderwijk, Aquat. Mammals, 2: 11. Reidarson, T.H., and McBain, J.F., 1994, The use of LDH isoenzymes to differentiate three medical conditions in cetaceans, in Proceedings of the International Association for Aquatic Animal Medicine, 25: 156. Reidarson, T.H., and McBain, J.F., 1995, The combined use of itraconazole and flucytosine in the treatment of chronic candida cystitis in a bottlenose dolphin (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 26: 13. Reidarson, T.H., and McBain, J.F., 1999, Hematologic, biochemical and endocrine effects of dexamethasone on bottlenose dolphins (Tursiops truncatus), J. Zoo Wildl. Med., 30: 310–312. Reidarson, T.H., Harrell, J.H., Rinaldi, M.G., and McBain, J.F., 1998, Bronchoscopic and serologic diagnosis of Aspergillus fumigatus pulmonary infection in a bottlenose dolphin (Tursiops truncatus), J. Zoo Wildl. Med., 29: 451–455. Reidarson, T.H., McBain, J.F., Dalton, L.M., and Rinaldi, M.G., 1999, Diagnosis and treatment of fungal infections in marine mammals, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 478–485. Ridgway, S.H, Green, R.F., and Sweeney, J.C., 1975, Mandibular anesthesia and tooth extraction in the bottlenose dolphin, J. Wildl. Dis., 11, 415–418. Riviere, J.E., 1999, Comparative Pharmacokinetics, Iowa State University Press, Ames, 307. Robeck, T.R., Dalton, L.M., and Young, W.G., 1996, Single daily dosing of amikacin in two beluga whales and an Asian small clawed otter, in Proceedings of the International Association for Aquatic Animal Medicine, 27: 19–21. Schroeder, J.P., 1983a, Techniques for improving reproductive efficiency in Tursiops, in Proceedings of the International Association for Aquatic Animal Medicine, 14: 10. Schroeder, J.P., 1983b, Apparent toxicity of ketoconazole for Tursiops truncatus during treatment of lobomycosis, in Proceedings of the International Association for Aquatic Animal Medicine, 14: 3. Schroeder, J.P., 1990, Reproductive aspects of marine mammals, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 353–369. Schroeder, J.P., 1993, Captive marine mammal reproduction, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 428–432. Schroeder, J.P., Cates, M.B., and Wallace, J.B., 1984, A Vibrio alginolyticus tissue infection in Tursiops truncatus, in Proceedings of the International Association for Aquatic Animal Medicine, 15.
0839_frame_C31 Page 726 Tuesday, May 22, 2001 1:37 PM
726
CRC Handbook of Marine Mammal Medicine
Smith, C.F., 1973, Dermatitis caused by Dermatophilus congolensis in polar bears (Thalarctos maritimus), Vet. Rec., 92: 533. Stetter, M., Mangold, B., Miller, M., Weber, M., and Capobianco, J., 1999, Successful treatment of lead toxicosis in a bottlenose dolphin (Tursiops truncatus), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 146–147. Stoskopf, M.K., 1990, Marine mammal pharmacology, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 139–162. Stoskopf, M.K., Moench, T., Thoen, C., and Charache, P., 1987, Tuberculosis in pinnipeds, in Proceedings 1st International Conference on Zoological and Avian Medicine, Honolulu, HI, 393. Sweeney, J.C., 1974a, Common diseases of pinnipeds, J. Am. Vet. Med. Assoc., 165: 805. Sweeney, J.C., 1974b, Procedures for clinical management of pinnipeds, J. Am. Vet. Med. Assoc., 165: 811. Sweeney, J.C., 1977, Intratracheal injection of antibiotics in the California sea lion, Zalophus californianus, and bottlenose dolphin, Tursiops truncatus, J. Wildl. Dis., 13: 49. Sweeney, J.C., 1985, Serum uptake and excretion rates for orally administered ampicillin and amoxicillin in large killer whales (Orcinus orca), in Proceedings of the International Association for Aquatic Animal Medicine, 16: 11. Sweeney, J.C., 1986a, Infectious diseases, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 777. Sweeney, J.C., 1986b, Clinical consideration of parasitic and noninfectious diseases, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 785. Sweeney, J.C., 1990, Surgery, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 215–233. Thurman, G.D., and Windsor, I.M., 1984, Serological testing for viral and mycoplasma antibodies in a captive dolphin as an aid in diagnosis and epidemiological control, in Proceedings of the International Association for Aquatic Animal Medicine, 15: 75. Townsend, F.I., 1999, Medical management of stranded small cetaceans, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 485–493. Townsend, F.I., and Petro, S., 1998, Significant panhypoproteinemia and regenerative anemia secondary to duodenitis in roughed-tooth dolphins (Steno bredanensis), in Proceedings of the International Association for Aquatic Animal Medicine, 29: 157. Townsend, F.I., Materese, F.J., and Sips, D.G., 1996, The use of liposomal amphotericin-B in the therapy of systemic zygomycosis, in Proceedings of the International Association for Aquatic Animal Medicine, 27: 18. Tyczkowska, K.L, Seay, S.S., Stoskopf, M.K., and Aucoin, D.P., 1992, Determination of ceftazidime in dolphin serum by liquid chromatography with ultraviolet-visible detection and confirmation by thermospray liquid chromatography-mass spectrometry, J. Chromatogr., 576: 305–313. Vandenbroek, D.J., Dierauf, L.A., Roletto, J., Koski, M.A., Amaya, L.M., and Gage, L.J., 1985, An epizootic of Leptospira (pomona) in California sea lions (Zalophus californianus), in Proceedings of the International Association for Aquatic Animal Medicine, 16: 122. Walsh, M.T., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., Murphy, D., and Innis, S.M., 1999, Pneumatosis intestinalis in orphan manatees (Trichechus manatus), diagnosis, pathological findings and potential therapy, in Proceedings of the International Association for Aquatic Animal Medicine, 30: 1. Williams, T.D., 1990, Cited in Stoskopf, M.K., 1990, Marine mammal pharmacology, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 139–162. Williams, T.D., 1993, Rehabilitation of sea otters, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 432–435. Williams, T.D., and Siniff, D.B., 1983, Surgical implantation of radiotelemetry devices in the sea otter (Enhydra lutra), Oral presentation, in Proceedings of the International Association for Aquatic Animal Medicine, 14: 31.
0839_frame_C31 Page 727 Tuesday, May 22, 2001 1:37 PM
Pharmaceuticals and Formularies
727
Williams, T.M., Davis, R.W., McBain, J.F., Tuomi, P., Wilson, R.K., McCormack, C.R., and Donoghue, S., 1995a, Diagnosing and treating common clinical disorders of oiled sea otters, in Emergency Care and Rehabilitation of Oiled Sea Otters, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 59–95. Williams, T.M., McBain, J.F., Tuomi, P., and Wilson, R.K., 1995b, Initial clinical evaluation, emergency treatments and assessment of oil exposure, in Emergency Care and Rehabilitation of Oiled Sea Otters, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 45–58. Yoshida, K., Yabe, K., Nishida, X., Yamamoto, N., Ohshime, C., Sekiguchi, M., Yamada, K., and Furuhama, K., 1998, Pharmacokinetic disposition and arthropathic potential of oral ofloxacin in dogs, J. Vet. Pharmacol. Ther., 21: 128–132. Young, S.J.F., and Huff, D.G., 1996, Fertility management in a female killer whale (Orcinus orca) with Altrenogest (Regu-mate ), in Proceedings of the International Association for Aquatic Animal Medicine, 27: 66. Young, S.J.F., Huff, D.G., and Anthony, J.M.G., 1999, A safe and cost effective oral care regime to control gingivitis and periodontal disease in the northern sea otter (Enhydra lutris lutris), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 67.
0839_frame_C31 Page 728 Tuesday, May 22, 2001 1:37 PM
0839_frame_C32 Page 729 Tuesday, May 22, 2001 1:38 PM
32 Euthanasia Leah L. Greer, Janet Whaley, and Teri K. Rowles
Introduction Veterinarians, biologists, and stranding network personnel are often faced with the need to euthanatize marine mammals to end an animal’s suffering. There are many considerations that must be addressed of when and how to euthanatize a marine mammal. When is an animal truly suffering? What method of euthanasia is most humane? How does one implement a method of euthanasia that requires either an accessible vein or specialized equipment? This chapter provides information on methods of euthanasia in all taxa of marine mammals so that informed decisions on techniques can be made. Also included is information on carcass disposal and tissue residue levels in chemically euthanatized marine mammals.
Stranded Animals A cetacean is considered stranded when it is found dead or live on land or is found in shallow water and unable to return to deeper water or in need of medical attention. Other marine mammals that spend normal periods of their lives on land (pinnipeds, walruses, sea otters, polar bears) are considered stranded when found dead or live, hauled out onshore, and unable to return to the water or in need of medical attention (Marine Mammal Protection Act, 1997). Many marine mammals are considered protected species around the world. When a stranded marine mammal is found alive, the responsible government agency (e.g., National Marine Fisheries Service for U.S. cetaceans and pinnipeds; Department of Conservation in New Zealand) or authorized stranding network personnel should be notified. The complex decision of whether the animal should be rehabilitated or euthanatized rests with the governing agency and its designated representatives, stranding network personnel. All stranded marine mammals must be given a physical examination to guide the initial assessment. Examination findings that may indicate euthanasia are the following: serious disabling locomotor injuries such as vertebral fractures; wounds that involve a large percentage of surface area or that have full penetration into the thoracic or abdominal body cavity; significant hemorrhage from the anus, genital opening, blowhole, or mouth; loss of reflexes from the anus, genital opening, blowhole, or tongue; marked prolonged hypothermia or hyperthermia, core body temperatures <35°C or >40°C (<95°F or >104°F), respectively; and extended length of time (over 12 hours) beached (Geraci and Lounsbury, 1993; Needham, 1993). Other factors that may contribute to the euthanasia decision include the location of the stranding and the logistics of humanely transporting the animal to a rehabilitation center or back into deep water.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
729
0839_frame_C32 Page 730 Tuesday, May 22, 2001 1:38 PM
730
CRC Handbook of Marine Mammal Medicine
A thorough record of the animal’s condition before euthanasia is essential adjunct information to necropsy reports. Each stranding should be respected as a unique event, and complete biological, medical, and environmental data should be obtained. Since many strandings are of public and media interest, thorough and careful communication of the animal’s condition and reason for euthanasia should be made to the stranding volunteers and public. This communication is especially important when the animal has been in rehabilitation for an extended period of time and the volunteers and the public have developed a close relationship with the animal.
Display and Collection Animals Marine mammals kept in captive environments, such as zoos and aquariums, often have greater access to veterinary care during their lifetime. Because these animals are usually intensively managed, an intimate relationship often develops between the animal, its caregivers, and perhaps visitors to the facility. The decision to euthanatize a charismatic marine mammal, particularly one in a display facility, is undoubtedly subject to public scrutiny. It is recommended that an open and positive relationship be established with everyone involved, including the media, at the onset of an illness. Thorough communication from the veterinarian explaining the extent of an illness, the differential diagnosis, and the perceived quality of life for the animal is usually well received. Such efforts should preclude the development of negative feelings that might arise when a popular animal is euthanatized.
Methods of Euthanasia Euthanasia may be achieved by one of three basic physiological mechanisms: (1) depression of neurons vital for life (e.g., typically by overdose of chemical anesthetics); (2) hypoxia, by either direct physical means (e.g., decapitation) or indirect means (e.g., paralytics); and (3) physical disruption of brain activity and destruction of neurons vital for life (e.g., captive bolt). There are many methods available to accomplish these results in mammals; however, marine mammals present unique circumstances. The available methods can be broadly classified as chemical (inhalant or injectable agents) or physical means. Although many methods will accomplish death, only a few are considered acceptable by published guidelines (Andrews et al., 1993; Close et al., 1996). In the 1993 Report of the American Veterinary Medical Association (AVMA) Panel of Euthanasia (Andrews et al., 1993), a panel of experts evaluated methods of euthanasia to determine humaneness and acceptability. The panel used the following criteria: 1. Ability to induce loss of consciousness and death without causing pain, distress, anxiety, or apprehension; 2. Time required to induce unconsciousness; 3. Reliability; 4. Safety to personnel; 5. Irreversibility; 6. Compatibility with requirement and purpose; 7. Emotional effect on observers or operators; 8. Compatibility with subsequent evaluation, examination, or use of tissue; 9. Drug availability and human abuse potential; 10. Age and species limitations; and 11. Ability to maintain equipment in proper working order (Andrews et al., 1993).
0839_frame_C32 Page 731 Tuesday, May 22, 2001 1:38 PM
Euthanasia
731
Although the guidelines were developed primarily with domestic animals in mind, all of these criteria should be considered when electing euthanasia of a marine mammal. A humane death is described as one that obtains rapid unconsciousness followed by cardiac or respiratory arrest (Andrews et al., 1993). Methods that do not create unconsciousness (e.g., paralytics, KCl, MgCl, hypothermia, cyanide, strychnine) are not considered humane to use alone for euthanasia in a conscious animal. Methods classified as acceptable by the AVMA panel are those that are considered humane when used alone in a conscious animal. However, if an animal is properly sedated to a level of unconsciousness, any method of euthanasia is considered humane. Based on the following discussion of the various methods available for euthanasia, the veterinarian and animal care personnel can make an informed decision on the most appropriate method to euthanatize an animal humanely in any given situation.
Injectable Agents Route of Administration Intravenous administration of an acceptable pharmaceutical agent is considered the most rapid and reliable means of obtaining humane euthanasia in mammals (Andrews et al., 1993; Close et al., 1996) and is the most common method used in marine mammals. Peripheral veins can be found in anatomical grooves of cetaceans (see Chapter 19, Clinical Pathology). The vessels lie under the dermis and can be accessed with superficial techniques, particularly in the tail fluke. When the vasculature starts to collapse in dying cetaceans, the ventral peduncle may be the most useful site for injection (see Chapter 19, Clinical Pathology). For small cetaceans a 1-in., 20-gauge needle is suitable, for larger cetaceans use a 1.5-in. needle, and for larger whales a needle of 2 in. or longer is needed (Sweeney, 1989). To access deeper vessels, a 6-in. needle can be used for an Orca-sized whale, and a 12- to 18-in. needle for a larger whale (RSPCA, 1997). Sites for venipuncture in different marine mammal groups are given in Chapter 19, Clinical Pathology. The disadvantages of the intravenous route are the difficulty in locating peripheral vasculature in debilitated or traumatized animals or animals in various stages of shock, and the potential danger to humans in restraining animals for access to vessels. If an injection cannot be administered intravenously, then less preferred routes can be used. Intraperitoneal administration is considered acceptable by AVMA standards. However, there are several problems that can be encountered, such as irritation of surrounding tissues and effects due to differential absorption leading to prolonged onset of action and variation in the effective dose (Smith et al., 1986). The thickness of the skin, blubber, and muscle must be taken into account when selecting needle length for an intraperitoneal route of administration, and access may be difficult in large mysticete whales. Intraperitoneal injection may be more appropriate for smaller animals; however, the human risks associated with restraint for injection remain. Intrahepatic administration of euthanasia solutions has been considered acceptable in cats (Grier and Schaffer, 1990; Andrews et al., 1993), and, after further study, may be shown to be a successful route of administration in some marine mammals. When compared with the intraperitoneal route, intrahepatic administration of sodium pentobarbital in cats resulted in minimal response to injection, moderate accuracy, low rate of excitability, and a significantly faster response, followed by cardiac standstill (Grier and Schaffer, 1990). Both intraperitoneal and intrahepatic administration may be less acceptable to the public or volunteers than intravenous dosing. Subcutaneous, intrathoracic, intrapulmonary, intrarenal, intrasplenic, and intrathecal routes irritate tissues, have a prolonged onset of action, and are considered unacceptable for administering
0839_frame_C32 Page 732 Tuesday, May 22, 2001 1:38 PM
732
CRC Handbook of Marine Mammal Medicine
euthanasia agents (Andrews et al., 1993; Close et al., 1996). Because the intracardiac route may be considered painful and unpredictable in a conscious animal, it is recommended only for anesthetized, moribund, or unconscious animals (Andrews et al., 1993; Close et al., 1997). Handlers may risk serious personal injury when restraining large marine mammals during administration of any injection, especially in uncontrollable environments (such as in the surf with polar bears, adult pinnipeds, and large whales). Therefore, sedating the animal prior to euthanasia will decrease the risk during handling. Sedation can be accomplished utilizing remote darting systems or intramuscular injections (see Chapter 29, Anesthesia, for intramuscular doses in marine mammals). Preliminary studies in three gray whales (Eschrichtius robustus) found that premedication with 0.02 to 0.03 mg/estimated kg body weight, or 15 mg/m of body length, of midazolam administered intramuscularly adequately sedated animals for intravenous euthanasia (Gulland, pers. comm.). Xylazine has been used in a gray whale, but the animal became excited and fractious; therefore, xylazine is not recommended for premedication in mysticetes (Gulland, pers. comm.).
Barbiturates Barbiturates are the most widely accepted mammalian euthanasia agents because of their rapid and targeted action (Andrews et al., 1993; Close et al., 1997). These drugs act by depressing the medullary respiratory and vasomotor centers to a degree that results in unconsciousness and respiratory and cardiac arrest. The onset of these reactions is quick, thus minimizing the discomfort to the animal. Some countries limit use of these drugs to people who are appropriately licensed (e.g., in the United States, individuals registered with the U.S. Drug Enforcement Agency), thereby limiting access to this drug in some stranding situations. The dosage for pentobarbitol-induced euthanasia for most species is 60 to 200 mg/kg. Immature gray whales 4 to 6 m in length have been successfully euthanatized with 180 to 230 ml of pentobarbital solution (390 mg/ml concentration) administered intravenously (Haulena and Gulland, pers. comm.). Pilot whales (Globicephala spp.) 4 to 6 m in length were successfully euthanatized with 120 ml of pentobarbital solution (390 mg/ml concentration) administered intravenously (Rowell, 1985). A dose of 10 mg/kg effectively induces deep anesthesia in cetaceans. This dose can induce apnea for a period long enough to cause hypoxia without the animal regaining consciousness (Sweeney, 1989), and would be considered humane euthanasia in circumstances where larger volumes (60 to 200 mg/kg) cannot be administered. The volume of pentobarbital can be reduced by premedication with acepromazine at 100 mg/m of body length (Needham, 1993) or midazolam at 15 mg/m of body length (Gulland, pers. comm.; Greer and Rowles, 2000). Many barbiturates other than pentobarbital have an acidic pH, and therefore are irritating if injected intraperitoneally.
Etorphine Etorphine has been used as an intramuscular alternative to intravenous euthanasia. Large animal Immobilon (etorphine and acepromazine) is not available in the United States, but can be obtained in many countries (Vericore, Novartis Animal Health UK Ltd, Litlington, Herts, U.K.). M-99 (etorphine HCl) (M99, Kruger-Med Pharmaceuticals, Johannesburg, South Africa) is the only form of etorphine that can be obtained in the United States. This drug is strictly regulated in several countries, including the United States, and few people are approved by the U.S. Drug Enforcement Agency to obtain and use this drug in the United States. Etorphine is an ultrapotent opioid and is considered up to 10,000 times more potent than morphine sulfate (Swan, 1993).
0839_frame_C32 Page 733 Tuesday, May 22, 2001 1:38 PM
Euthanasia
733
The potency of etorphine poses risk to personnel handling the drug, especially in the large doses needed for euthanasia (Morkel, 1993). Personnel can absorb etorphine through broken skin and mucous membranes (mouth, eyes, and nose). Etorphine should never be used unless a second person trained in handling opioid accidents and emergencies and a first-aid kit are present. Another hazard associated with etorphine is the potential for secondary toxicity if an animal or human ingests tissues from carcasses that have been euthanatized with etorphine. Following euthanasia with etorphine, the carcass must be properly disposed of to prevent any risk of tissue ingestion by animals or people. The dose of Immobilon used for euthanasia is approximately 0.5 ml/1.5 m in dolphins, and 4 ml/1.5 m in whales (Greenwood and Taylor, 1980; RSPCA, 1997; Barnett et al., 1999). Etorphine (M-99) is manufactured as 1 or 10 mg/ml solutions and is available only from Wildlife Pharmaceuticals in the United States. The dose of etorphine for immobilization can range from 0.5 to 5 µg/kg, but euthanasia dosages for most marine mammals have not been determined.
T-61 T-61 is an injectable mixture of a local anesthetic, a hypnotic agent, and curariform drug— N-2-(m-methoxyphenyl)-2-ethylbutyl-l-γ-hydroxybutyramide (20%), 4,4′-methylene biscyclohexyltri-methyl ammonium iodide (5%), and tetracaine HCl (0.5%) in aqueous solution with formamide. This drug should only be used intravenously because there are concerns about differential absorption when administered by any other route. There has been concern that the curariform component may take effect before the onset of unconsciousness, causing distress to the animal. Studies in dogs and rabbits have shown that a loss of consciousness occurs simultaneously with paralysis (Hellebrekers et al., 1990), making this agent acceptable in these species. This drug is no longer available in the United States, but it is not controlled in many countries and therefore may be available. The dose extrapolated from small animals is 0.3 ml/kg intravenously (Hyman, 1990).
Paralytics Paralytic agents have been reportedly used in stranded marine mammals, primarily because these drugs are not all controlled substances. The mechanism of action involves muscle paralysis, respiratory restriction, and hypoxia induction. Animals that have received paralytics as euthanasia agents suffocate while maintaining consciousness. This process can be slow and prolonged in diving species that can withstand long periods of apnea. The addition of potassium chloride with succinyl choline has been used to induce cardiac arrest (i.e., the potassium ion is cardiotoxic) and thereby shorten the period that paralytics may take to induce death (Hyman, 1990). There is no advantage to using these drugs over other chemical agents. An intravascular or intraperitoneal route is still required, and animals often exhibit muscular rigidity, spasms, vocalizations, and gasping while still maintaining consciousness. Paralytics, and potassium and magnesium salts, are explicitly listed as inhumane methods of euthanasia (Andrews et al. 1993; Close et al., 1997) due to the high expectation of fear, struggling, or pain before death occurs. The International Whaling Commission report of the workshop on humane killing techniques for whales (1980) concluded that paralytic agents should not be used for killing whales since they do not produce loss of consciousness. Accidental injection of personnel handling paralytic agents is considered a life-threatening event. Paralytic agents should never be used unless a person trained in treatment of paralytic drug accidents and an appropriate first-aid kit are present.
0839_frame_C32 Page 734 Tuesday, May 22, 2001 1:38 PM
734
CRC Handbook of Marine Mammal Medicine
Inhalants Inhalant anesthetic agents such as halothane, isoflurane, methoxyflurane, and enflurane are considered humane methods of euthanasia. These agents are more easily applied for euthanasia of captive animals, smaller animals (sea otters, pinnipeds), or animals already anesthetized. The disadvantage of these agents are the expensive delivery systems required for administration, the extended period of time required in breath-holding species, and exposure hazards to personnel. Examples of personnel risks from some inhalant agents are explosions (ether), narcosis (halothane), hypoxemia (nitrogen, carbon dioxide), spontaneous abortion (halothane), and addiction (nitrous oxide). Carbon dioxide, delivered in a closed chamber, is commonly used for euthanasia of laboratory animals. Carbon dioxide concentrations must reach levels high enough to induce unconsciousness and subsequent death. It is doubtful that these levels would be reached quickly in breath-holding animals or species with adaptations to high concentrations of carbon dioxide. Carbon dioxide has also been shown to cause an excitatory phase before death in cats and dogs, is not recommended in species larger than a cat, and may not be as painless as previously believed (National Research Council, 1992; Andrews et al., 1993; Close et al., 1997).
Physical Methods Several physical methods of euthanasia have been employed in marine mammals. For a physical method of destruction to be considered humane, it must fulfill the requirement of rapidly inducing relatively painless unconsciousness before death. Only methods that quickly and relatively painlessly destroy the brain or brain stem are considered humane methods of euthanasia. All other physical methods of euthanasia (e.g., exsanguination, suffocation, bilateral thoracotomy, gunshot to heart) are only considered humane if used in a heavily sedated, unconscious, or moribund animal or as a secondary confirmation of euthanasia. There may be adverse public reaction to the use of some physical methods of euthanasia.
Ballistics A scientific approach to the use of ballistics has not been reported in polar bears (Ursus maritimus), pinnipeds, walruses (Odobenus rosmarus), or sea otters (Enhydra lutris); however, the anatomy of the target organs for euthanasia of these species is not significantly different from terrestrial mammals. If done correctly, ballistics should cause instantaneous unconsciousness (e.g., instant destruction of the brain) followed by respiratory and cardiac arrest. When ballistics are used in field conditions, the caliber of the firearm should be appropriate for the species. Personnel should be trained to hit specific target organs (e.g., neck, heart) in field conditions. Since, in some cases, ballistics do not cause unconsciousness before death, this method is considered killing or harvesting rather than euthanasia (Andrews et al., 1993). Ballistics have been evaluated for euthanasia of cetaceans. Several anatomical parameters in cetaceans make the use of ballistics challenging. The skin, blubber, and muscle of the forehead (melon) are arranged such that kinetic energy from a projectile is absorbed, dampening the impact. The anterior (frontal) surface of the cetacean skull is concave with extensive sinuses, increasing the likelihood of bullet deflection (Barzdo and Vodden, 1983). The extensive muscle on the nuchal, parietal, and occipital regions of the skull makes occipital shooting ineffective. Use of ballistics in mass strandings can be distressful to the surviving animals and to the personnel responding to the event. Exposure to the noise, visual destruction, agonal vocalizations,
0839_frame_C32 Page 735 Tuesday, May 22, 2001 1:38 PM
Euthanasia
735
and possible release of pheromones by frightened animals may exacerbate anxiety and fear in the conscious animals (National Research Council, 1992). In smaller cetacean species, there are two different documented approaches. The first is a shot aimed through the blowhole at a 45° angle directly down and back (ventroposteriorly) to an area behind the pectoral flippers. The second is a horizontal shot lateral to the brain, just above (dorsal) the center of the eye–ear line (RSPCA, 1997). The preferred shot is the lateral approach, particularly in larger-sized cetaceans since the skull is closest to this body surface. The ventroposterior shot can be used in bottlenose dolphins (Tursiops truncatus) shooting preferably when the blowhole is open, but not placing the muzzle of the firearm in the blowhole (pressure can cause firearm explosion). The firing range should be 0.4 to 1.0 m from the head (Geraci and Lounsbury, 1993; Blackmore et al., 1995; RSPCA, 1997). Bullets shot at point-blank range are subjected to greater yaw when penetrating the thick soft tissue that surrounds a cetacean brain, and this method is therefore not recommended (RSPCA, 1997). Shooting cetaceans through the thorax will likely result in persistent consciousness and a slow death (Geraci and Lounsbury, 1993). There are conflicting reports on the type of firearm to use in these smaller cetaceans. The RSPCA Stranded Cetaceans, Guidelines for Veterinary Surgeons, 1997, states that a shotgun or a .22-caliber rifle should never be used. A shotgun with buckshot or a .22-caliber projectile may not reliably penetrate the skull. Bullets that are solid or jacketed, a minimum of .30 caliber, and a minimum of 140 grains (9.8 g) are recommended (Geraci and Lounsbury, 1993; RSPCA, 1997). Hollow or soft bullets do not reliably penetrate the skull. Use of high-powered rifles also poses a risk to humans on rocky beaches, where richochets of penetrating bullets may occur. A safe and humane shot with a high-powered rifle requires someone who is trained and adequately skilled to destroy the brain accurately and rapidly kill the animal. To address the concerns of using high-powered rifles, a study was performed on carcasses of a common dolphin (Delphinus delphis) and five pilot whales (≤ 5.7 m) using a shotgun with 28-g lead slug or 28-g buckshot. The authors concluded that shooting smaller cetaceans (less than 4 m) from the dorsal ventral approach with buckshot was effective (Blackmore et al., 1995, their Fig. 7a). In larger cetaceans (4 to 6 m), this approach did not penetrate the skull, although buckshot from the lateral approach of the brain appeared to be effective (Blackmore et al., 1995). The use of normally available ballistics in larger whales is challenging and not recommended in whales larger than 9 m, sperm whales (Physeter macrocephalus) of any size, or baleen whales other than minke whales (Balaenoptera acutorostrata), due to the anatomy of the head and blowhole (Barzdo and Vodden, 1983; Geraci and Lounsbury, 1993; Needham, 1993). The brain is deeply buried in these larger cetaceans. For any projectile to be effective it would need to penetrate approximately 1.2 m of blubber, muscle, and bone, and still maintain enough kinetic energy to destroy the brain and cause immediate unconsciousness and death. The Department of Conservation in New Zealand (Marsh and Bramber, 1999) have reported the development of a specialized round and firearm for the humane euthanasia of sperm whales. They describe a specially designed 14.5 × 114 mm anti-aircraft, 61 g, 12 L14 leadloy boreriding bullet with a flat tip. A firearm was also extensively modified to use this round effectively. The result was an 11.8-kg firearm that had a 2.4-m recoil, which must be operated standing sideways. Operators require training and practice to prevent serious injury to themselves. In field studies, two sperm whales were euthanatized. One whale died after a single shot, and the second was rendered insensible by the first shot. In the second instance, a second shot gave the appearances of a dead whale, but the animal resumed breathing for another 2.5 hours. Further studies are being done to determine accurate target areas in the sperm whale.
0839_frame_C32 Page 736 Tuesday, May 22, 2001 1:38 PM
736
CRC Handbook of Marine Mammal Medicine
When using ballistics for euthanatizing cetaceans, three main components must be evaluated: the size and anatomy of the animal, the firearm and projectile to be used, and the skill of the marksman. If any of these variables are less than ideal, then this method should not be used. In the RSPCA Stranded Cetaceans: Guidelines for Veterinary Surgeons (1997), the authors suggest that it may be more humane to leave the animal to die on its own rather than applying any substandard method of euthanasia, especially in larger whales like sperm or baleen whales. The gravitational weight on the internal organs will likely induce a more humane death than repeated rounds of projectiles fired inaccurately, but may take a prolonged time (RSPCA, 1997).
Explosives Explosives have been used in attempts to euthanatize larger whales that are difficult to euthanatize by other methods. These methods are usually considered less acceptable, because of the tremendous soft-tissue damage, excessive noise, and required expertise in the application of explosives (i.e., human safety factor). The noise and visual disturbance is especially unacceptable in mass strandings when other conscious animals, stranding network volunteers, and the public are present (see also Carcass Disposal, below). There are two different techniques for using explosives. A charge can be placed externally, caudal to the blowhole, and sand bagged to direct the implosion down toward the brain. This technique causes extensive tissue destruction and does not reliably cause instantaneous unconsciousness (RSPCA, 1992). Alternatively, a charge placed inside the mouth (by a pole) at the base of the brain has reportedly worked as well (Geraci and Lounsbury, 1993; Needham, 1993). The positioning inside the mouth must be accurate, since the impact of blast effects decreases rapidly with distance (RSPCA, 1992). Explosives can be dangerous and may be prohibited in public areas. The authors of the RSPCA (1992) and the Victorian whale rescue plan (1986) do not recommend this method for euthanasia of large whales.
Verification of Death It is imperative that death be verified. The absence of a heartbeat is the only reliable confirmation of death in mammals; however, with large marine mammals in field situations, it may not always be possible to detect a heartbeat. If there is any doubt about confirmation of death, a secondary physical means of euthanasia can be performed to ensure death (Close et al., 1996). Physical methods include bilateral thoracotomy, exsanguination, and gunshot through the heart or brain. In most cases, exsanguination is more acceptable to the public than other described methods.
Carcass Disposal A thorough necropsy can facilitate carcass disposal (smaller pieces) or complicate carcass disposal (decreased ease of towing). Carcass disposal is less of a problem with most pinnipeds, otters, and small cetaceans but becomes problematic with large whales. In most cases, smaller carcasses can be transported for rendering, burial, or incineration. For large carcasses, the options are limited. A carcass may be left alone, buried, towed to sea and sunk or released, moved, or rendered. Previous attempts to burn or blow up carcasses created more problems than solutions and are not recommended (Geraci and Lounsbury, 1993). A carcass left on a remote beach will decompose quickly, and those that are sunk at sea provide habitat and food for numerous marine species. However, this should not be done if there is any concern about high concentrations of euthanasia solution in the carcass.
0839_frame_C32 Page 737 Tuesday, May 22, 2001 1:38 PM
Euthanasia
737
Little information has been published to date on tissue residue levels of anesthetic drugs in marine mammal carcasses; however, leaving such a carcass has been presumed to be potentially harmful to scavenging animals. Preliminary results in three gray whales and one pilot whale euthanatized with pentobarbital at 20 to 40 mg/estimated kg of body weight (25 ml/m of body length) showed that the tissues residue levels ranged from 1.04 (muscle) to 0.001 (cerebrum) µg/g of tissue (Greer and Rowles, 2000). The highest tissue residue levels were found in muscle and liver. Based on the highest level in muscle, a dog would have to consume 85 kg of muscle/kg of body weight to reach a lethal ingested dose (Clarke et al., 1981). A carcass that is buried should be at a site approved by the local authorities or beach owners. The body cavity should be opened, then buried deep enough to prevent reexposure of the tissues and digging scavengers from finding the carcass. Towing and releasing at sea is problematic, since bloated carcasses tend to float and may re-beach themselves at a later date or become a navigation hazard. Of about ten gray whale carcasses towed out to sea in California in 2000, six returned to the beach (Cordaro, pers. comm.). Cetacean carcasses should be towed by the tail, have the body cavity opened, be hauled far enough offshore to prevent drifting back, and have enough ballast attached to sink it (Geraci and Lounsbury, 1993). However, there may be cases in which this cannot occur. Carcasses that return to the beach can be costly (i.e., second disposal costs) and may significantly alter stranding statistics. Therefore, any carcass that is towed out to sea should be marked in some manner, such as tail fluke or lateral thoracic notching, so that it can be recognized as a previously stranded animal. Alternatively, an animal can be moved to another site for further study or more appropriate disposal. Some carcasses may need to be cut into smaller pieces for adequate disposal. Rendering plants, commercial incinerators, and veterinary schools may accept marine mammal carcasses. These collaborations and connections for carcass disposal should be pursued before an actual event occurs. Commercial trade in marine mammal parts is prohibited under the Endangered Species Act and the Marine Mammal Protection Act; therefore, carcasses or parts of carcasses cannot be sold.
Acknowledgments The authors thank Frances Gulland, Marty Haulena, Bill McLellan, and stranding network volunteers of The Marine Mammal Center and University of North Carolina, Wilmington, for providing data on euthanasia doses and methods, and collecting tissue residue studies, and Dorcas O’Rourke and Ed Ramsay for their critical comments on this chapter.
References Andrews, E.J., Bennet, B.T., and Clark, J.D., 1993, Report of the AVMA panel on euthanasia, J. Am. Vet. Med. Assoc., 202: 230–247. Barnett, J.E.F., Jepson, P.D., and Patterson, I.A.P., 1999, Drug-induced euthanasia of stranded cetaceans, Vet. Rec., 145: 292. Barzdo, J., and Vodden, P., 1983, Report of Stranded Whale Workshop. A Practical and Humanitarian Approach, RSPCA, Horsham, U.K. Blackmore, D.K., Madie, P., Bowling, M.C. et al., 1995, The use of a shotgun for the euthanasia of stranded cetaceans, N.Z. Vet. J., 43: 158–159. Clarke, M.L., Harvey, D.G., and Humphrey, D.J. (Eds.), 1981, Barbiturates and other hypnotics, sedatives and tranquillizing drugs, in Veterinary Toxicology, 2nd Ed., Bailliere Tindall, Cassell Ltd, London, U.K., 106–107.
0839_frame_C32 Page 738 Tuesday, May 22, 2001 1:38 PM
738
CRC Handbook of Marine Mammal Medicine
Close, B., Banister, K., and Baumans, V., 1996, Recommendations for euthanasia of experimental animals, Lab. Anim., 30: 293–316. Close, B., Banister, K., and Baumans, V., 1997, Recommendations for euthanasia of experimental animals, Lab. Anim., 31: 1–32. Geraci, J.R., and Lounsbury, V.J., 1993, Marine Mammals Ashore. A Field Guide for Strandings, Texas A&M Sea Grant Publication, Galveston, 305 pp. Greenwood, A.G., and Taylor, D.C., 1980, Humane handling of stranded cetaceans, Vet. Rec., 106: 345. Greer, L., and Rowles, T., 2000, Humane euthanasia of stranded marine mammals, in Proceedings of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine, Joint Conference, New Orleans, LA, September 17–21, 374–375. Grier, R.L., and Schaffer, C.B., 1990, Evaluation of intraperitoneal and intrahepatic administration of a euthanasia agent in animal shelter cats, J. Am. Vet. Med. Assoc., 197: 1611–1615. Hellebrekers, L.J., Baumans, V., Bertens, A.P., and Hartman, W., 1990, On the use of T-61 for euthanasia of domestic and laboratory animals; an ethical evaluation, Lab. Anim., 24: 200–204. Hyman, J., 1990, Euthanasia in marine mammals, in CRC Handbook of Marine Mammal Medicine: Health, Disease, and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 265–266. Marsh, N., and Bramber, C., 1999, Development of a specialized round and firearm for the humane euthanasia of stranded sperm whales (Physeter macrocephalus) in New Zealand, Report to the 51st Meeting of the International Whaling Commission, IWC/51/WK5. Morkel, P., 1993, Prevention and management of capture drug accidents, in The Capture and Care Manual. Capture, Care, Accommodation and Transportation of Wild African Animals, McKenzie, A.A. (Ed.), Wildlife Decision Support Services and the South African Veterinary Foundation, Pretoria, South Africa, 100–113. National Research Council, 1992, Euthanasia, in Recognition and Alleviation of Pain and Distress in Laboratory Animals, National Academy Press, Washington, D.C., 102–116. Needham, D.J., 1993, Cetacean strandings, in Zoo and Wild Animal Medicine: Current Therapy 3, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 415–425. Rowell, S.F., 1985, Stranded whales, Vet. Rec., 116: 167. RSPCA (Royal Society for the Prevention of Cruelty to Animals), 1992, Stranded Whales, Dolphins and Porpoises. A First Aid Guide, RSPCA, Horsham, U.K., 18. RSPCA (Royal Society for the Prevention of Cruelty to Animals), 1997, Stranded Cetaceans: Guidelines for Veterinary Surgeons, RSPCA, Horsham, U.K., 14–16. Smith, A.W., Houpt, K.A., and Kitchell R.L., 1986, Report of AVMA panel on euthanasia, J. Am. Vet. Med. Assoc., 188: 252–268. Swan, G.E., 1993, Drugs used for the immobilization, capture, and translocation of wild animals, in The Capture and Care Manual: Capture, Care, Accommodation and Transportation of Wild African Animals, McKenzie, A.A. (Ed.), Wildlife Decision Support Services and the South African Veterinary Foundation, Pretoria, South Africa, 2–64. Sweeney, J.C., 1989, What practitioners should know about whale strandings, in Kirk’s Current Veterinary Therapy, 10th ed., Kirk, R. (Ed.), W.B. Saunders, Philadelphia, 721–727. Warneke, R.M., (Ed.), 1986, Victorian whale rescue plan. A contingency plan for strandings of cetaceans (whales, dolphins and porpoises) on the Victorian coastline, Fisheries and Wildlife Service, Department of Conservation, Forests and Lands, Melbourne,Victoria.
0839_frame_C33.fm Page 739 Tuesday, May 22, 2001 1:38 PM
VII Marine Mammal Well-Being
0839_frame_C33.fm Page 740 Tuesday, May 22, 2001 1:38 PM
0839_frame_C33.fm Page 741 Tuesday, May 22, 2001 1:38 PM
33 U.S. Federal Legislation Governing Marine Mammals Nina M. Young and Sara L. Shapiro
Federal Legislation and Regulations—Discussion Nina M. Young
Introduction Federal legislation and regulations touch the lives of every researcher and veterinarian in the marine mammal field at one time or another. Knowing and understanding these laws and regulations and how to navigate the federal permit process is critical, and can help facilitate the conduct of a researcher’s or clinician’s work. The U.S. legislation governing marine mammals, as well as the international treaties the legislation implements, can be confusing to those unfamiliar with them. For this reason, it is essential to consider the legal responsibilities of undertaking activities involving marine mammals. The legislation and regulation of human activities relating to animals have their origins in many ancient civilizations of the world. These laws and rules of conduct regarding human relationships to, and interactions with, animals were usually centered on certain species that held some particular cultural, religious, sporting, or nutritional significance to the society regulating such activities (Cooper, 1987). Since the 19th century, the idea that animals require some form of protection has led to an exponential growth in the interest in animal law. In the 20th century, particularly during the 1970s, the establishment of many local, national, and international legislative efforts resulted in the laws and regulations that exist in the United States today. These laws and regulations are due largely to the advocacy of conservation and animal welfare groups, motivated by the public concern for the environment in the late 1960s and early 1970s. During that period, the focus on commercial whaling sparked the public’s growing fascination with whales, dolphins, and other marine mammals. The simultaneous emphasis of “marine mammal species” and “the marine mammal as an individual” has helped mold the federal legislation that at present regulates marine mammals. This chapter deals specifically with federal legislation in the United States related to marine mammals, and is not intended to be a discussion or analysis of the legalistic basis upon which the legislation governing these marine mammals is based. Rather, it is to serve as an outline of the major laws and regulations in the United States at present affecting people’s interactions with marine 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
741
0839_frame_C33.fm Page 742 Tuesday, May 22, 2001 1:38 PM
742
CRC Handbook of Marine Mammal Medicine
mammals, and to give the reader some practical guidelines for dealing with this legislation. U.S. legislation related to marine mammals is still relatively new to the legal world, and as a consequence, is still evolving. Before relying on any portion of this chapter, reference should be made to the U.S. Code (USC) and the Code of Federal Regulations (CFR). Once a piece of legislation or amendments to a particular law become law, these new laws or amendments are published in the USC. In addition, the text of the law and the floor debate are published in the Congressional Record, which is the daily journal of Congress’s action on the floor of both the House of Representatives and the Senate. Laws such as the Marine Mammal Protection Act (MMPA) or the Endangered Species Act (ESA) are contained in the USC. Each law then needs certain implementing regulations, which are the Executive Branch agencies’ attempts to both interpret and implement the law. Regulations are developed through a process of public notice and comment rule-making, mandated under the Administrative Procedures Act. A notice and text of a proposed regulation is published in the Federal Register; this notice provides the public with the opportunity to review and comment on a particular regulation within a specified time period. Once the regulation becomes final, it is published in the Federal Register as a final rule, and then in the CFR. Finally, one can also contact the relevant governmental agency or agencies for the most recent information pertaining to the activity under consideration. One of the most useful Web sites for this information is http://thomas.loc.gov.
The Responsible Regulating Agencies Before discussing the various pieces of legislation that affect marine mammals, it is best to outline briefly the Executive Branch governmental agencies that administer these laws and regulations. The administration of the various federal laws in the United State that affect marine mammals is essentially divided among three federal agencies. Activities that involve what Congress terms domestic marine mammal species (those found in the United States) are administered by the U.S. Fish and Wildlife Service (FWS) of the U.S. Department of the Interior (DOI), and include walruses (Odobenus rosmarus), manatees ( Trichechus manatus), dugongs (Dugong dugon), sea otters (Enhydra lutris), and polar bears (Ursus maritimus). The National Marine Fisheries Service (NMFS) of the National Oceanic and Atmospheric Administration (NOAA) of the U.S. Department of Commerce (DOC) administers activities that involve the remaining marine mammal species, referred to by Congress as oceanic species, and including all other pinnipeds and the cetaceans. A word of caution: Do not be confused by the groupings of “domestic” and “oceanic.” There is no biological basis for the groupings, which merely represent an arbitrary decision by the U.S. Congress. Nor are the oceanic species (pinnipeds at least) wholly aquatic. Both the FWS and the NMFS operate under the authority granted them by the ESA and the MMPA. Both the FWS and the NMFS administer requests through their permitting divisions for “prohibitions against taking” (see the ESA discussion below) or “waiving the moratorium” (see the MMPA discussion below), including granting or denying requests for exemptions; issuing permits; carrying out research and management programs; enforcing provisions of the MMPA and the ESA; participating in international programs, agreements, and treaties; issuing rules and regulations to carry out their missions to conserve and protect marine mammals; cooperating with the states to decide whether to act on requests for the return of management authority to those states; and cooperating with conservation organizations, the public, other federal agencies, the Marine Mammal Commission (MMC) and its Committee of Scientific Advisors, and many constituent groups, particularly scientific researchers and the public display community. In addition, the FWS lists and delists species as endangered or threatened and undertakes other activities under the ESA. The NMFS also has responsibilities under the ESA to the species for which it is
0839_frame_C33.fm Page 743 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
743
responsible. By mutual agreement, the FWS and the NMFS are party to an agreement with the Animal and Plant Health Inspection Service (APHIS) at the U.S. Department of Agriculture (USDA) for administering the MMPA and the Animal Welfare Act (AWA). The APHIS administers activities that involve all aspects of care and maintenance for any captive marine mammal. The APHIS operates under the authority granted it by the AWA. In administering captive care and maintenance of marine mammals, the APHIS is responsible for the organization and implementation of the AWA. This includes, but is not limited to, the promulgation of captive care rules and regulations; licensing and registration of persons subject to the AWA (which includes dealers, carriers, exhibitors, research facilities, etc.); prelicensing inspections, compliance inspections, and animals-in-transit inspections; and complaint investigations of those licensed facilities under the AWA (Vehrs, 1985). The APHIS also interacts with the FWS, the NMFS, and the APHIS under the MMPA and the AWA (see Chapters 40 through 45). The MMC was established under Title II of the MMPA. The MMC “is charged with the responsibility for developing, reviewing, and making recommendations on actions and policies for all federal agencies with respect to marine mammal protection and conservation and for carrying out a research program” (16 USC 1402). The MMC is an independent agency of the Executive Branch, and consists of three part-time commissioners, who are appointed by the President. The MMC makes appointments to the Committee of Scientific Advisors on Marine Mammals (16 USC 1401). This body is largely responsible for producing the work that encompasses, but is not limited to, reports on marine mammal species of special concern, management problems, international aspects of marine mammals, research programs, and marine mammal/fisheries interactions. The MMC interacts with and comments on the activities of the FWS, the NMFS, and the APHIS regarding marine mammals (Marine Mammal Commission, 1999). Although the FWS, the NMFS, and the APHIS are not necessarily bound by recommendations of the MMC, they do tend to heed the recommendations the MMC forwards to them in the course of their administrative activities affecting marine mammals. These activities are compiled by the FWS, the NMFS, the APHIS, and the MMC in annual reports available to the general public upon request (generally about February of each year). Notice of such reports is usually made in the Federal Register. The interplay and interactions among the various governmental agencies involved with marine mammals are complex, and can be confusing and frustrating, particularly to those not familiar with how the system(s) works. This confusion can be further exacerbated by recognizing that two agencies can have authority over the same marine mammal species, under several laws at the same time (i.e., both the FWS and the NMFS share similar responsibilities for administration of the Hawaiian monk seal (Monachus schauinslandi), because it is a marine mammal whose habitat is on a National Wildlife Refuge, and it is a highly endangered species under the ESA). It is important to keep in mind that the laws discussed here define and regulate any human activities (rather than regulating the marine mammals themselves) within the United States that affect marine mammals or their environment. The laws and regulations outlined below, therefore, are always open to redefinition and reinterpretation, particularly regarding issuance of permits, and it is imperative for individuals who intend to become involved with marine mammals to update themselves regularly on the current status of the laws and their regulations.
The Endangered Species Act (16 USC 1531–1544; 50 CFR 1539 ff)* The ESA is the most comprehensive U.S. wildlife conservation statute, protecting threatened and endangered species from being killed or injured, prohibiting trade in such species, and *Available at http://www.fws.gov.
0839_frame_C33.fm Page 744 Wednesday, May 23, 2001 11:00 AM
744
CRC Handbook of Marine Mammal Medicine
protecting the habitat upon which these species depend. In addition, the ESA creates special obligations for federal agencies to use their authorities to conserve threatened and endangered species. The ESA was first passed in 1969, and was amended in 1973 to incorporate the United States as a party to the Endangered Species Convention (CITES, outlined below) (USFWS, 2000). Congress periodically reviews the ESA, amends it, and reauthorizes appropriations (funds) to implement the responsibilities under the law. The ESA was reauthorized in 1978, 1982, and 1988. It is has been up for reauthorization for nearly 10 years, but Congress has been deadlocked by controversies over the ESA and its impact on private property rights and public resource use and management. Meanwhile, all the substantive provisions of the ESA remain in effect, with Congress adopting annual appropriations for ESA implementation, while debate on the future of the law’s provisions continues. Listing, Critical Habitat, and Recovery Plans
For a species (which includes subspecies and distinct populations of vertebrate animals) to be protected under the ESA, it must first be listed pursuant to Section 4 of the ESA by the Secretary of the Interior, as either endangered (any species in danger of extinction throughout all or a portion of its range) or threatened (any species likely to become endangered in the foreseeable future). In the case of marine species, the Secretary of Commerce makes a listing determination and forwards it to the Secretary of the Interior, if a species is to be added to the list. Species may receive the designation of endangered or threatened, if the Secretary of the Interior, or the Secretary of Commerce, determines such by virtue of any of the following five factors: 1. 2. 3. 4. 5.
The present or threatened destruction, modification, or loss of use of the species habitat or range; The overexploitation of the species for commercial, recreational, scientific, or educational purposes; Predation or disease problems; Existing regulatory mechanisms found to be inadequate; or Other natural or anthropogenic factors affecting the continued existence of the species.
Individuals of such federally listed species are included whether they are living or dead, or whether they are parts of the animal (such as tissues, teeth, or bones) or the entire animal. Species are removed from the endangered or threatened designation by an improvement or alteration in their status, or by extinction, followed by a formal delisting action. Any interested person may petition to have a species considered for addition to or removal from the ESA, and must provide information with the petition that indicates the reasons the petitioned action is warranted. Any decision regarding a species under the ESA is to be made “solely on the basis of the best available scientific and commercial data” (16 USC 1533 (b)(1)(A)). When a species is listed, critical habitat (those areas essential to the survival and recovery of the species) is to be designated to the “maximum extent prudent and determinable.” More than 1600 species worldwide are listed as threatened or endangered under the ESA, including more than 1000 species in the United States, of which only four marine mammals currently have a critical habitat designated. Once species are listed, Recovery Plans (blueprints for bringing species back to the point where they no longer need to be listed as threatened or endangered) are supposed to be developed and implemented. Recovery Plans have been developed for about 75% the U.S. listed species, but including just one marine mammal. Protection for Listed Species
Section 9 of the ESA prohibits anyone (including private persons, corporations, state and federal agencies) from “taking” endangered species in the United States, or in territorial or high seas of the United States. “Take” is defined as killing, injuring, harassing, harming, or capturing an endangered species. Significant habitat modification, which disrupts essential breeding, feeding,
0839_frame_C33.fm Page 745 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
745
or sheltering activities of endangered species, is also prohibited. Under DOI regulations, threatened terrestrial species generally receive the same protection as endangered species. For a threatened marine species, however, the DOC must issue regulations for the particular species, specifying what protections apply to it. Furthermore, within the jurisdiction of any state, more restrictive state laws or regulations (if such exist) with regard to listed species prevail relative to “taking” and other actions involving threatened and endangered species. Permits
Under Section 10 of the ESA, permits are available that allow for the incidental take of threatened or endangered species, provided that a habitat conservation plan to minimize and mitigate the impact of the take is approved by the Secretary of the Interior or the Secretary of Commerce for those species, including marine mammals, under their jurisdiction and administration. In addition, “enhancement of survival” permits are available, allowing the take of endangered species for scientific research, endangered species propagation, to enhance the conservation and survival of a threatened or endangered species, and to provide for Alaskan native subsistence. Specifically, the Assistant Administrator (AA) of the NMFS, or the Director of the FWS, may issue permits for incidental take and scientific purposes, to enhance the propagation and/or survival of listed species. The ESA permit authorizes (under such terms and conditions as the AA or Director may prescribe) “taking,” importing, or conducting other actions with respect to listed species otherwise strictly prohibited by Section 9 of the ESA. Permits for threatened species may be issued for the same purposes as endangered species permits, with the addition of economic hardship, zoological exhibition, educational purposes, or special purposes consistent with the goals of the ESA. The Director of the FWS or the AA for the NMFS makes the final determination on each permit application. The type of information required for a permit application, instructions for completing the application, and the applicable federal regulations authorizing such permits can be found at http://www.nmfs.noaa.gov or http://www. fws.gov (see also Marine Mammal Permits: Frequently Asked Questions at the end of this chapter). Consultations
Section 7 of the ESA creates an additional duty on the part of all federal agencies to use their authorities to further the conservation of threatened and endangered species. Specifically, federal agencies must consult with the Secretary of the Interior (the FWS) or the Secretary of Commerce (the NMFS) to ensure that any action that any federal agency authorizes, funds, or carries out is not likely to jeopardize the continued existence of a listed species or adversely modify or destroy its critical habitat. As a result of Section 7 consultation, the FWS or the NMFS will provide the federal agency proposing an action with a biological opinion regarding the effects of the proposed action on listed species or critical habitat. The biological opinion may suggest reasonable and prudent measures (RPMs) and conservation recommendations to mitigate for the impacts of the proposed action on a species or critical habitat. If the proposed action is likely to result in jeopardy to a listed species or adverse modification or destruction of critical habitat, the biological opinion will require reasonable and prudent alternatives (RPAs) that will allow the action to go forward without jeopardizing the species or destroying critical habitat. If the agency undertaking the action is unwilling to implement the RPAs, it must either refrain from the action or seek an exemption from the ESA requirements by applying to the Executive Branch cabinet-level Endangered Species Committee (often referred to as the “god squad”). Enforcement
Section 11 of the ESA provides both civil and criminal penalties for violations of the ESA. These penalties include fines ranging from $500 to $50,000 and jail terms from 6 months to a
0839_frame_C33.fm Page 746 Tuesday, May 22, 2001 1:38 PM
746
CRC Handbook of Marine Mammal Medicine
year. In addition, the ESA allows citizens to bring suit to enforce its provisions against anyone that a citizen believes is violating the law. Implementation of the Convention on International Trade in Endangered Species of Wild Fauna and Flora
The ESA, under Section 8, implements the U.S. international obligations as a party to the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES). Originally established on March 6, 1973, CITES went into effect July 1, 1975, and now has approximately 150 countries as signatory parties. The primary purpose of CITES is to regulate international trade in wild animals and plants, which are listed in three appendices to the Convention. Species threatened with extinction are listed in Appendix I and, with few exceptions, trade involving these species is prohibited. Appendix II lists those species that may become threatened. The Convention allows for the parties regulating the export of species domestically to obtain support of other parties in doing so. Marine mammal species that are currently listed as being endangered or threatened under the ESA, or that are included in the CITES Appendixes, are given in Table 1. TABLE 1 Status of Marine Mammals Common Name
Scientific Name
ESA Status
CITES Appendix
E
I I I
A. CETACEA Mysticeti: The Baleen Whales Balaenidae: The Bowhead and Right Whales Bowhead whale Southern right whale Northern right whale
Balaena mysticetus Eubalaena australis Eubalaena glacialis
E
Balaenopteridae: The Rorqual Whales Minke whale Sei whale Bryde’s whale Blue whale Finback whale Humpback whale
Balaenoptera acutorostrata Balaenoptera borealis Balaenoptera edeni Balaenoptera musculus Balaenoptera physalus Megaptera novaeangliae
E E E
I I I I I I
R
I
E
Eschrichtiidae: The Gray Whale Gray whale
Eschrichtius robustus Neobalaenidae: The Pygmy Right Whale
Pygmy right whale
Caperea marginata
I
Odontoceti: The Toothed Whales, Dolphins and Porpoises Delphinidae: The Dolphins Commerson’s dolphin Black dolphin Heaviside’s dolphin Hector’s dolphin Common dolphin; saddleback dolphin Pygmy killer whale Short-finned pilot whale Long-finned pilot whale
Cephalorhynchus commersonii Cephalorhynchus eutropia Cephalorhynchus heavisidii Cephalorhynchus hectori Delphinus delphis Feresa attenuata Globicephala macrorhynchus Globicephala melas (Globicephala meleana)
II II II II II II II II
0839_frame_C33.fm Page 747 Tuesday, May 22, 2001 1:38 PM
747
U.S. Federal Legislation Governing Marine Mammals
TABLE 1 Status of Marine Mammals (continued) Scientific Name Risso’s dolphin Fraser’s dolphin; Sarawak dolphin Atlantic white-sided dolphin White beaked dolphin Peale’s dolphin Hourglass dolphin Pacific white-sided dolphin Dusky dolphin Northern right whale dolphin Southern right whale dolphin Irrawaddy dolphin Killer whale; Orca Melon-headed whale False killer whale Tucuxi Indo-Pacific humpbacked dolphin Atlantic humpbacked dolphin Pantropical spotted dolphin Clymene dolphin Striped dolphin Atlantic spotted dolphin Spinner dolphin Rough-toothed dolphin Bottlenose dolphin
Grampus griseus Lagenodelphis hosei Lagenorhynchus acutus Lagenorhynchus albirostris Lagenorhynchus australis Lagenorhynchus cruciger Lagenorhynchus obliquidens Lagenorhynchus obscurus Lissodelphis borealis Lissodelphis peronii Orcaella brevirostris Orcinus orca Peponocephala electra Pseudorca crassidens Sotalia fluviatilis Sousa chinensis Sousa teuszii Stenella attenuata Stenella clymene Stenella coeruleoalba Stenella frontalis Stenella longirostris Steno bredanensis Tursiops truncatus
ESA Status
D(1) D(2)
CITES Appendix II II II II II II II II II II II II II II II II II II II II II II II II
Monodontidae: The Narwhal and White Whale Beluga whale; white whale Narwhal
Delphinapterus leucas Monodon monoceros
D(4)
II II
Phocoenidae: The Porpoises Spectacled porpoise Finless porpoise Harbor porpoise Vaquita; cochito Burmeister’s porpoise Dall’s porpoise
Australophocaena dioptrica Neophocaena phocaenoides Phocoena phocoena Phocoena sinus Phocoena spinipinnis Phocoenoides dalli
E
II I II I II II
Physeteridae: The Sperm Whales Pygmy sperm whale Dwarf sperm whale Sperm whale
Kogia breviceps Kogia simus Physeter macrocephalus (Physeter catodon)
E
II II I
Platanistidae: The River Dolphins Amazon River dolphin Chinese River dolphin; Beiji Ganges River dolphin; Ganges susu Indus River dolphin; Indus susu Franciscana; La Plata River dolphin
Inia geoffrensis Lipotes vexillifer Platanista gangetica Platanista minor Pontoporia blainvillei
E E
II I I I II (Continued)
0839_frame_C33.fm Page 748 Tuesday, May 22, 2001 1:38 PM
748
CRC Handbook of Marine Mammal Medicine
TABLE 1 Status of Marine Mammals (continued) Scientific Name
ESA Status
CITES Appendix
Ziphiidae: The Beaked Whales Arnoux’s beaked whale Baird’s beaked whale Northern bottlenose whale Southern bottlenose whale Sowerby’s beaked whale Andrew’s beaked whale Hubbs’ beaked whale Blainville’s beaked whale; dense beaked whale Gervais’ beaked whale; Gulf Stream beaked whale Ginkgo-toothed whale Gray’s beaked whale Hector’s beaked whale Strap-toothed whale True’s beaked whale Longman’s beaked whale Pygmy beaked whale Stejneger’s beaked whale Shepherd’s beaked whale Cuvier’s beaked whale
Berardius arnuxii Berardius bairdii Hyperoodon ampullatus Hyperoodon planifrons Mesoplodon bidens Mesoplodon bowdoini Mesoplodon carlhubbsi Mesoplodon densirostris
II II I I II II II II
Mesoplodon europaeus
II
Mesoplodon ginkgodens Mesoplodon grayi Mesoplodon hectori Mesoplodon layardii Mesoplodon mirus Mesoplodon (Indopacetus) pacificus Mesoplodon peruvianus Mesoplodon stejnegeri Tasmacetus shepherdi Ziphius cavirostris
II II II II II II II II II II
B. CARNIVORA 1. PINNIPEDIA: THE SEALS, SEA LIONS, AND WALRUSES Odobenidae: The Walruses Walrus
Odobenus rosmarus
III
Otariidae: The Eared Seals South American fur seal New Zealand fur seal; West Australian fur seal Galapagos fur seal Antarctic fur seal Juan Fernández fur seal South African fur seal; Cape fur seal Guadalupe fur seal Subantarctic fur seal; Amsterdam Island fur seal Northern fur seal Steller sea lion; northern sea lion (southeastern population) Steller sea lion; northern sea lion (western population) Australian sea lion South American sea lion Hooker’s sea lion California sea lion
Arctocephalus australis Arctocephalus forsteri
II II
Arctocephalus galapagoensis Arctocephalus gazella Arctocephalus philippii Arctocephalus pusillus Arctocephalus townsendi Arctocephalus tropicalis
II II II II I II
T
Callorhinus ursinus Eumetopias jubatus
D(3) T
Eumetopias jubatus
E
Neophoca cinerea Otaria byronia Phocarctos hookeri Zalophus californianus
0839_frame_C33.fm Page 749 Tuesday, May 22, 2001 1:38 PM
749
U.S. Federal Legislation Governing Marine Mammals
TABLE 1 Status of Marine Mammals (continued) Scientific Name
ESA Status
CITES Appendix
E E E
II I I I
Phocidae: The True Seals Hooded seal Bearded seal Gray seal Leopard seal Weddell seal Crabeater seal Northern elephant seal Southern elephant seal Mediterranean monk seal Hawaiian monk seal Caribbean monk seal; West Indian monk seal Ross seal Caspian seal Ribbon seal Harp seal Ringed seal Saimaa seal (subspecies of ringed seal) Larga seal; spotted seal Baikal seal Harbor seal
Cystophora cristata Erignathus barbatus Halichoerus grypus Hydrurga leptonyx Leptonychotes weddellii Lobodon carcinophagus Mirounga angustirostris Mirounga leonina Monachus monachus a Monachus schauinslandi Monachus tropicalis Ommatophoca rossii Phoca caspica Phoca fasciata Pagophilus groenlandicus Phoca hispida Phoca hispida saimensis Phoca largha Phoca sibirica Phoca vitulina
E
2. NONPINNIPEDS Mustelidae: The Otters Sea otter Marine otter
b
T E
Enhydra lutris b Lutra (Lontra) felina
I I
Ursidae: The Bears Polar bear
b
II
Ursus maritimus
C. SIRENIA: MANATEES AND DUGONG Trichechidae: The Manatees Amazonian manatee West Indian manatee; Florida manatee West African manatee
b
E E
I I
T
II
b
E
b
I (NonAustralian) II (Australian)
Trichechus inunguis b Trichechus manatus b
Trichechus senegalensis Dugongidae: The Dugongs
Dugong
Dugong dugon
Dugong
Dugong dugon
E
Steller’s sea cow (Extinct)
Hydrodamalis gigas
X
Key: E = endangered; T = threatened; D = depleted [D(1) = ETP stock; D(2) = Mid-Atlantic coastal migratory stock; D(3) = Pribilof Island stock; D(4) = Cook Inlet stock]; R = recovered; X = extinct. Note: All species are under the jurisdiction of the NMFS except as noted below: a Both the NMFS and the FWS share jurisdiction. b Under FWS jurisdiction only.
0839_frame_C33.fm Page 750 Tuesday, May 22, 2001 1:38 PM
750
CRC Handbook of Marine Mammal Medicine
Individuals desiring to file a CITES permit application for a marine mammal should first obtain a copy of the regulations at 50 CFR 23 for the Endangered Species Convention in addition to those listed for the ESA. CITES permit applications are submitted either to the NMFS or the FWS, and permits are issued either by the Secretary of Commerce or the Secretary of the Interior, depending on the marine mammal species involved (see Table 2 for the NMFS and FWS contact information). It is important to note that 50 CFR Parts 13 and 17, 50 CFR 222, and 50 CFR 23 are highly integrated regulations and all criteria must be met in filing an application for either an ESA or CITES permit.
The Marine Mammal Protection Act (16 USC 1361–1362, 1371–1384, 1401–1407; 50 CFR 216 (NMFS); 50 CFR 18 (FWS))* The MMPA is the most comprehensive marine mammal conservation and management legislation in the world. Passed in 1972 to rectify the consequences of human impacts on marine mammals, the MMPA, enforced by the U.S. DOC and DOI, governs every interaction within U.S. jurisdiction between an individual person and a marine mammal. Its purpose is to protect marine mammal species of “great international significance, aesthetic and recreational as well as economic.” The species included under the MMPA are whales, dolphins, porpoises, seals, sea lions, walruses, manatees, dugongs, sea otters, and polar bears. The MMPA Moratorium on Taking
The goal of the MMPA is to protect and promote the growth of marine mammal populations commensurate with sound policies of resource management and to “maintain the health and stability of the marine ecosystem.” Congress also mandated that marine mammals be protected and managed so that they continue to be significant parts of their ecosystems, and that they should not be allowed to “diminish below their Optimum Sustainable Population” (OSP). A species or population stock is below its OSP level if it is listed as endangered or threatened under the ESA or if it is designated as “depleted” under the MMPA. “Optimum Sustainable Population means, with respect to any population stock, the number of animals that will result in the maximum productivity of the population or the species, keeping in mind the carrying capacity of the habitat and the health of the ecosystem of which they form a constituent element” (16 USC 1362(9); 50 CFR 216.3 (1994)). Thus, OSP is a population size that falls within a range, from the population level of a given species or stock that is the largest supportable within the ecosystem, to the population level that results in maximum net productivity. Maximum net productivity is the greatest net annual increment in population numbers or biomass resulting from additions to the population due to reproduction and/or growth, less losses due to natural mortality. Through the MMPA, Congress sought to achieve broad protection for marine mammals by establishing a moratorium on importation and take. The MMPA also states that the “incidental kill or incidental serious injury of marine mammals permitted in the course of commercial fishing operations be reduced to insignificant levels approaching a zero mortality and serious injury rate” or zero mortality rate goal (ZMRG) (16 USC 1387). Exemptions and Permits for Incidental Take
The MMPA allows the Secretary of Commerce or the Interior to waive the moratorium, by issuing a permit for taking a marine mammal species, if the best available scientific evidence reveals that such take would not disadvantage a specific marine mammal population. Actions that may receive a permit include the incidental taking of marine mammals in commercial fisheries, the taking and importation for the purpose of scientific research, public display, or *Available at http://www.nmfs.noaa.gov.
0839_frame_C33.fm Page 751 Tuesday, May 22, 2001 1:38 PM
751
U.S. Federal Legislation Governing Marine Mammals
TABLE 2 NMFS and FWS Offices NATIONAL MARINE FISHERIES SERVICE Headquarters Office of Protected Resources Permits Division, F/PR1 National Marine Fisheries Service, NOAA 1315 East-West Highway Silver Spring, MD 20910-3226 (301) 713-2289 Regional Offices AK Regional Administrator, Alaska Region National Marine Fisheries Service, NOAA P.O. Box 21668 Juneau, AK 99802-1668 Phone (907) 586-7221; Fax (907) 586-7249
AZ, CA, HI, NV, NM Regional Administrator, Southwest Region National Marine Fisheries Service, NOAA 501 West Ocean Blvd., Suite 4200 Long Beach, CA 90802-4213 Phone (310) 980-4001; Fax (310) 980-4018
CT, DE, IL, IN, ME, MD, MA, MI, MN, NH, NJ, NY, OH, PA, RI, VT, VA, WV, WI Regional Administrator, Northeast Region National Marine Fisheries Service, NOAA One Blackburn Drive Gloucester, MA 01930-2298 Phone (508) 281-9250; Fax (508) 281-9371
Pacific Area Office Coordinator, Pacific Area Office National Marine Fisheries Service, NOAA 2570 Dole Street, Room 106 Honolulu, HI 96822-2396 Phone (808) 943-1221; Fax (808) 943-1240
CO, ID, MT, ND, OR, SD, UT, WA, WY Regional Administrator, Northwest Region National Marine Fisheries Service, NOAA 7600 Sand Point Way, NE, BIN C15700, Bldg. 1 Seattle, WA 98115-0070 Phone (206) 526-6150; Fax (206) 526-6426
National Marine Mammal Laboratory Director, National Marine Mammal Laboratory National Marine Fisheries Service, NOAA 7600 Sand Point Way, NE, BIN C15700, Bldg. 1 Seattle, WA 98115-0070 Phone (206) 526-6150; Fax (206) 526-6426
AL, AR, FL, GA, IA KS, KY, LA, MS, MO, NC, NE, OK, PR, SC, TN, TX, VI Regional Administrator, Southeast Region National Marine Fisheries Service, NOAA 9721 Executive Center Drive North St. Petersburg, FL 33702-2432 Phone (813) 570-5301; Fax (813) 570-5300 U.S. FISH AND WILDLIFE SERVICE OFFICES Washington Office U.S. Fish and Wildlife Service Division of Endangered Species Main Interior Building 1849 C Street, NW Washington, D.C. 20240 Phone (202) 208-4545 (Continued)
0839_frame_C33.fm Page 752 Tuesday, May 22, 2001 1:38 PM
752
CRC Handbook of Marine Mammal Medicine
TABLE 2 NMFS and FWS Offices (continued) Regional Offices Region 1 (WA, OR, CA, ID, NV, HI) Eastside Federal Complex 911 N.E. 11th Avenue Portland, OR 97232 Phone (503) 231-6121
Region 5 (ME, NH, MA, RI, CT, NY, NJ, DE, MD, VA, VT, NY, PA, WV) 300 Westgate Center Drive Hadley, MA 01035 Phone (413) 253-8200
Region 2 (AZ, NM, TX, OK) 500 Gold Avenue, SW P.O. Box 1306 Albuquerque, NM 87103 Phone (505) 248-6911
Region 6 (ND, SD, NE, KS, CO, UT, WY, MT) Denver Federal Building 134 Union Blvd. Lakewood, CO 80228 P.O. Box 25486 Denver, CO 80225 Phone (303) 236-7917
Region 3 (OH, MI, IN, IL, WI, MN, IA, MO) Federal Building Fort Snelling Twin Cities, MN 55111 Phone (612) 713-5360
Region 7 (AK) 1011 E. Tudor Road Anchorage, AK 99503 Phone (907) 786-3909
Region 4 (NC, SC, GA, FL, KY, TN, AL, MS, AR, LA, PR, VI) 1875 Century Blvd. Suite 200 Atlanta, GA 30345 Phone (404) 679-7287
National Fish and Wildlife Forensics Laboratory 1490 East Main Street Ashland, OR 97520 Phone (541) 482-4191
enhancing the recovery of a population, and/or the importation of polar bear parts taken from sport hunts in Canada. The Marine Mammal Permits and Documentation Division of the NMFS develops and implements policies, procedures, and regulations for permits to take marine mammals for research, public display, commercial and educational photography, and enhancing the survival or recovery of a marine mammal species or stock. It also issues general authorizations for scientific research; formulates, organizes, and executes the national program for tracking captive marine mammals; and assists the various NMFS regional offices with marine mammal viewing guidelines. The Permits Division is also responsible for coordinating with the FWS on permits for imports and exports of species listed under the CITES and serves as a liaison on marine mammal permit matters with the MMC, the FWS, the APHIS, other federal agencies, the academic community, public display institutions, and environmental and animal welfare organizations. The NMFS issues permits for scientific research and enhancement of protected marine species and for the capture and first-time import of marine mammals for public display. Prior to issuing a permit, NMFS requests public comment on all permit applications through publication of a notice in the Federal Register. Additionally, the NMFS maintains an inventory of marine mammals held for public display, scientific research, or enhancement. Permit applications for activities involving marine mammal species are submitted to the FWS or the NMFS (see Table 2). Detailed instructions required for a marine mammal permit application are available from the Office of Protected Species of the NMFS or on the NMFS Web site:
0839_frame_C33.fm Page 753 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
753
http://www.nmfs.noaa.gov
or from the FWS or its Web site: http://www.fws.gov
Individuals contemplating submitting a permit application under the MMPA are advised to obtain a copy of the MMPA (USC 1361-1362, 1371-1384, 1401-1407) and the respective regulations relative to the NMFS and the FWS, and to call their local NMFS or FWS office to discuss their application with the permitting staff, before beginning to compile their application materials. Prospective applicants should realize that processing of applications takes a minimum of 120 days. Therefore, it is recommended that applications be submitted well in advance of the proposed activity. Helpful instructions for submitting a marine mammal permit application are available from the Office of Protected Species, the NMFS. As with the ESA, first-time applicants will find it very helpful to recruit the aid of an individual experienced with the filing of such applications and/or to contact their regional NMFS marine mammal program leader. The permitting processes for marine mammals under the MMPA and the ESA are currently under extensive review by the NMFS and the FWS. The reader is advised to ensure that the information provided is up-to-date, accurate, and that the application language has been reviewed by the government agency before submission of the permit application, if at all possible. Reauthorizations of the MMPA
The MMPA was passed in 1972 and was amended in 1973 (primarily to incorporate the CITES provisions), and again in 1976, 1981, 1988, and 1994 (Marine Mammal Commission, 1999). The November 1988 MMPA amendments by Congress primarily deal with marine mammal public display, survival enhancement of population stocks, and marine mammal–fisheries interactions (Marine Mammal Commission, 1989). In 1994, Congress enacted amendments that clarified the role of the NMFS and the FWS with regard to captive marine mammals, limiting the authority of these agencies over marine mammals once the animals were removed from the wild, and vesting the humane handling, housing, care, treatment, and transportation of captive marine mammals with the APHIS under the Animal Welfare Act (AWA) (Marine Mammal Commission, 1995). The amendments also simplified the procedures for authorizing transfers of marine mammals among display facilities. The NMFS is in the process of proposing regulations to comply with new and changed provisions in the MMPA as a result of the 1994 Amendments. Marine Mammal Strandings and Health
One of the unique aspects of the MMPA is that it grants the legal authority allowing the NMFS to enter into agreements necessary to carry out the goals of the MMPA. This provision of the MMPA allows for the taking of stranded marine mammals without prior authority or permit being required, provided that such taking is for the protection of the welfare of the marine mammal, the health and welfare of the public, or the nonlethal removal of nuisance animals. The provision only applies to designated local, state, or federal governments. A parallel exception for endangered or threatened species is found in 50 CFR 17.21(c). A letter of agreement (LOA) is usually required to engage in the taking of stranded marine mammals, and is obtained by writing one’s Regional Office of the NMFS. There is a national Marine Mammal Stranding Network coordinated through the regional offices of the NMFS as well (see Chapter 4, Stranding Networks). This network largely comprises volunteer coalitions of private individuals, researchers, aquaria and oceanaria, research laboratories, and other similar institutions interested in marine mammal strandings. Participants operate under LOAs from regional NMFS offices. On November 4, 1992, the Marine Mammal Health and Stranding Response Act was signed into law. The two main reasons for the law were (1) concern that knowledge of the connection
0839_frame_C33.fm Page 754 Tuesday, May 22, 2001 1:38 PM
754
CRC Handbook of Marine Mammal Medicine
between marine mammal health and the various components of the marine environment was insufficient to allow adequate understanding of the causes of unusual marine mammal mortality events; and (2) concern that responses to unusual marine mammal mortality events were often uncoordinated as a result of insufficient contingency planning. The law, now Title IV of the MMPA, mandates that the Secretary of Commerce establish a “Marine Mammal Health and Stranding Response Program” to (1) facilitate the collection and dissemination of data on the health and health trends of marine mammals in the wild; (2) correlate the data on marine mammal health with available data on physical, chemical, and biological environmental parameters; and (3) coordinate effective responses to unusual mortality events (see Chapter 5, Unusual Mortality Events). Title IV also requires the Secretary of Commerce to undertake the following: • Collect and update information on procedures for rescuing and rehabilitating stranded marine mammals, and for collecting, preserving, and ensuring the integrity of marine mammal tissue samples; • Develop objective criteria for determining the point at which a rehabilitated marine mammal can be returned to the wild; • Compile and analyze, on a regional basis, information on stranded marine mammals, including the species and numbers involved, the condition of the animals, and the causes of any illnesses or death; • Establish an expert working group (the Working Group on Marine Mammal Unusual Mortality Events) to provide advice and guidance for determining when unusual mortality events are occurring, for developing a contingency plan to respond to such events, and for determining when actions taken in response to such events are no longer needed; • Establish a marine mammal Unusual Mortality Event Fund to compensate stranding network participants and others for costs incurred while responding to unusual mortality events; • Develop a national contingency plan to be used to respond to unusual marine mammal mortalities; • Formalize the establishment of a National Marine Mammal Tissue Bank, which is to contain samples collected from marine mammals involved in unusual mortality events, taken by Alaskan natives for subsistence purposes, and/or taken incidental to commercial fisheries; along with guidance for tissue collection, preservation, archiving, and quality control; and • Issue guidance for the analyses of tissue samples, to monitor and measure overall health trends in representative marine mammal populations, to identify the levels and effects of potentially harmful contaminants, and to determine the frequency and causes of any abnormal lesions (see Chapter 21, Necropsy; Chapter 23, Noninfectious Diseases).
The NMFS has established the Marine Mammal Health and Stranding Response Program, and this has greatly improved the response of stranding networks to stranded marine mammals, as well as emergency responses to unusual mortality events and investigations into the causes of such events. The program has also resulted in the establishment of a National Marine Mammal Tissue Bank, numerous training sessions for stranding network participants, and a greater awareness of the importance of health-related research to the conservation of marine mammal populations. The Marine Mammal Health and Stranding Response Act is a program vital to the NMFS effort to conserve and protect marine mammals. Although the MMPA and the ESA tend to support the activities of one another, the MMPA is more stringent in that it does not allow for the live taking of depleted marine mammal stocks, except for scientific research or enhancement of survival or recovery of a species or stock. For example, public display is not an authorized activity for a depleted species. Similarly, the MMPA does not allow the importation (take) of any pregnant, nursing, or juvenile (less than 8 months old) marine mammal, unless such importation is necessary for the protection or welfare of the animal.
0839_frame_C33.fm Page 755 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
755
The Animal Welfare Act (7 USC 2131–2156)* The AWA was first passed in 1966, and amended in 1970, 1976, 1985, and 1990. The primary function of the AWA is to ensure that animals intended for use in research facilities, public displays, and exhibitions, or as pets are provided with humane care and treatment. Hence, the AWA is a protection act that focuses on the individual animal and does not emphasize the conservation of species or habitat. The AWA is administered by the USDA, and implemented by the APHIS. Although originally covering animals traditionally used as pets and research specimens (e.g., dogs, rabbits, etc.), the AWA allows for inclusion of other animals as deemed appropriate by the Secretary of Agriculture. Marine mammals, along with most other warmblooded exotic animals, have been included under the AWA, and regulations for their care have been in effect since September 20, 1979. The Law
The AWA requires that minimum standards of care and treatment be provided for certain animals bred for commercial sale, used in research, transported commercially, or exhibited to the public. Individuals who operate facilities in these categories must provide their animals with adequate care and treatment in the areas of housing, handling, sanitation, nutrition, water, veterinary care, and protection from extreme weather and temperatures. Although federal requirements establish acceptable regulations, they are not ideal and these businesses are encouraged to exceed the specified minimum regulations. Licensing and Registration
The AWA also requires that all individuals, businesses, and organizations dealing with animals covered under the law must be licensed or registered with the APHIS. Marine mammal rescue and rehabilitation facilities are exempt from these requirements; however, the NMFS is devising its own set of facility regulations for these facilities. Research Facilities
Research facilities must establish an Institutional Animal Care and Use Committee (IACUC) to oversee the use of animals in experiments. This Committee is responsible for ensuring that the facility remains in compliance with the AWA and provides documentation to the APHIS in all areas of compliance. The Committee must be composed of at least three members, including one veterinarian and one person who is not affiliated with the facility in any way. The the AWA does not permit the APHIS to interfere with research procedures or experimentation, only noncompliance. Regulated research facilities include hospitals, colleges and universities, diagnostic laboratories, and many private firms in the pharmaceutical and biotechnology industries. AWA Enforcement
The APHIS ensures that all regulated transportation companies, exhibitors, and research facilities are licensed or registered. APHIS also searches for unlicensed or unregistered facilities. Before the APHIS will issue a license, the applicant must be in compliance with all regulations under the AWA. To ensure that all licensed and registered facilities continue to comply with the AWA, the APHIS inspectors make unannounced inspections at least once annually. If an inspection reveals deficiencies in meeting the AWA rules and regulations, the inspector instructs the facility to correct the problems within a given time frame. If deficiencies remain
*Available at http://www.aphis.usda.gov.
0839_frame_C33.fm Page 756 Tuesday, May 22, 2001 1:38 PM
756
CRC Handbook of Marine Mammal Medicine
uncorrected at the unannounced follow-up inspection, the APHIS documents the deficiencies of the facility and considers possible legal action. The APHIS also conducts reviews and investigates alleged violations. Some cases are resolved with Official Notices of Warning or agency stipulation letters, which set civil penalties for the infractions. Civil penalties include cease-and-desist orders, fines, and license suspensions or revocations. If APHIS officials determine that an alleged AWA violation warrants additional action, the APHIS submits all evidence to the USDA for further legal review. Regulations
The regulations under which the AWA is implemented are 9 CFR 1, 2, 3, and 4. The APHIS receives its authority through the APHIS Authorization Act (21 USC 134-134h, 101-105, and 111). Subject to the general provisions of the AWA, the captive care and maintenance of marine mammals are regulated through 9 CFR 3, Subpart E—Specifications for the Humane Handling, Care, Treatment, and Transportation of Marine Mammals (Regulations). In 1995 and 1996, the APHIS convened various marine mammal experts, industry specialists, and animal welfare organizations to create a negotiated rule-making advisory committee for marine mammals. The rule-making process involved several sessions designed to allow the committee to create regulations that reflected in great detail the type of facilities in which marine mammals may be kept, the minimum space requirements for housing the various species of marine mammals, feeding, sanitation, attendant and employee qualifications, veterinary care, other related husbandry care, carrier, intermediate handler restrictions and requirements, and transport handling requirements. Although there are many general provisions in the regulations (i.e., housekeeping, pest control, etc.), there are specific provisions that mandate measurable levels that must be met or maintained. For example, frozen food to be fed to marine mammals must not be stored in temperatures above 18°C (0°F), unless being thawed for use; kitchen areas must be sanitized using specific procedures; thawed food must be kept iced or refrigerated and must be fed within 24 hours of removal from the freezer. Particular attention is paid to the water quality parameters of primary enclosures (see Chapter 35, Water Quality). The coliform bacteria count of the water must not exceed a most probable number (MPN of bacteria in 100 ml of water) of 1000; if it does, a prescribed method for further water testing is given. It is therefore imperative that individuals responsible for the care and maintenance of captive marine mammals be conversant with both the intent and the particulars of the AWA regulations. As stated above, facilities that house or otherwise handle marine mammals must be licensed by the APHIS through the AWA, and are subject to routine inspection to ensure compliance and maintenance of their licenses. If one wishes to house marine mammals, the facility for doing so must first be inspected and certified that it meets the above regulations before the permit to acquire the particular marine mammals under the MMPA can be granted. Such certification usually only applies to the marine mammal species in question, and recertification is required if, at some time in the future, another species is under consideration for holding at the facility. Facilities under construction can receive provisional certification to avoid holding up the processing of permit applications under the MMPA, provided that they are inspected and approved prior to marine mammals arriving at the facility. Space Requirements
Of all the restrictions given in the regulations for marine mammals, perhaps the most complex, frustrating, and confusing requirements are those for minimum space requirements. This is primarily because the term marine mammal covers a very broad and diverse group of animals,
0839_frame_C33.fm Page 757 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
757
and the requirements were originally developed along traditional taxonomic divisions of cetaceans, pinnipeds, sirenians, sea otters, and polar bears. The cetacean and pinniped taxonomic groups are then divided into Group I and Group II categories. Group I animals tend to be more “inshore” and, therefore, can be maintained in smaller, shallower spaces. Group II animals tend to be more “offshore,” and tend to require larger, deeper spaces. Sea otters and polar bears have additional requirements for dry resting spaces. Mathematical formulas are provided in the body of the regulations whereby the calculation for minimum surface area, minimum volume, minimum depth, and minimum horizontal dimension (MHD) of the pool can be calculated for the minimum holding of two animals. Similar formulas are provided for the dry resting area requirements, with Groups I and II each having its own formulas for calculating specific requirements. Additional formulas are provided for calculating space for more than two marine mammals of the same species in a single enclosure, or when there is a mixture of Group I and Group II marine mammals in the same enclosure. It is important to note that a single marine mammal of any species requires the same space as that required for two marine mammals of the same species. If two marine mammals of different species are housed together, the species requiring the larger minimum space is to be used to determine if the enclosure meets the space requirements. A particularly important measurement is the MHD used to calculate the surface area and pool volume of marine mammal enclosures. The MHD is derived from multiplying a specific factor that differs between Group I and II marine mammals by the average adult length of the particular marine mammal under consideration. The average adult length is based on both the species and the sex of the animal for pinnipeds, but only on the species for other marine mammals. The average adult length is also used for calculating the minimum depth of the enclosure pools (for specific formulas, see the APHIS regulations at 9 CFR 3.100). Marine mammals may be temporarily housed in enclosures not in compliance with the space requirement of the regulations for routine husbandry and training purposes, or for the transfer of animals. Such enclosures may also be used when the professional staff deems it necessary (e.g., isolation and medical treatment). Theoretically, marine mammals housed in temporary situations, such as rehabilitation facilities, are currently outside the regulatory authority of the AWA. The APHIS does investigate complaints about temporary facilities, and does become involved if the animals at such facilities are on public display or being used in research or for teaching purposes. However, the NMFS or the FWS (depending on the marine mammal species involved) takes responsibility for issuing rehabilitation centers LOAs granting permission to rescue and medically care for beached or stranded marine mammals (see Chapter 4, Stranding Networks). Overlap among the Agencies and the Various Laws
Because of the tremendous amount of administrative and enforcement overlap among the NMFS, the FWS, and the APHIS in administering the laws affecting marine mammals, these three agencies have entered into a memorandum of understanding that outlines their relative responsibilities and authorities under the MMPA and the AWA. The primary goals of the memorandum are to ensure that regulations for marine mammals are applied consistently; that appropriate and consistent guidance is provided to individuals responsible for marine mammals in captivity; and that the responsibilities of the respective agencies are met in an effective manner with a minimum duplication of effort. The memorandum of understanding became effective in September 1979, and remains in effect until one party to the agreement notifies the others of its intent to terminate the agreement.
0839_frame_C33.fm Page 758 Tuesday, May 22, 2001 1:38 PM
758
CRC Handbook of Marine Mammal Medicine
The Lacey Act of 1901 (16 USC 3371–3378) The Lacey Act (LA) was first passed in 1901 (18 USC 43-44, the original Lacey Act) and was amended in 1948, 1949, 1960, 1969, and most recently in 1981(16 USC 3371–3378; Pub. L. 97-79, as amended). Elements of the repealed acts addressing illegal trade of fish, wildlife, and plants were consolidated into the 1981 amendments. Thus, the LA in its current form primarily prohibits the importation, exportation, transportation, selling, receiving, acquiring, or purchasing of any fish, wildlife, or plant that has been taken or possessed in violation of any U.S. or Indian Tribal law, treaty, or regulation. It also prohibits any individual from engaging in interstate or foreign commerce with any fish or wildlife taken, possessed, transported, or sold in violation of any law or regulation of any state or in violation of any foreign law. The Secretary of Commerce is authorized to issue regulations including, but not limited to, cooperating with the Secretary of the Interior for the marking and labeling of packages containing fish or wildlife. The Secretary of Commerce is authorized to assess civil penalties not in excess of $10,000 per violation by persons engaging in conduct prohibited by this Act. Civil penalties assessed by the Secretary may be reviewed by the appropriate district court of the United States within 30 days of the assessment. The Act further provides for criminal action to be taken against persons found to be in violation of the LA. In addition, the LA allows for the seizure of all vessels, vehicles, aircraft, and other equipment used to aid in the criminal violation of the LA. Pursuant to the LA, the Secretary of Commerce may make available rewards for information furthering the intent of this Act. The LA does not specifically address any concerns regarding marine mammals, except where they would have been obtained or transported illegally (50 CFR 246).
The Fur Seal Act (16 USC 1151–1153) The Fur Seal Act (FSA) was passed in 1966 for the implementation of the Interim Convention on Conservation of North Pacific Fur Seals. The provisions of the FSA only apply to the north Pacific fur seal (Callorhinus ursinus; northern fur seal). The regulations that implement the FSA, 50 CFR 216, essentially allow for the administration of the Pribilof Islands, the taking of fur seals, and the procedures for issuing, amending, or rescinding display permits for fur seals. The FSA is administered through a permit process nearly identical to the one for the MMPA, and the instructions, format, and procedures used in submitting, reviewing, and approving such permits are likewise similar.
Conclusion This section has briefly discussed the laws and regulations of the U.S. federal government regulating the activities typically associated with marine mammals. It addresses existing federal legislation, and does not include any reference to the many local and state laws that may also be applicable. Many local and state governments have laws and regulations (in addition to the federal laws discussed in this section) that provide for the protection and humane treatment of animals. At present, federal marine mammal law preempts any state law, unless the state requests a return of power and the federal government concurs and grants such. Some coastal states, particularly Florida and California, have developed state and local marine mammal programs, modeled after the MMPA and consistent with federal law; these programs are the initial steps in requesting return of marine mammal management to the state level. Other states, such as Massachusetts, are considering development of similar programs. Thus, it can be expected, sooner rather than later, that additional coastal states may move toward requesting return of laws that regulate marine mammal activities.
0839_frame_C33.fm Page 759 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
759
The number and scope of international agreements and conventions are continually expanding, as nations decide to cooperate more in worldwide efforts to conserve natural resources (Lyster, 1985). The International Convention for the Regulation of Whaling once applied only to the harvesting of whales; the parties to this Convention are now showing greater interest in other forms of human activity dealing with cetaceans in general, such as the effects of whalewatching boats, environmental contaminants, and climate change on various populations of whales. The Convention for the Conservation of Polar Bears and the Convention on the Conservation of Migratory Species of Wild Animals are international agreements among nations related to activities concerning specific marine mammals. Similarly, conventions such as the Convention for the Protection and Development of the Marine Environment of the Wider Caribbean Region (the Cartagena Convention), which calls for sweeping protection and preservation of large expanses of aquatic environments, will undoubtedly help to establish even greater conservation for the marine environment and marine mammals. Conservation of marine mammals is complicated by the fact that to conserve the world’s oceans, the laws and regulations governing marine species are in an almost constant state of change, evolving to achieve added conservation benefit, not only for marine mammals, but also for commercial fish stocks and marine ecosystems as a whole. Consequently, these laws and regulations will continue to be integral factors in marine mammal research, display, rehabilitation, and conservation. One must expect that marine mammal legislation in several forms will exist for some time to come and that these evolving processes are important tools for achieving the necessary protection for these animals and the habitats upon which they depend for survival. It is therefore important that anyone already or soon to become involved with marine mammals, be it for public display, scientific research, strandings, or recovery, have a basic familiarity with this increasingly important and growing body of legislation and regulations. This section is provided for information purposes only. Marine mammal legislation and regulation are of a rather fluid nature, and before fully relying on any portion of this chapter as presented, the reader should reference the official reports concerning the most current laws and implementing regulations in the USC and CFR, and the pertinent instructions and informational materials available from the governmental regulating agencies concerning marine mammals (see http://www.nmfs.noaa.gov and http://www.fws.gov).
Definitions and Abbreviations Pertaining to U.S. Marine Mammal Legislation Sara L. Shapiro and Nina M. Young
Administrative Procedures Act (5 USC 551–559, 701–706, 1305, 3105, 3344, 4301, 5362, 7521; 60 Stat. 237, as amended)–Public Law 79-404, signed June 11, 1946: Outlines administrative procedures that federal agencies must follow. These steps include identification of information to be made public; publication of material in the Federal Register; maintenance of records, including those involving certain meetings and hearings; attendance and notification requirements for specific meetings and hearings; issuance of licenses; and review of agency actions. Animal and Plant Health Inspection Service (APHIS): A division of the USDA that regulates the treatment of marine mammals held in captivity. The APHIS determines health standards, such as tank size and water quality, and implements the Animal Welfare Act. Animal Welfare Act (AWA) (7 U.S.C. 2131 et seq.): Federal statute created in 1966 to “(1) ensure that animals intended for use in research facilities or for exhibition purposes or for use as pets, are provided humane care and treatment, (2) assure the humane treatment
0839_frame_C33.fm Page 760 Tuesday, May 22, 2001 1:38 PM
760
CRC Handbook of Marine Mammal Medicine
of animals during transportation in commerce, and (3) protect the owners of animals from the theft of their animals by preventing the sale or use of animals which have been stolen.” The AWA is a protection act that focuses on the individual animal and does not emphasize the conservation of species or habitat. The responsibility for administering the AWA lies with the USDA, and is implemented by the APHIS. Bona fide research: Defined in the MMPA as scientific research on marine mammals, “the results of which likely would be accepted for publication in a refereed scientific journal; are likely to contribute to the basic knowledge of marine mammal biology or ecology; or are likely to identify, evaluate, or resolve conservation problems.” Code of Federal Regulations (CFR): Regulations created by various federal agencies to support and explain federal statutes (laws). For purposes of this document, the FWS and the NMFS have created wildlife and fisheries regulations to support and clarify sections of the MMPA and the ESA. The wildlife and fisheries regulations pertaining to marine mammals and endangered species can be found in 50 CFR 1–599. Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES): An international treaty that was enacted in 1973 to protect wildlife against overexploitation and to prevent international trade from threatening species with extinction. The CITES currently has a membership of over 150 countries. These countries regulate commercial international trade in an agreed-upon list of endangered species and monitor trade in other species that might become endangered. Under the CITES, a species is listed at one of three levels of protection. Each level has different permitting requirements (see Table 1). • Appendix I: • Appendix II: • Appendix III:
Species threatened with extinction that are or may be affected by trade. Species that may become threatened with extinction if not regulated. Species listed by a participating country for the purpose of obtaining international cooperation in controlling trade.
For more information on the CITES, visit its Web site: http://www.cites.org
or the FWS Web site: http://international.fws.gov/permits/permits.html
Endangered Species Act (ESA) (16 USC 1531 et seq.): Federal statute created in 1973 to “conserve to the extent practicable the various species of fish or wildlife and plants facing extinction … to provide a means whereby the ecosystems upon which endangered or threatened species depend may be conserved, to provide a program for the conservation of such endangered species and threatened species.” Enhancement permits: As defined in the MMPA, “permits issued for the recovery of a species or stock where taking or importation: a) is likely to contribute significantly to maintaining or increasing distribution or numbers necessary to ensure the survival or recovery of the species or stock; and b) is consistent with any conservation plan adopted by the Secretary (NMFS or FWS) for the species or stock, or if there is no conservation or recovery plan in place, with the Secretary’s evaluation of action required to enhance the survival or recovery of the species or stock.” Fur Seal Act (FSA) (16 USC 1151–1153): Statute that was enacted to administer fur seal activities on the Pribilof Islands in Alaska, the “take” of northern fur seals (Callorhinus ursinus), and the procedures for issuing, amending, or rescinding northern fur seal display permits.
0839_frame_C33.fm Page 761 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
761
Harass: Under the ESA, FWS regulations (50 CFR 17.3) define harass as “an intentional or negligent act or omission which creates the likelihood of injury to wildlife by annoying it to such an extent as to significantly disrupt normal behavioral patterns, which include, but are not limited to, breeding, feeding or sheltering.” Harassment: Specifically defined in the MMPA as any act of pursuit, torment, or annoyance that has the potential to injure (Level A) or disturb (Level B) a marine mammal or marine mammal stock in the wild. Harm: Under the ESA, FWS regulations (50 CFR 17.3) define harm as “an act which actually kills or injures wildlife. Such act may include significant habitat modification or degradation, where it actually kills or injures wildlife by significantly impairing essential behavioral patterns, including breeding, feeding or sheltering.” The Lacey Act (LA) (16 USC 3371–3378): Prohibits the import, export, transport, sale, receipt, acquisition, or purchase of any fish, wildlife, or plant that has been taken or possessed in violation of any U.S. or Indian Tribal law, treaty, or regulation. It also prohibits any individual from engaging in interstate or foreign commerce with any fish or wildlife taken, possessed, transported, or sold, in violation of any law or regulation of any state or in violation of any foreign law. Marine Mammal Commission (MMC): Established under Title II of the MMPA, the Commission is charged with the responsibility for developing, reviewing, and making recommendations on actions and policies for all federal agencies with respect to marine mammal protection and conservation and for carrying out a research program. The MMC is an independent agency of the U.S. Executive Branch and consists of three part-time commissioners, who are appointed by the President. The Commission makes appointments to the Committee of Scientific Advisors on Marine Mammals. Marine Mammal Protection Act (MMPA) (16 USC 1361 et seq.): The federal statute created in 1972 to conserve and protect marine mammals by regulating activities of U.S. citizens and activities of all persons carried on within the jurisdiction of the United States. National Marine Fisheries Service (NMFS) (or NOAA Fisheries): Part of the U.S. DOC NOAA, the NMFS administers NOAA programs that support the domestic and international conservation and management of living marine resources. NMFS provides services and products to support domestic and international fisheries management operations, fisheries development, trade and industry assistance activities, enforcement, protected species and habitat conservation operations, and the scientific and technical aspects of the NOAA marine fisheries program. National Oceanic and Atmospheric Administration (NOAA): Organization whose mission is to predict environmental changes, protect life and property, provide decision makers with reliable scientific information, foster global environmental stewardship, and conserve and wisely manage the nation’s coastal and marine resources. Take: • MMPA definition: “To harass, hunt, capture, or kill or attempt to harass, hunt, capture, or kill any marine mammal.” • ESA definition: “To harass, harm, pursue, hunt, shoot, wound, kill, trap, capture, or collect, or to attempt to engage in any such conduct.”
U.S. Fish and Wildlife Service (FWS): The service within the U.S. DOI that is the principal federal agency responsible for conserving, protecting, and enhancing fish, wildlife, and plants and their habitats for the continuing benefit of the American people. Among its key functions,
0839_frame_C33.fm Page 762 Tuesday, May 22, 2001 1:38 PM
762
CRC Handbook of Marine Mammal Medicine
the FWS enforces federal wildlife laws, protects endangered species, manages migratory birds, restores nationally significant fisheries, conserves and restores wildlife habitat such as wetlands, and helps foreign governments with their international conservation efforts. Contact Information
The FWS and the NMFS implement both the MMPA of 1972 (16 USC 1361 et seq.) and the ESA of 1973 (16 USC 1531 et seq.). The FWS Office of Management Authority (OMA) has jurisdiction over the following species: 1. 2. 3. 4. 5. 6. 7. 8.
Sea otter (Enhydra lutris) Marine otter (Lutra felina) Polar bear (Ursus maritimus) Walrus (Odobenus rosmarus) Dugong (Dugong dugon) Amazonian manatee (Trichechus inunguis) West African manatee (Trichechus senegalensis) West Indian manatee (Trichechus manatus)
The FWS/OMA also regulates the international trade of all marine mammal parts through the CITES. Please refer all questions to: Office of Management Authority U.S. Fish and Wildlife Service 4401 N. Fairfax Drive, Room 700 Arlington, VA 22203 Phone: (703) 358-2104 (800) 358-2104 Fax: (703) 358-2281 http://international.fws.gov/permits/permits.html
The NMFS regulates the “taking” of all other marine mammal species (i.e., cetaceans and all other pinnipeds). Refer all permitting questions regarding these species to: Permits Division (F/PR1) Office of Protected Resources National Marine Fisheries Service 1315 East-West Highway Silver Spring, MD 20910 Phone: (301) 713-2289 Fax: (301) 713-0376 http://www.nmfs.noaa.gov/prot_res/prot_res.html
Marine Mammal Permits: Frequently Asked Questions (FAQs) Sara L. Shapiro
Congress is required to reauthorize both the MMPA and the ESA every 5 years. The following questions (Q) and answers (A) may change as a result of these reauthorizations.
The Marine Mammal Stranding Networks 1. Q: Who can rescue and rehabilitate marine mammals? A: Persons wishing to rescue or rehabilitate distressed marine mammals must obtain an LOA from either FWS or NMFS. The only people not required to obtain an LOA are government
0839_frame_C33.fm Page 763 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
763
employees who are performing such activities in the course of their duties as employees. This requirement is further explained in Section 109(h) of the MMPA and in the Code of Federal Regulations (50 CFR 216.22). Rehabilitation can occur only at an APHIS-licensed facility. 2. Q: Who can euthanatize a marine mammal in severe distress? A: Section 109(h)(1) of the MMPA authorizes qualified Marine Mammal Stranding Network members to euthanatize humanely nonendangered marine mammals in severe distress. Only qualified stranding network members, government employees, and veterinarians who have special permission granted directly from either the FWS or the NMFS can euthanatize endangered marine mammals. This permission may be granted over the telephone. 3. Q: What are permits needed for, and what is the process involved in placing an animal in a public display facility rather than releasing it? A:
Section 104(c)(7) of the MMPA allows a person or facility to retain a nonreleasable marine mammal provided that person or facility obtains a public display, scientific research, or enhancement permit. Endangered or threatened marine mammals cannot be held in captivity for the purpose of public display.
For NMFS-regulated species (cetaceans and most pinnipeds): Ac c o r d i n g t o 5 0 C F R 216.27, any marine mammal held for rehabilitation must be released within 6 months unless the attending veterinarian determines that release would be (a) detrimental to the wild population; (b) detrimental to the animal; or (c) premature. The attending veterinarian must evaluate the animal every 6 months to determine whether or not it should be released. If the attending veterinarian determines that the animal should not be released, within 30 days the person with authorized custody must (1) request authorization (from the NMFS) to retain or transfer custody or (2) humanely euthanize or arrange for disposition as authorized by the NMFS. Any rehabilitated beached or stranded marine mammal that is neither endangered nor threatened and is placed in public display following a nonreleasability determination must be held in captive maintenance according to all the public display requirements (50 CFR 216.27(b)(5)). For FWS-regulated species: Contact the FWS/OMA ((800) 358-2104) for information about retaining nonreleasable marine mammals. 4. Q: Can a marine mammal bone found on the beach be kept by the finder? A: The following applies only to marine mammals that are neither endangered nor threatened. One may collect and keep any marine mammal hard parts (bones, teeth, or ivory) found on a beach or land within 0.25 mile of the ocean (including bays and estuaries). Within 30 days of collection, all parts must be identified and registered with either the NMFS or the FWS. Registration must include (a) owner’s name, (b) description of the part, and (c) the date and location of collection. One may not retain parts from endangered or threatened species and must relinquish them to either the FWS or the NMFS. For more information, see 50 CFR 216.26 and 50 CFR 18.26 or call FWS ((800) 358-2104) or NMFS ((301) 713-2289). 5. Q: Who is responsible for cleaning up a rotting marine mammal carcass from a beach? A: Local government officials are authorized to dispose of marine mammal carcasses and remains that will not be salvaged or cataloged for scientific research or enhancement purposes.
Scientific Research and Enhancement Permits 1. Q: What permits are necessary to ship marine mammal carcasses or parts around the country or the world?
0839_frame_C33.fm Page 764 Tuesday, May 22, 2001 1:38 PM
764
CRC Handbook of Marine Mammal Medicine
A: To transfer samples within the United States, one must possess a valid scientific research or enhancement permit. If one plans to import or export samples, one must also possess the appropriate CITES permit(s). It takes a minimum of 120 days to apply for and receive a scientific research or enhancement permit, and a minimum of 60 days for a CITES permit. Each CITES Appendix classification has different permitting requirements. To obtain the proper permits, contact the NMFS ((301) 713-2289) and the FWS OMA ((800) 358-2104). If one plans to ship samples only once or twice, consider becoming a coinvestigator on another researcher’s or institution’s permit. 2. Q: What permits does a researcher at an academic institution need to obtain marine mammal samples for a project? How are the samples obtained? A: An NMFS or FWS scientific research or enhancement permit is needed to collect, obtain, or import samples. Once obtained, the researcher or institution who possesses the samples can ship them to other researchers. To import or export marine mammal parts, one must have a CITES permit (see question 1 above). Within the United States, parts may also be loaned for the purpose of scientific research, education, or maintenance in a properly curated, professionally accredited scientific collection.
For NMFS-regulated species only: Section 216.37 in 50 CFR describes the procedure for transferring, importing, and exporting marine mammal parts. It is legal to ship (transfer, import, or export) marine mammal samples if: (a) the person transferring the part receives no remuneration for the part; (b) The recipient possesses a marine mammal scientific research or enhancement permit and any appropriate CITES permits to import or export samples; and (c) the part is transferred for the purpose of scientific research, education, or maintenance in a properly curated, professionally accredited scientific collection. 3. Q: If one wants to do some physiological studies on marine mammals, can animals that are being rehabilitated or that are on public display be used? A: Generally, it is preferable for an applicant to conduct physiological studies on rehabilitated or display animals as opposed to capturing or “taking” animals from the wild. To conduct scientific research on any marine mammal, one must apply for a scientific research permit from the appropriate agency (the NMFS or the FWS). In addition, any activities beyond normal, daily animal husbandry practices require a scientific research permit. Because the application process includes a mandatory 30-day public comment period, allow a minimum of 120 days for the entire process. All applications are sent to the MMC for review and comment. Call the NMFS or the FWS for application materials or visit their Web sites.
Public Display Permits 1. Q: While a marine mammal is being rehabilitated, can it be placed on public display? A:
A marine mammal can be placed on public display incidental to any scientific research or enhancement (e.g., rehabilitation) activities. An endangered species cannot be held in captivity solely for the purpose of public display. For further information, contact the FWS or the NMFS.
2. Q: What are the requirements for holding a marine mammal for public display? A:
According to Section 104(c)(2)(A) of the MMPA, an individual or facility may obtain a permit for public display if: (a) the facility offers a program for education or conservation purposes that is based on professionally recognized standards of the public display community; (b) the facility is registered or holds a license issued under 7 USC 2131 et seq. (USDA/APHIS license); and (c) the facility is open to the public on a regularly scheduled
0839_frame_C33.fm Page 765 Tuesday, May 22, 2001 1:38 PM
U.S. Federal Legislation Governing Marine Mammals
765
basis, and that access is not limited or restricted other than by charging of an admission fee. As with the scientific research and enhancement permits, one should allow a minimum of 120 days to process a public display permit and must contact either the NMFS or the FWS for application instructions.
Other Permits 1. Q: If one wants to conduct relatively benign research, such as behavioral observations and passive listening, is it still necessary to apply for a permit and wait 120 days for processing? A:
For nonendangered or nonthreatened species, the NMFS has devised a more streamlined way to obtain the authority to conduct benign research activities such as behavioral observations. This process applies only to species under the jurisdiction of the NMFS and does not apply to endangered or threatened species. As stated in the definitions section, the MMPA recognizes two forms of harassment, Level A and Level B. Level A harassment includes activities that have the potential to injure marine mammals, and Level B harassment is any activity that has the potential to disturb a marine mammal. To obtain the authority to conduct Level B harassment, you need to acquire an LOA under the general authorization process. One should allow 30 to 45 days for this process and should contact the NMFS for further information.
For all endangered or threatened species and all FWS-regulated species: One must apply for a scientific research or enhancement permit and allow 120 days for processing. Refer all questions to the FWS or the NMFS. 2. Q: A commercial film crew wants to document research. Do they need a permit? A: Yes. They have two options. They can apply for a commercial or educational photography permit, or the researcher can add the camera crew’s activities to his or her permit. If they conduct activities under the authority of the researcher’s permit, they would be prohibited from influencing the research activities in any manner. If they apply for a commercial or educational photography permit, they would be restricted to activities classified as Level B harassment and their final products must be made available to the public. If the research involves endangered species, it is recommended that the photography crew’s activities be added to the researcher permit. 3. Q: A new anesthesia has just been made available. If a researcher already has a permit to anesthetize animals in the course of research, can the permit be changed to include the use of this new drug? A: One would need to apply for a major amendment to the existing permit. There are two types of permit amendments: major and minor. According to 50 CFR 216.39, a major amendment is any change to the permit conditions regarding: (a) the number and species of marine mammals being “taken”; (b) the manner in which the animals may be “taken” (activities); (c) location of the research activities; and (d) permit duration (if longer than 1 year). Minor amendments include the addition of coinvestigators, extensions of less than 1 year, and changes in dates of the field season. It is necessary to allow at least 120 days to process major amendments because the process includes a mandatory 30-day public comment period and review by the MMC.
Acknowledgments The authors thank Leslie Dierauf, Marilee Menard, Kris Vehrs, Monica Farris, Charles Hamilton, Ruth Johnson, and Judity Leiby for their reviews of this chapter.
0839_frame_C33.fm Page 766 Tuesday, May 22, 2001 1:38 PM
766
CRC Handbook of Marine Mammal Medicine
References Animal Welfare Enforcement, 1985, Fiscal Year 1984, Animal and Plant Health Inspection Service, U.S. Department of Agriculture, Washington, D.C., May, 112 pp. CITES, 1995, Appendices I, II, and III to the Convention on International Trade in Endangered Species of Wild Fauna and Flora, February 16. Convention on International Trade in Endangered Species of Wild Fauna and Flora, 1987, 1985 Annual Report, U.S. Fish and Wildlife Service, U.S. Department of the Interior, Washington, D.C., May, 88 pp. Cooper, M.E, 1987, An Introduction to Animal Law, Academic Press, Orlando, FL, 230 pp. Endangered Species Act of 1973 (50 CFR 17.11 and 17.12). Lyster, S., 1985, International Wildlife Law, Grotius Publications Unlimited, Cambridge, U.K., 325 pp. Marine Mammal Commission, 1989, Annual Report of the Marine Mammal Commission, Calendar Year 1988, Marine Mammal Commission, Washington, D.C. Marine Mammal Commission, 1995, Annual Report of the Marine Mammal Commission, Calendar Year 1994, Marine Mammal Commission, Washington, D.C. Marine Mammal Commission, 1999, Annual Report of the Marine Mammal Commission, Calendar Year 1998, Marine Mammal Commission, Washington, D.C. Marine Mammal Protection Act of 1972, 1985, Administration January 1, 1984 to December 31, 1985, U.S. Department of the Interior, U.S. Fish and Wildlife Service, Washington, D.C., May, 87 pp. Marine Mammal Protection Act of 1972, 1986, Annual Report, 1985/6, U.S. Department of Commerce, National Marine Fisheries Service, National Oceanic and Atmospheric Administration, Washington, D.C., June, 102 pp. Marine Mammal Protection Act of 1972, 1995 (50 CFR 216.15), Compiled and annotated by the Marine Mammal Commission, Washington, D.C., February. Shoemaker, A.H., 1985, AAZPA Manual of Federal Wildlife Regulations, Protected Species, Vol. 1, American Association of Zoological Parks and Aquariums, Wheeling, WV, 173 pp. USFWS (U.S. Fish and Wildlife Service), 2000, Digest of Federal Resource Laws of Interest to the U.S. Fish and Wildlife Service, Endangered Species Act of 1973, available at http://laws.fws.gov/lawsdigest/index.html. Vehrs, K.L., 1985, AAZPA Manual of Federal Wildlife Regulations, Laws and Regulations, Vol. 2, American Association of Zoological Parks and Aquariums, Wheeling, WV, 97 pp.
0839_frame_C34 Page 767 Tuesday, May 22, 2001 1:44 PM
34 Public Health Daniel F. Cowan, Carol House, and James A. House
Introduction Reports of transmission of disease from marine mammals to humans are scarce; however, as humans are increasingly in contact with marine mammals, the possibility of encountering new diseases must be considered (Tryland, 2000). Lack of reports in the literature may indicate lack of occurrence of disease, but may also reflect lack of recognition by physicians or failure to report for a variety of reasons. Until recently, only hunters and scientists were likely to have close physical contact with marine mammals, and the public’s exposure was limited to zoos or aquaria with animals behind barriers. However, in the last decade, human contact with marine mammals has changed so that a broader range of people are potentially exposed to zoonoses. Sophisticated oceanaria have improved housing for marine mammals, and inner-city aquaria bring education opportunities and tourism dollars into urban restoration projects. The display animals in oceanaria are generally healthy, and their care is well regulated, for the benefit of both the animals and the humans they contact. However, they may carry infections that are not apparent as clinical disease. Many facilities now feature interactive programs, allowing the public to feed and/or pet the trained performers, thus increasing the degree of contact between marine mammal and humans. Increasing public interest in marine mammals has resulted in the emergence of popular “swim-with-the-dolphin or manatee” programs (see review by Samuels et al., 2000). These programs offer tourists and, in some cases, handicapped children, the opportunity to encounter free-ranging dolphins or manatees, or those in large, ocean-connected enclosures. The wild dolphins are presumed healthy, but are not usually subject to routine veterinary care. Appealing as they might be, wild animals are indeed wild, and the possibility of injury and disease transmission is always present. Also, the dolphins, while healthy themselves, may harbor organisms as part of their normal flora that may be potentially pathogenic to humans. Another recent development is the formation of centers and networks dedicated to the collection and rehabilitation of stranded marine mammals. Wild animals are often infected with parasites and other potential disease agents, including viruses, bacteria, protozoa, and fungi. Workers in programs for the rescue and rehabilitation of stranded marine mammals are exposed to animals that must be presumed sick. Most rescue programs rely heavily on volunteers who donate a few hours a week, so the number of individuals exposed to stranded animals is considerable. The training volunteers receive varies from facility to facility, and avoiding injury and exposure of the volunteers is variably successful. Handlers supporting sick animals in small pools are exposed to water that may quickly become contaminated with urine
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
767
0839_frame_C34 Page 768 Tuesday, May 22, 2001 1:44 PM
768
CRC Handbook of Marine Mammal Medicine
and feces. However, perhaps the most likely situation for transmission of disease from marine mammals to humans is during the post-mortem examinations of large whales. During such operations, humans may be literally immersed in a marine mammal, exposing mucous membranes and cut surfaces to a variety of potential pathogens. These increasing contacts between humans and marine mammals raise significant public health issues, as it is not just theoretically possible for humans to contract infection from marine mammals. For example, in one survey of a sealing fleet, 10% of the crew was affected by seal finger (Rodahl, 1953). Handlers of marine mammals may be at some risk of exposure to parasite ova from fish fed to marine mammals, or from ova in the stool of the mammals (Meyers, 1970). Direct transmission of metazoan parasites from marine mammals to humans is probably not a serious consideration, because of the complex life cycles of parasites, but direct or indirect contact with viruses, bacteria, protozoa, and fungi may result in human infection. This chapter addresses known pathogens associated with marine mammals, certain infectious diseases of marine mammals, and human diseases that are known to be, or may be, caused by transmissible agents from marine mammals. Although at this time, with a few exceptions, transmission may seem to be hypothetical, awareness of risk and taking of appropriate precautions may prevent the hypothetical from becoming the actual.
Viral Infections Poxviruses Humans may acquire parapox virus infections through contact with seals with seal pox (see Chapter 15, Viral Diseases). The first clinical symptom is observed 10 to 20 days postexposure. A red area (a macule) appears and typically persists for 24 hours, progressing into a papule with a raised pale center, reflecting edema. This may further progress to a vesicular stage. The delicate vesicles often rupture, particularly when they occur in areas subject to abrasion. After a few days, the papule or vesicle becomes a pustule as leukocytes accumulate in the lesion. The pustule dries up over a period of 1 to 5 weeks and an eschar scab is formed. However, lesions may persist for several months or as long as a year before healing is complete (Hicks and Worthy, 1987). The lesions and scabs contain infectious virus and are the source of new infections of animals and humans. Even dried scabs may contain viable virus for months. Seal pox virus has been isolated from both the host animals and their human contacts (Hicks and Worthy, 1987). Stranded seals, which are often stressed, poorly nourished, and heavily parasitized, frequently exhibit clinical seal pox. Seal pox infections can spread within a stranding center with overcrowded conditions, as sanitary and quarantine measures are difficult to achieve (see Chapter 41, Seals and Sea Lions). Volunteers rescuing and treating stranded seals should wear gloves whenever handling animals. Although human-to-human transmission of seal pox has not been reported, care should be taken to prevent exposure. There is no treatment for seal pox infection other than topical supportive care to prevent complications.
Calicivirus There is a single report of a laboratory worker who showed a deep skin lesion after working with a marine calicivirus, San Miguel sea lion virus (SMSV) serotype five (Smith et al., 1978a,b). Co-workers developed rising titers to additional serotypes of this group of viruses. SMSVs are marine caliciviruses common in both fish and marine mammals in the Pacific Ocean (Smith et al., 1983) (see Chapter 15, Viral Diseases). These viruses, of which there are 18 or more
0839_frame_C34 Page 769 Tuesday, May 22, 2001 1:44 PM
Public Health
769
serotypes, cause vesicular lesions in the mouths and on the flippers of California sea lions (Zalophus californianus). The first isolations were reported from California sea lions on San Miguel Island in 1973 (Smith et al., 1973). SMSVs have been shown to cause vesicular exanthema of swine (VES), a disease first recognized in the United States in 1932. A recent isolate from California sea lions was shown experimentally to infect swine and cause a disease indistinguishable from VES (Van Bonn et al., 2000). VES was eradicated in the United States in the late 1950s after requirements for cooking garbage fed to swine were implemented. It should be noted that during the national eradication program, there were no reports of human infection, even though tens of thousands of VES-infected pigs were slaughtered. In addition, Alaskan natives handling SMSV-infected seals did not develop disease or antibody titers after harvesting infected seals (Smith et al., 1978a). The lack of disease and antibody in people who were heavily exposed places this group of viruses low on the scale of probable zoonotic agents.
Influenza Both humans and marine mammals are susceptible to infection with avian-origin influenza viruses. Four people developed conjunctivitis after post-mortem examination of infected seals, and one person developed conjunctivitis caused by influenza A/seal/Mass/1/80(H7N7) after a harbor seal (Phoca vitulina) sneezed on him (Webster et al., 1981a,b). A reciprocal case, human sneezing on seal, is theoretically possible. Recently, influenza B was isolated from harbor seals in the Netherlands, several years after a human epidemic (Osterhaus et al., 2000) (see Chapter 15, Viral Diseases).
Rabies Rabies virus infects many animals, including humans, with high mortality in unvaccinated individuals. The first and only documented case in a marine mammal occurred in a ringed seal (Phoca hispida) in Norway in 1981 (Odegaard and Krogsrud, 1981). Although rare, any animal demonstrating neurological signs should be treated with appropriate precautions. Morbilliviruses, which cause the distempers, also produce neurological signs in seals, dolphins, porpoises, and a wide range of other cetaceans, but these marine morbilliviruses have not been shown to be infecious to humans, although some other morbilliviruses are.
Bacterial Infections Many bacterial species are found in marine waters and may be recovered from marine mammals, either as pathogens or as part of a complex normal flora. Bacterial infection is thought to be the main cause of disease and death in marine mammals, especially in captivity (Howard et al., 1983). The vast majority of bacteria associated with marine mammals are not of public health concern (see Chapter 16, Bacterial Diseases). However, a few are known pathogens of humans and some could be adventitious infectious agents for persons with compromised immune systems, or may be inoculated into bites, cuts, or abrasions.
Vibrio spp. Marine organisms known to produce severe or fatal infections in humans include the halophilic Vibrio spp. (V. fulnificus, V. parahemolyticus, V. damsela, V. cholerae, V. fluvialis, V. pelagius, V. furnissi, V. alginolyticus, V. metchnikovii, V. gazogenes, V. mimicus, V. hollisae) (Fujioka et al., 1988; Buck and Schroeder, 1990; Woods and Guiterrez, 1993), and Edwardsiella tarda (Janda and
0839_frame_C34 Page 770 Tuesday, May 22, 2001 1:44 PM
770
CRC Handbook of Marine Mammal Medicine
Abbott, 1993). These organisms are common in the marine environment and are frequently encountered in cetaceans (Howard et al., 1983; Cowan et al., 1998), but are less often encountered in pinnipeds (Johnson et al., 1998; Thornton et al., 1998). Vibrio infections are frequently mixed. Three clinical syndromes in human infection have been identified: gastroenteritis, wound infection, and primary septicemia. In one survey of human cases in Florida, gastroenteritis accounted for about half of all cases, wound infection about a quarter, and about 17% were primary septicemia. Wound infection and septicemia are highly seasonal, April through October (at least in Florida), relating to the seasonal abundance of V. fulnificus and V. parahemolyticus. Wound infections are largely occupational, occurring in people who fish and others working around the water, whereas 68% of cases of gastroenteritis and 83% of cases of septicemia were associated with ingestion of raw oysters. Septicemia is clearly associated with preexisting conditions (Hlady and Klontz, 1996). Middle ear infections, especially with V. alginolyticus and V. parahemolyticus, may be acquired from exposure to contaminated seawater, especially if the patient has a perforation of the tympanic membrane (Hornstrup and Gahrn-Hansen, 1993).
Edwardsiella spp. Only one of the three species of Edwardsiella, E. tarda, is known to be pathogenic for humans, causing gastroenteritis, wound infection (cellulitis and gas gangrene), septicemia, meningitis, cholecystitis, and osteomyelitis. Infection is usually attributed to exposure to the aquatic environment, exotic animals, including marine mammals, reptiles, and amphibia, and eating raw fish. As with Vibrio infection, preexisting liver disease, iron overload, and immune impairment predispose to infection (Janda and Abbott, 1993).
Clostridium spp. The Clostridium spp. are spore-forming obligate anaerobic bacilli, ubiquitous in the environment in soil, sewage, marine sediments, decaying animals and plant products, and the intestinal tracts of many animals. Although more than 80 species are known, those implicated in human disease include C. botulinum, C. tetani, C. perfringens, C. difficile, C. sordelli, C. novyi, C. histolyticum, C. septicum, C. bifermentans, C. sporogenes, C. tertium, C. ramosum, C. butyricum, and C. baratyii (Woods and Guiterrez, 1993). In human medicine, most disease is toxic or enterotoxic, related to C. botulinum and C. difficile; however, in the context of this discussion, the main risk would seem to be wound infection. Many species of Clostridium have been cultured from blood, lesions, and intestinal tract of stranded bottlenose dolphins (Tursiops truncatus) in the Gulf of Mexico (Cowan et al., 1998) but appear less common in California pinnipeds (Thornton et al., 1998).
Leptospira Leptospirosis occurs in harbor seals (Stamper et al., 1998; Stevens et al., 1999), California sea lions, and northern fur seals (Callorhinus ursinus) (Gulland, 1998). It should be noted that veterinarians and veterinary technicians have been made ill through contact with fluids and tissues during necropsy of sea lions infected with L. interrogans pomona (Smith et al., 1978b).
Streptococcus β-Hemolytic streptococci (Lancefield group L) have been identified as playing an important role in infections of harbor porpoise (Phocoena phocoena) of the North and Baltic Seas (Swenshon et al., 1998). While Lancefield group L are well known as pathogens of a variety of animals, causing mastitis and various infections, they have only rarely been recognized as causing
0839_frame_C34 Page 771 Tuesday, May 22, 2001 1:44 PM
Public Health
771
infections of humans. These infections may include bacteremia and endocarditis in debilitated persons (Ellner, 1970; Bevanger and Stamnes, 1979) and cellulitis, wound infections, impetigo, and paronychia in meat handlers (Barnham and Neilson, 1987).
Brucella Brucella infection of the placenta with abortion has been reported in bottlenose dolphins. The organism, B. delphini, appears to be readily transmissible among dolphins, and has also been cultured from the lung of a bottlenose dolphin at necropsy (Miller et al., 1999). Brucella infection occurs in other cetaceans and seals (Ross et al., 1996). A substantial percentage of marine mammal serum samples (about 30%) react positively on tests used to detect antibody to Brucella spp., and a number of Brucella isolates have been obtained from marine mammals. However, only B. delphini has been associated with reproductive failure in marine mammals. One laboratory worker in the United Kingdom became ill when handling a marine Brucella isolate, and responded positively to a 6-week course of rifampin and doxycycline (Brew et al., 1999).
Erysipelothrix rhusiopathiae Red indurated patches on the skin of marine mammals or humans may indicate infection with E. rhusiopathiae/insidiosa. In humans, the disease is generally localized and is called erysipeloid. (Do not be confused by the term erysipelas in the human literature, which is a superficial cellulitis caused by Group A β-hemolytic streptococci.) In classic cases of disease caused by E. rhusiopathiae, a diamond pattern on the skin may be observed. Typical clinical signs of erysipelas are swelling and pain, but more generalized illness (polyarthritis, septicemia, or pneumonia) is also recognized (Medway, 1980; Woods and Gutierrez, 1993). Erysipelothrix rhusiopathiae was once thought to be the cause of seal finger, but new evidence and antibioticresistance patterns implicate Mycoplasma as the probable cause. The Gram-positive or Gramvariable organism E. rhusiopathiae is readily isolated from fish, so the source of infection for persons working with marine mammals may be fresh or frozen fish used for feed. Broadspectrum antibiotics are generally effective in treating the disease.
Mycobacterium spp. Mycobacterium marinum (syn. M. platypoecilus, M. balnei), originally described from fish, was first recognized as a human pathogen in 1951 (Norden and Linell, 1951). Mycobacterium marinum has been reported to have been transmitted to a handler by a dolphin bite (Flowers, 1970). The handler was bitten on a finger during a training session. About 2 months later, firm fluctuant swellings appeared in the vicinity of the original wound. Viscid pus was aspirated from one lesion. Cultures taken then and a month later yielded pure growth of M. marinum. The lesions healed over several months. Infections with M. marinum are uncommon, but are described in the literature (Woods and Gutierrez, 1993). They tend to heal spontaneously but may take up to 2 years to do so. Most inoculations are on the elbow, knee, foot, toe, or finger, and lesions may be verrucose or ulcerated. Extracutaneous manifestations are rare, and include synovitis, osteomyelitis, and ocular and laryngeal lesions; in immunocompromised persons, the disease may become disseminated (Woods and Gutierrez, 1993). Mycobacterium bovis has been reported to cause pulmonary tuberculosis in a seal trainer. Cultural characteristics, biochemical reactions, sodium dodecyl sulfate polyacrilamide gel electrophoresis, and restriction endonuclease analysis linked his infection to three seals with which he had worked 2 years earlier. None of the seals was overtly sick despite post-mortem findings showing
0839_frame_C34 Page 772 Tuesday, May 22, 2001 1:44 PM
772
CRC Handbook of Marine Mammal Medicine
extensive tissue involvement (Thompson et al., 1993). Mycobacterium tuberculosis was diagnosed in a captive colony of mixed pinnipeds by pathology, culture, and tuberculin skin testing (Forshaw and Phelps, 1991). It was also diagnosed in wild pinnipeds that stranded off Australia and Argentina (Romano et al., 1995; Woods et al., 1995). An instance of disseminated M. chelonei infection in a manatee (Trichechus inunguis) has also been reported (Boever et al., 1976). Howard et al. (1983) cite cases of cutaneous mycobacteriosis in a manatee and its handler, attributed to M. chelonei.
Coxiella burnetii There is an interesting report that the causative agent of Q fever (C. burnetii) was identified in the placenta of a Pacific harbor seal (Phoca vitulina richardsi) (Lapointe et al., 1999). The animal was euthanatized because of protozoal encephalitis, and the intracellular Gram-negative bacteria were found during a histopathological study of the tissues collected at necropsy.
Other Mixed Infections The most common organisms identified in two surveys of bacterial isolates from collection and stranded marine mammals, apart from coliforms, were Clostridium (eight species, 21 isolates), Vibrio (eight species, 51 isolates), Citrobacter (two species, ten isolates), and Edwardsiella tarda (five isolates) (Howard et al., 1983; Cowan et al., 1998). The majority of these species are known to be pathogenic or potentially pathogenic for humans. In another study, 21 different species of bacteria were cultured from dolphins living in ocean pens in southern California (Johnson and Fung, 1969). These included six species of Pseudomonas, two Proteus, three Streptococcus, two Staphylococcus, and Edwardsiella tarda, among others. This is in contrast to experience with pinnipeds from the California coast, in which the major pathogens encountered in sea lions, elephant seals (Mirounga angustirostris), and harbor seals are enteric organisms, mainly Escherichia coli, Klebsiella pneumoniae, K. oxytoca, Proteus spp., Pseudomonas spp., and Enterococcus spp., as well as nine different serotypes of Salmonella (most commonly S. newport), β-hemolytic streptococci, and Staphylococcus aureus. Infection with Vibrio spp. and Clostridium spp. was less common, while infection with Listeria ivanovii, not recognized on the Gulf Coast, was especially prevalent in lesions from harbor seals (Johnson et al., 1998; Thornton et al., 1998). Other bacterial infections reported in marine mammals are Pseudomonas mallei (many species), P. pseudomallei [many species, notably an epizootic of melioidosis of dolphins, pilot whales (Globicephala spp.), and harbor seals in Hong Kong], Pasturella multocida (California sea lion), Neisseria spp. (dolphins), Nocardia asteroides (pilot whale), N. brasiliensis (bottlenose dolphin), N. caviae (bottlenose dolphin), N. paraguayensis (dolphin) (Medway, 1980), and actinomycosis (Sweeney et al., 1976). However, none of these has as yet been associated with disease of humans as a result of transmission from a marine mammal.
Mycoplasma Infections The most common serious ailment associated with bites from seals or sea lions is seal finger, also called spaeck finger, sealer’s finger, speck finger, or blubber finger (Candolin, 1953). A mycoplasma, M. phocacerebrale, was isolated in 1990 from both the front teeth of a healthy seal and the finger lesion of a woman it had bitten at the New England Aquarium in Boston, Massachusetts (Madoff et al., 1991; Stadtlander and Madoff, 1994). The organism was originally isolated from diseased seals in 1988 from the North Sea and the Baltic Sea (Giebel et al., 1991; Baker et al., 1998). Koch’s postulates are not satisfied at this time, so there is only circumstantial evidence that seal finger is caused by one or more mycoplasmas that may be present in both healthy and sick seals.
0839_frame_C34 Page 773 Tuesday, May 22, 2001 1:44 PM
Public Health
773
In one survey in 1950, over 10% of a Norwegian sealing fleet was affected with seal finger (Rodahl, 1953). Usually within 1 to 8 days, but up to 21 days later, the area around the puncture wound becomes markedly swollen and extremely painful, with the joint nearest the wound becoming inflamed and immobile. The skin may become dark and discolored, but there is less erythema than in erysipeloid (Mass et al., 1981). Recurring and worsening symptoms, regional lymphadenitis and lymphadenopathy (Sargent, 1980), demineralization of the bone, and loss of mobility and permanent disability (Mass et al., 1981) may result without effective treatment. Seal finger may also be transmitted by a cut from a contaminated knife, or from infection of previously existing open wounds. There is no evidence of transmission to humans from frozen infected seal tissues. Penicillin is not effective in treating seal finger. Tetracycline, 150 to 500 mg four times per day for as long as 4 to 6 weeks (Mass et al., 1981), or doxycycline at 200 mg/adult loading dose, followed by 100 mg/day (Gulland, pers. comm.) is recommended.
Fungal Infections A large number of fungal species have been recovered from marine mammals (see Chapter 17, Mycotic Diseases). Opportunistic fungi identified in marine mammals include Candida albicans, other Candida spp., Aspergillus fumigatus, Cryptococcus neoformans, Cladophialophora bantiana, Apophysomyces elegans, Saksenaea vasiformis, Mucor, Rhizopus, and other Zygomycetes, and Fusarium spp. Primary (endemic) pathogens include Blastomyces dermatiditis, Coccidioides immitis, Histoplasma capsulatum, and Lacazia loboi (Medway, 1980; Migaki and Jones, 1983; Cowan et al., 1998; Jensen et al., 1998; Reidarson et al., 1999; Wunschmann et al., 1999; Haubold et al., 2000). Animals with immune suppression from morbillivirus disease may suffer severe, disseminated fungal infection, frequently from Aspergillus spp., as a terminal event. An unusual, perhaps unique case of disseminated sporotrichosis (Sporothrix shenckii) has been reported in a Pacific white-sided dolphin (Lagenorhynchus obliquidens) (Migaki et al., 1978). Dermatophytosis caused by Epidermophyton floccosum has been reported in manatees (Dilbone, 1965) and Microsporum canis has been recovered from scaling, pustular lesions in a harbor seal (Farnsworth et al., 1975). Since infection with fungi requires spores from the environment, rather than the vegetative stages found in marine mammals, direct transmission to humans from animals seems unlikely. To date, only one instance of direct association of transmission of a fungus infection from a dolphin to a human has been reported. This is a case of Lobo’s disease transmitted from a captive bottlenose dolphin to a handler (Symmers, 1983). Lobo’s disease (once called keloidal blastomycosis) is a skin infection producing chronic, treatment-resistant, thick nodular swellings of the superficial dermis and epidermis, occasionally with ulceration. It is caused by a fungus, Loboa loboi (syn. Lacazia loboi). Although the lesions in humans and dolphins are quite similar, there are subtle morphological differences in the organisms in the lesions (Haubold et al., 2000). In humans, Lobo’s disease is a disease of the Central and South American tropics, and in dolphins it ranges from the Gulf of Mexico, mainly Florida, to South America.
Protozoal Infections Toxoplasma gondii Toxoplasmosis (infection with T. gondii), has been reported in a variety of marine mammals, including a West Indian manatee (Buergelt and Bonde, 1983), several pinnipeds (Migaki et al., 1977; Holshuh et al., 1985), stranded Atlantic bottlenose dolphins (Inskeep et al., 1990),
0839_frame_C34 Page 774 Tuesday, May 22, 2001 1:44 PM
774
CRC Handbook of Marine Mammal Medicine
Pacific spinner dolphins (Stenella longirostris) (Dubey and Beattie, 1988; Migaki et al., 1990), belugas (Delphinapterus leucas) (Mikaelian et al., 2000), and a long-time captive killer whale (Orcinus orca) (Cowan, unpubl. data). The animals were found in both the Atlantic and Pacific Oceans, were in collections and free-ranging, and the lesions ranged from incidental to disseminated and fatal. The mode of transmission in these animals is not known, and while humans are certainly susceptible to toxoplasmosis, there are no reports of transmission to humans from a marine mammal.
Cryptosporidium spp. Cryptosporidium oocysts morphologically, immunologically, and genetically indistinguishable from C. parvum and C. duodenalis obtained from infected domestic cattle have been recovered from feces of California sea lions, suggesting that the sea lion could serve as a reservoir for environmental transmission of this organism (Deng et al., 2000).
Giardia spp. Giardia spp. cysts have been isolated from fecal material from harp seals (Pagophilus groenlandicus), gray seals (Halichoerus grypus), and harbor seals in eastern Canadian waters (Measures and Olson, 1999), from ringed seals in western Arctic Canada (Olson et al., 1997), and from California sea lions in northern coastal California (Deng et al., 2000). The transmission of these organisms between marine mammals and humans has not been demonstrated, and little is known about the strain types found in marine mammals. However, their potential presence in feces should be remembered when handling marine mammals.
Potential for Transmission of Infectious Disease from Marine Mammals to Humans Several observations emerge from personal experience and review of the literature. One is that wild animals may be very sick with infectious disease and yet show remarkably few signs of disease until shortly before death. The implication is that reasonable precautions against transmission of infection must be taken around all marine mammals, whether they appear to be healthy or not. Face protection should be worn whenever an animal’s breath can be exhaled into the face of a handler. Food handlers and tank cleaners should take full precautions. Persons with open cuts or abrasions should not be permitted exposure to blood or secretions from marine mammals, or the water in which they live. Pregnant women and people with chronic illness, especially liver disease, or any form of immune suppression, should not be permitted close exposure to marine mammals. Gloves and other protective gear should be worn during post-mortem examination. Review of the possibilities of infection, except for a few virus infections, indicates that documented instances of infection are rare, except in the case of inoculation, as in seal finger. Although uncommon, infection of a marine mammal with a species of Mycobacterium may have a relatively high risk of transmission to a handler. These infected animals may be very difficult to recognize clinically, before culture. In 10 years of contact with many dolphin necropsies and with rehabilitating live strandings with the Texas Marine Mammal Stranding Network, the authors have not recognized a single instance of transmission of infection from dolphin to human. This might relate to the general good health and high resistance of people working with these animals, but might also be related to serotypes of the potential bacterial pathogens. Identification by culture alone does not mean that the organism is an agent of great risk under
0839_frame_C34 Page 775 Tuesday, May 22, 2001 1:44 PM
Public Health
775
ordinary circumstances. Streitfield and Chapman (1976) examined dolphins and healthy human attendants from two oceanaria by bacteriological culture. Although coagulase-positive Staphylococcus spp. were recovered from blowholes of eight dolphins (one with respiratory tract infection) and from 14 people, antibiotic sensitivity and phage typing indicated that the Staphylococcus spp. found on dolphins and humans were not shared. Types were consistent within the dolphins, and within the humans, but did not occur across the species. Infection from a marine mammal source may begin subtly, without preceding incident, or following an encounter such as a bite or scratch. When seeking treatment for any possibly infectious condition, the history of contact with a marine mammal should be presented, since it is not something that the average physician would think to ask. As a general practice, a full clinical history of the bite victim and the biting animal should be taken. If the biting animal was showing neurological symptoms, the possibility of rabies should be the physician’s concern. The animal can be euthanatized, and brain tissues can be submitted to a public health laboratory certified for rabies diagnosis. The submission information should clearly state “A Rabies Suspect.” The bite victim’s medical history will allow the physician to determine if a tetanus booster is advisable. If the history indicates that the patient is immunologically compromised, a wider range of microorganisms should be considered as potentially pathogenic in that individual.
Acknowledgments The authors thank Bill Van Bonn, Teri Rowles, Jan Kovach, and Jim Hurley for their helpful comments in reviewing this chapter.
References Baker, A.S., Ruoff, K.L., and Madoff, S., 1998, Isolation of Mycoplasma species from a patient with seal finger, Clin. Infect. Dis., 27: 1168–1170. Barnham, M., and Neilson, D.J., 1987, Group L beta-hemolytic streptococcal infection in meat handlers: Another streptococcal zoonosis? Epidemiol. Infect., 99: 257–264. Bevanger, L., and Stamnes, T.L., 1979, Group L streptococci as the cause of bacteremia and endocarditis, a case report, Acta Pathol. Microbiol. Scand. B, 87: 301–302. Boever, W.J., Thoen, C.O., and Wallach, J.D., 1976, Mycobacterium chelonei infection in a Natterer manatee, J. Am. Vet. Med. Assoc., 169: 927–929. Brew, S.D., Perrett, L.L., Stack, J.A., and MacMilllan, A.P., 1999, Human exposure to Brucella recovered from a sea mammal, Vet. Rec., 144: 483. Buck, C.D., and Schroeder, J.P., 1990, Public health significance of marine mammal disease, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 163–173. Buergelt, C.D., and Bonde, R.K., 1983, Toxoplasmic meningoencephalitis in a West Indian manatee, J. Am. Vet. Med. Assoc., 183: 1294–1296. Candolin, Y., 1953, Seal finger and its occurrence in the gulfs of the Baltic Sea, Acta Chir. Scand., 177(Suppl.):62. Cowan, D.F., Turnbull, B.S., and Haubold, E.M., 1998, Organisms cultured from stranded cetaceans: Implications for rehabilitation and for safety of handlers, in Proceedings of the 29th International Association for Aquatic Animal Medicine, San Diego, CA, 148–149. Cusak, P.K., and Bullock, B., 1973, Ulcerative dermatitis and pneumonia associated with Aeromonas hydrophila infection in the bottle-nosed dolphin, J. Am. Vet. Med. Assoc., 163: 578–579. Deng, M.Q, Peterson R.P., and Cliver, D.O., 2000, First findings of Cryptosporidium and Giardia in California sea lions (Zalophus californianus), J. Parasitol., 86: 490–494. Dilbone, R.P., 1965, Mycosis in a manatee, J. Am. Vet. Med. Assoc.,147: 1095–1096.
0839_frame_C34 Page 776 Tuesday, May 22, 2001 1:44 PM
776
CRC Handbook of Marine Mammal Medicine
Dubey, J.P., and Beattie, C.P., 1988, Toxoplasmosis of Animals and Man, CRC Press, Boca Raton, FL, 220 pp. Ellner, P.D., 1970, Endocarditis due to groups L. Streptococcus, Ann. Intern. Med., 72: 547–548. Farnsworth, R.J., McKeever, P.J., and Fletcher, J.A., 1975, Dermatomycosis in a harbor seal caused by Microsporum canis, J. Zoo Anim. Med., 8: 26–28. Flowers, D.J., 1970, Human infection due to Mycobacterium marinum after a dolphin bite, J. Clin. Pathol., 23: 475–477. Forshaw, D., and Phelps, G.R., 1991, Tuberculosis in a captive colony of pinnipeds, J. Wildl. Dis., 27: 288–295. Fujioka, R.S., Greco, S.B., Cates, M.B., and Schroeder, J.P., 1988, Vibrio damsela from wounds in bottlenose dolphins, Tursiops truncatus, Dis. Aquat. Organ, 4: 1–4. Giebel, J., Meier, J., Binder, J., Flossdorf, J., Poveda, J.B., Schmidt, R., and Kirchhoff, H., 1991, Mycoplasma phocarhinis sp. nov. and Mycoplasma phocacerebrale sp. nov., two new species from harbor seals (Phoca vitulina L.), Int. J. Syst. Bacteriol., 41: 39–44. Gulland, F.M.D., 1998, Leptospirosis in marine mammals, in Zoo and Wild Animal Medicine, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 469–471. Haubold, E.M., Cooper, C.R., Jr., Wen, J.W., McGinnis, M.R., and Cowan, D.F., 2000, Comparative morphology of Lacazia loboi (syn Loboa loboi) in dolphins and humans, Med. Mycol., 38: 9–14. Hicks, B.D., and Worthy, G.A., 1987, Sealpox in captive gray seals (Halichoerus grypus) and their handlers, J. Wildl. Dis., 23: 1–6. Hlady, W.G., and Klontz, K.C., 1996, The epidemiology of Vibrio infections in Florida, 1981–1993, J. Infect. Dis., 173: 1176–1183. Holshuh, H.J., Sherrod, A.E., Taylor, C.R., Andrews, B.F., and Howard, E.B., 1985, Toxoplasmosis in a feral northern fur seal, J. Am. Vet. Med. Assoc., 187: 1229–1230. Hornstrup, M.K., and Gahrn-Hansen, B., 1993, Extra intestinal infections caused by Vibrio parahemolyticus and Vibrio alginolyticus in a Danish county, 1987–1992, Scand. J. Infect. Dis., 25: 735–740. Howard, E.B., Britt, J.O., Jr., Matsumoto, G.K., Itahara, R., and Nagano, C.N., 1983, Bacterial diseases, in Pathobiology of Marine Mammal Diseases, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 69–118. Inskeep, W., Gardiner, C.H., Harris, R.K., Goldston, R.T., and Dubey, J.P., 1990, Toxoplasmosis in Atlantic bottle-nosed dolphins, J. Wildl. Dis., 26: 377–382. Janda, J.M., and Abbott, S.L., 1993, Infections associated with the genus Edwardsiella: The role of Edwardsiella tarda in human disease, Clin. Infect. Dis., 17: 742–748. Jensen, E.D., Van Bonn, W., Lipscomb, T., and Ridgway, S.H., 1998, Disseminated histoplasmosis in Atlantic bottlenose dolphins, in Proceedings of the 29th Annual International Association for Aquatic Animal Medicine, San Diego, CA, 64. Johnson, D.G., and Fung, J., 1969, Bacterial flora of wild and captive porpoises, J. Occup. Med., 11: 276–277. 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. Kennedy-Stoskopf, S., Stoskopf, M.K., Eckhaus, M.A., and Strandberg, J.D., 1986, Isolation of a retrovirus and a herpesvirus from a captive California sea lion, J. Wildl. Dis., 22: 156–164. Lapointe, J.-M., Gulland, F.M., Haines, D.M., Barr, B.C., and Duignan, P.J., 1999, Placentitis due to Coxiella burnetii in a Pacific harbor seal (Phoca vitulina richardsi), J. Vet. Diagn. Invest., 11: 541–543. Madoff, S., Ruoff, K., and Baker, A.S., 1991, Isolation of a Mycoplasma species from a case of seal finger, presented at 91st General Meeting of the American Society for Microbiology, Poster G12. Mass, D.P., Newmeyer, W.L., and Kilgore, E.S., 1981, Seal finger, J. Hand Surg., 6: 610–612. Measures, L.N., and Olson M., 1999, Giardiasis in pinnipeds from eastern Canada, J. Wildl. Dis., 35: 779–782. Medway, W., 1980, Some bacterial and mycotic diseases of marine mammals, J. Am. Vet. Med. Assoc., 177: 831–834. Meyers, B.J., 1970, Nematodes transmitted to man by fish and aquatic mammals, J. Wildl. Dis., 6: 266–271. Migaki, G., and Jones J.R., 1983, Mycotic diseases in marine mammals, in Pathobiology of Marine Mammal Diseases, Howard, E.B. (Ed.), CRC Press, Boca Raton, FL, 1–27.
0839_frame_C34 Page 777 Tuesday, May 22, 2001 1:44 PM
Public Health
777
Migaki, G., Allen, J.F., and Casey, H.W., 1977, Toxoplasmosis in a California sea lion (Zalophus californianus), Am. J. Vet. Res., 38: 135–136. Migaki, G., Font, R.L., Kaplan, W., and Asper, E.D., 1978, Sporotrichosis in a Pacific white-sided dolphin (Lagenorhynchus obliquidens), Am. J. Vet. Res., 39: 1916–1919. Migaki, G., Sawa, T.R., and Dubey, J.P., 1990, Fatal disseminated toxoplasmosis in a spinner dolphin (Stenella longirostris), Vet. Pathol., 27: 463–464. Mikaelian, I., Boisclair, J., Dubey, J.P., Kennedy, S., and Martineau, D., 2000, Toxoplasmosis in beluga whales from the St. Lawrence estuary: Two case reports and a serological survey, J. Comp. Pathol., 122: 73–76. Miller, W.G., Adams, G.L., Ficht, T.A., Cheville, N.F., Payeur, J.P., Harley, D.R., House, C., and Ridgway, S.O., 1999, Brucella induced abortions and infection in bottlenose dolphins Tursiops truncatus, J. Zoo Wildl. Med., 30: 100–110. Neese, A., Beckmen, K.B., Lowenstine, L.J., and Jang, S., 1993, Listeria ivanovii infection in five phocid pups, Abstract, Proceedings, 10th Biennial Conference on the Biology of Marine Mammals, November 11–15, 193: 81. Norden, A., and Linell, F., 1951, A new type of pathogenic Mycobacterium, Nature, 168: 826. Odegaard, O.A., and Krogsrud, J., 1981, Rabies in Svalbard: Infection diagnosed in arctic fox, reindeer, and seal, Vet. Rec., 109: 141–142. Olson, M.E., Roach, 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., Yang, H., Spijkers, H.E.M., Groen, J., Teppema, J.S., and van Steenis, G., 1985, The isolation and partial characterization of a highly pathogenic herpesvirus from the harbor seal, Arch. Virol., 86: 239–251. Osterhaus, A.D.M.E., Rimmelzwaan, G.F., Martina, B.E.E., Bestebroer, T.M., and Fouchier, R.A.M., 2000, Influenza B virus in seals, Science, 288: 1051–1053. Reidarson, T.H., McBain, J.F., Dalton, L.M., and Rinaldi, M.G., 1999, Diagnosis and treatment of fungal infections in marine mammals, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 478–485. Rodahl, K., 1953, Speck finger, a severe finger infection observed in arctic sealers, Wes. J. Surg., 61: 39–43. Romano, M.I., Alito, A., Bigi, F., Fisanotti, J.C., and Cataldi, A., 1995, Genetic characterization of mycobacteria from South American wild seals, Vet. Microbiol., 47: 89–98. Ross, H.M., Jahans, K.L., MacMillan, A.P., Reid, R.J., Thompson, P.M., and Foster, G., 1996, Brucella species infection in North Sea seal and cetacean populations, Vet. Rec., 138: 647–648. Samuels, A., Bejder, L., and Heinrich, S., 2000, A review of the literature pertaining to swimming with wild dolphins, Marine Mammal Commission, Contract No. T74463123, Bethesda, MD, 57 pp. Sargent, E., 1980, Tetracycline for seal finger, J. Am. Med. Assoc., 244: 437. Smith, A.W., Akers, T.G., Madin, S.O., and Vedros, N.A., 1973, San Miguel sea lion virus isolation, preliminary characterization and relationship to vesicular exanthema of swine, Nature, 244: 108–110. Smith, A.W., Prato, C., and Skilling, D.E., 1978a, Caliciviruses infecting monkeys and possibly man, Am. J. Vet. Res., 39: 287–289. Smith, A.W., Vedros, N.A., Akers, T.G., and Gilmartin, W.G., 1978b, Hazards of disease transfer from marine mammals to mammals: Review and recent findings, J. Am. Vet. Med. Assoc., 173: 1131–1133. Smith, A.W., Skilling, D.E., and Ridgway, S., 1983, Calicivirus B induced vesicular disease in cetaceans and probable interspecies transmission, J. Am. Vet. Med. Assoc., 183: 1223–1225. Stadtlander, C.T.K.-H., and Madoff, S., 1994, Characterization of cytopathogenicity of aquarium seal mycoplasmas and seal finger mycoplasmas by light and scanning electron microscopy, Int. J. Med. Microbiol., Virol., Parasitol. Infect. Dis. (Germany), 280: 458–467. Stamper, M.A., Gulland, F.M.D., and Spraker, T., 1998, Leptospirosis in rehabilitated Pacific harbor seals from California, J. Wildl. Dis., 34: 407–410. Stevens, E., Lipscomb, T.P., and Gulland, F.M.D., 1999, An additional case of leptospirosis in a harbor seal, J. Wildl. Dis., 35: 150.
0839_frame_C34 Page 778 Tuesday, May 22, 2001 1:44 PM
778
CRC Handbook of Marine Mammal Medicine
Streitfield, M.M., and Chapman, C.G., 1976, Staphylococcus aureus infections of captive dolphins (Tursiops truncatus) and oceanarium personnel, Am. J. Vet. Res., 37: 303–305. 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. Swenshon, M., Lammler, C., and Siebert, U., 1998, Identification and molecular characterization of betahemolytic streptococci isolated from harbor porpoises (Phocoena phocoena) of North and Baltic Seas, J. Clin. Microbiol., 36: 1902–1906. Symmers, W.S., 1983, Possible case of Lobo’s disease acquired in Europe from a bottle-nosed dolphin (Tursiops truncatus), Bull. Soc. Pathol. Exotic, 7: 777–784. Thompson, P., Cousins, D.V., Gow, B.L., Collins, D.M., Williamson, B.H., and Dagnia, H.T., 1993, Seals, seal trainers and mycobacterial infection, Am. Rev. Respir. Dis., 147: 164–167. Thornton, S.M., Nolan, S., and Gulland, F.M.D., 1998, Bacterial isolates from California sea lions (Zalophus californianus), harbor seals (Phoca vitulina), and northern elephant seals (Mirounga angustirostris) admitted to a rehabilitation center along the central California coast, 1994–1995, J. Zoo Wildl. Med., 29: 171–176. Thurman, G.D., and Windsor, I.M., 1984, Serological testing for viral and mycoplasma antibodies in a captive dolphin as an aid in diagnosis and epidemiological control, in 15th Annual International Association for Aquatic Animal Medicine, 75. Tryland, M., 2000, Zoonoses of arctic marine mammals, Infect. Dis. Rev., 2: 55–64. Van Bonn, W.G., Jensen, E.D., House, C., House, J.A., Burrage, T., and Gregg, D.A., 2000, Epizootic disease in captive California sea lions, J. Wildl. Dis., 36: 500–507. Webster, R.G., Geraci, J., Petursson, G., and Skirnisson, K., 1981a, Conjunctivitis in human beings caused by influenza A virus of seals, N. Engl. J. Med., 304: 911. Webster, R.G., Hinshaw, V.S., Bean, W.J., Van Wyke, K.L., Geraci, J.R., St. Aubin, D.J., and Petursson, G., 1981b, Characterization of an influenza A virus from seals, Virology, 113: 712–724. Woods, G.L., and Gutierrez, Y., 1993, Diagnostic Pathology of Infectious Diseases, Lea & Febiger, Philadelphia, 56 pp. Woods, R., Cousins, D.V., Kirkwood, R., and Obendorf, D.L., 1995, Tuberculosis in a wild Australian fur seal (Arctocephalus pusillus doriferus) from Tasmania, J. Wildl. Dis., 31: 83–86. Wunschmann, A., Siebert, U., and Weiss, R., 1999, Rhizopus mycosis in a harbour porpoise from the Baltic Sea, J. Wildl. Dis., 35: 569–573.
0839_frame_C35 Page 779 Tuesday, May 22, 2001 1:45 PM
35
Water Quality Kristen D. Arkush
Introduction Wild marine mammals inhabit dynamic aquatic environments, ranging from Arctic waters to tropical bays. Maintaining good water quality for captive animals is crucial to ensuring their health and well-being. Much of what is known about the basic husbandry requirements of captive marine mammals is derived from the collective experience of zoos, parks, and aquariums. Intuitively, recreating conditions of a species’ natural habitat is the most straightforward approach to captive management. However, in facilities in which natural seawater, for example, is limited, mechanical and biological processes must be developed and/or applied to provide alternatives for water conservation and reuse. This chapter provides an overview of some of the principal issues regarding water quality for marine mammals.
Environmental Considerations Facility design for captive marine mammal habitats has evolved from simple exhibits that emphasized ease of maintenance to those that more closely resemble native habitat of the occupants. Facilities built in the mid-1980s and thereafter have increased exhibit size and versatility, while trying to create more naturalistic habitats (Murphy, 1984; Charfauros, 1986; Hewlett, 1986; Hewlett and Hewlett, 1986; Jones, 1986; Proctor, 1986; Krajniak, 1987). Although more expensive, these exhibits are designed to favor behavioral enrichment. Appropriate social groupings might be considered the most important factor affecting the overall health of cetaceans (McBain, 1999). Most dolphins and small-toothed whales are social species, and efforts to house these species with conspecifics are critical to their care and management. Areas for special needs, such as medical pools or isolation areas for neonates, are frequently included in facility design. Moreover, increased versatility can allow for mixed species displays (Sweeney and Samansky, 1995). Fundamental design considerations for marine mammal holding facilities are based principally on species-specific requirements for salinity (fresh water, brine, artificial seawater, or seawater), temperature, and lighting. Beyond species-specific concerns, water composition may also affect process design, such as the efficacy of sterilization (Spotte, 1991). The Animal Welfare Act, organized and implemented by the U.S. Department of Agriculture (USDA) Animal and Plant Health Inspection Service (APHIS), has established minimum standards for aspects of marine mammal care including space requirements, water quality, nutrition and food preparation, social grouping, transport, and medical treatment (see Chapter 33, Legislation). Many facilities are designed today to exceed those requirements.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
779
0839_frame_C35 Page 780 Tuesday, May 22, 2001 1:45 PM
780
CRC Handbook of Marine Mammal Medicine
Space General space requirements for cetaceans, pinnipeds, sirenians, polar bears, and sea otters are described in APHIS regulations (USDA, 1999), with values for minimum horizontal dimension, depth, and water volume. Pinnipeds, polar bears (Ursus maritimus), and sea otters (Enhydra lutris) also require dry resting and social activity areas. It is important to integrate back holding areas for training purposes and medical support, which may include shallow areas, feeding stations, slide-out areas, and stretcher stations (Sweeney and Samansky, 1995). Medical isolation pools are highly desirable, and should allow for easy access to animals and equipment, variable water depth, and variable salinity (Walsh et al., 1989; Sweeney and Samansky, 1995). While design considerations should include space to allow for normal behavior and activity, pool configuration and/or system components may present risk to the animals. For example, in poorly constructed facilities, animals may consume pool components as materials disintegrate or as animals disassemble parts (Sweeney, 1990). If chlorine is used for water sterilization, adequate air circulation at water level is necessary to prevent noxious gas accumulation and potential inhalation injury (Amundin, 1986; Geraci, 1986). Fecal contamination of pool waters from haul-out areas, particularly in pinniped facilities, can be minimized by positioning drains so that material does not flow into the pool (Dierauf, 1990). Additionally, animals can be kept in an appropriate holding area at night to reduce fecal contamination of the exhibit area (Sweeney and Samansky, 1995).
System Water Source Water systems are described as open (flow-through), semiclosed, or closed. Water source, volume, and quality often dictate system choice. In an open system, the water supply is continuous and enters from a natural source, flows through the pool, and exits with no intentional recirculation. Typically, open systems do not require mechanical filtration, but filters may be added to improve water clarity and reduce intake of fouling organisms or organic material. Semiclosed systems rely on a lower replacement rate of the pool water. Granular media filtration (e.g., sand or a mixture of anthracite, sand, and garnet) is necessary to improve clarity. Closed systems require the most-intensive water treatment, since all the water is reused. Sterilization, temperature control, solids removal, and color reduction are processes incorporated into system design to maintain acceptable water quality. Water changes or additions are made as needed in closed systems.
Temperature Specific temperature ranges for marine mammals are derived principally from the husbandry experience of zoos and aquaria. The USDA regulations (USDA, 1999) stipulate that air and water temperatures encountered by marine mammals must “not adversely affect their health and comfort”; yet acceptable ranges are not provided. Sweeney and Samansky (1995) present general guidelines for water temperature minima and maxima for polar, temperate, and tropical species of pinnipeds and cetaceans. Species-specific ranges are provided elsewhere (Geraci, 1986; Dierauf, 1990; Faulk, 1990; White and Francis-Floyd, 1990; Williams, 1990; Worthy, 1990; Walsh and Bossart, 1999). Facilities should incorporate heaters and chillers into system designs to modulate both air and water temperatures, particularly if subject to diurnal and seasonal extremes in outdoor enclosures.
0839_frame_C35 Page 781 Tuesday, May 22, 2001 1:45 PM
Water Quality
781
Lighting Lighting intensity and duration should resemble conditions encountered by the animal in its natural habitat. This may entail supplying additional lighting for species housed indoors, or providing shelter to control overexposure in outdoor facilities. Lighting for indoor facilities must “provide uniformly distributed illumination which is adequate to permit routine inspections, observations, and cleaning of all parts of the primary enclosure” (USDA, 1999). It is generally desirable to provide natural lighting, but a mixture of candescent and incandescent light similar to the periodicity of an animal’s natural environment is acceptable (Geraci, 1986). Interestingly, polar species tolerate local ambient cycles without detriment (Sweeney and Samansky, 1995).
Salinity and pH Salinity is the measure of all dissolved substances per kilogram of seawater after carbonate has been converted to oxide, bromine and iodine are replaced with chlorine, and organic matter has been oxidized at 480°C (Spotte, 1991). In practice, however, salinity does not vary significantly from the total mass of dissolved solids. Indirect measures of salinity can be made with a refractometer or by conductivity. Alternatively, hydrometer readings or specific gravity values can be obtained and converted to salinity or salinity equivalents (Spotte, 1992). Cetaceans are housed nearly exclusively in artificial or natural seawater. For transport or medical examinations, however, fresh water is generally used to keep cetaceans cool and wet out of water, principally because seawater is corrosive and can easily damage electronic equipment and transporting vehicle components. Pinnipeds are also housed in enclosures with access to seawater. While otariids have been maintained in freshwater enclosures, they may develop ophthalmic injury (e.g., corneal edema) if denied access to salt water (Dunn et al., 1996). Members of the Sirenidae inhabit salt or brackish water with some exposure to freshwater sources such as springs, rivers, or runoff (Walsh and Bossart, 1999). Captive manatees have been held in either fresh water or seawater (with freshwater drinking source) without consequence. Increased skin sloughing has been observed in animals reared in fresh water when moved between seawater and fresh water (Walsh and Bossart, 1999). Marine mammals have been maintained in waters over a wide pH range, at least 6.5 to 8.5, without injury (Spotte, 1991). Although there may be no direct effect of pH on animal health, measuring pH of a marine mammal pool is useful for monitoring chemical processes. For example, as pH increases, a greater mass of chlorine-based oxidant must be added to achieve sterilization. Control of pH is dynamic, but pH can be modulated with the addition of muriatic or hydrochloric acid (lowers pH) or carbonate salts (raises pH). The pH of closed or semiclosed systems declines over time, but bicarbonate and carbonate salts may be added to restore alkalinity (Spotte, 1991).
Filtration There are two types of filtration commonly integrated into process design for marine mammal pool waters: biological and mechanical filtration. Biological filtration is a process by which nitrogenous organic compounds are mineralized (converted into inorganic nitrogen) and either removed from solution by biochemical reduction and lost to the atmosphere or partially reduced and taken up by algae (Spotte, 1992). Most nitrogen in organic matter is mineralized to ammonia (often referred to as NH4-N), which is toxic to most aquatic animals in high concentrations. However, marine mammals can tolerate ammonia levels that would be toxic for fish. In practice, biological filtration is used principally to remove biological waste products
0839_frame_C35 Page 782 Tuesday, May 22, 2001 1:45 PM
782
CRC Handbook of Marine Mammal Medicine
and control bacteria. In the absence of macroalgae (as is usually the case in marine mammal pool waters), bacteria are responsible for biological filtration. Heterotrophic bacteria use mostly organic substances as energy sources, whereas autotrophic bacteria focus on inorganic compounds. Bacteriological filters usually consist of bacteria attached to a solid matrix such as gravel (where the bacterial populations can become dense), but may be as simple as bacterial colonization of the surfaces of submerged system components (e.g., tank walls, pipes). Organic material is added to marine mammal systems continuously. Sources include the influent water, the addition of food, production of urine and feces, and even the animals themselves (e.g., sloughed skin). The sum of all dissolved organic carbon (DOC) plus particulate organic carbon (POC) is referred to as the total organic carbon (TOC). The DOC contains refractory organic matter, such as humus, which is not degraded by biological filtration. Humus accounts for approximately 10 to 50% of the TOC in ocean water (Harvey et al., 1983). It is a class of complex geopolymers derived from carbohydrates, proteins, fatty acids, and lignin (Gjessing, 1976; Christman and Gjessing, 1983; Harvey et al., 1983) that impart a yellow color to seawater. Through the process of adsorption, DOC components, including humus (the adsorbate), can be removed by being bound to an adsorbent. A common and effective adsorbent is activated carbon, which is derived from materials such as animal bones, coal, and wood (Anderson et al., 1981; Loper et al., 1985). The efficiency of adsorption is determined by (1) mass transfer of the adsorbate into the adsorbent, (2) contact time, (3) type and amount of adsorbate, (4) selectivity of the adsorbent, and (5) presence and composition of the biological film on the adsorbent surface (Spotte, 1979). Spotte and Adams (1984) evaluated the adsorptive capacities of various activated carbon sources in artificial seawater systems and found substantial variability among hardwood, bone char, coconut, and anthracite. They suggested that TOC removal efficiency cannot be “predicted”; rather materials should be tested after systems are operational and contain marine mammals. Activated carbon has a limited life span (limited adsorptive capacity) and must be chemically regenerated (a process that is not typically costeffective) or simply replaced. Flocculation is a process by which TOC is precipitated and trapped in the filters as flocs and sediments. Alum and cationic polyelectrolytes (positively charged polymers) are common flocculants. Since flocculation aggregates and increases a portion of the POC and even converts some of the DOC to particulate matter, it improves filtration efficiency and thus reduces oxidant demand (Robinson, 1979; Gregory, 1989). Mechanical filtration, specifically granular media filtration, is used to remove particulate waste and POC. Two types of filter systems exist: pressurized and vacuum (sometimes referred to as gravity feed) filters. Water is pumped under pressure into the former, and the media are usually housed in a steel or fiberglass-reinforced plastic tank. Vacuum filters can be housed in open tanks. A pump may be positioned after the filter unit, so that water is pulled through the media; thus the term gravity feed is not very accurate. Granular media filters are defined by grain size of the filtrant and/or media composition. Rapid sand filters contain either fine sand (<0.5 mm grain size) or coarse sand (>5.0 mm grain size), whereas high-rate filters contain dual media, usually anthracite plus sand, or multimedia, containing anthracite, sand, garnet, and/or ilemnite (Spotte, 1992). In dual and multimedia filters, materials are layered coarse (on top) to fine (bottom) for greatest particulate removal efficiency. Granular media filters are cleaned by backwashing, where filtration is stopped and flow of water is reversed to dislodge waste material. This concentrated wastewater is voided from the system, and then filtration is resumed. To increase cleaning efficiency, auxiliary air scour and surface wash may be added (Spotte, 1992). Air scour agitates the bed through the injection of air below the lowest filtrant material. Surface washers use agitators and water jets positioned above the filtrant to dislodge waste particulates (Tchobanoglous and Schroeder, 1985).
0839_frame_C35 Page 783 Tuesday, May 22, 2001 1:45 PM
Water Quality
783
Foam fractionation may be incorporated to enhance DOC and POC removal (Spotte, 1992). Some organic compounds behave as surfactants, concentrating at the air–water interface. When air bubbles are generated in a contact chamber, surfactants are drawn to the bubbles, where their hydrophobic ends “attach.” This foam can be collected in a column and voided from the system water.
Microorganisms (as Pathogens and/or Indicators of Water Quality) Numerous microorganisms exist in marine mammal pool waters, entering the system from the influent water, the animals, husbandry personnel, and food items. A variety of microbial species have been cultured from both wild and captive animals and their respective environments (see Chapter 16, Bacterial Diseases). Indeed, some species are directly beneficial to the animals (e.g., intestinal microflora) or provide indirect benefits (e.g., heterotrophic species of biological filters). Good medical management of marine mammals includes knowledge of the typical flora of the animals and bacterial species that exist in the water system. Furthermore, it is important to monitor flora over time, as shifts in the microflora of the animals may occur (Buck and Spotte, 1986a; Buck et al., 1989). For example, marine Vibrio spp. occur commonly in captive animals and may be transmitted to new hosts both from tank mates and from the pool surfaces (Kaneko and Colwell, 1978; Buck and Spotte, 1986b). Limiting the introduction and propagation of potentially pathogenic microbes, both through effective water sterilization processes and good husbandry practices, can protect the animals and personnel against exposure. Guidelines on the safe handling and preparation of fish fed to marine mammals are reported elsewhere (Crissey, 1998; see Chapter 36, Nutrition). Since 1979, APHIS has been responsible for the oversight of water quality criteria at marine mammal exhibits and holding facilities under provisions of the Animal Welfare Act (Goff, 1979). Regulations stipulate that water samples be taken from the primary enclosures daily to determine pH, chlorine, and any other chemicals added to maintain water quality standards. Facilities using natural seawater are exempt from these daily measurements unless chemicals are added. Regarding bacterial monitoring of pool waters, samples must be collected at least weekly to enumerate coliform bacteria. The aquatic portions of the enclosures must not exceed a most probable number (MPN) of 1000 coliform bacteria/100 ml of water. If a sample exceeds that limit, two additional water samples must be analyzed within 48 hours, and the average of all three must meet the standard. “Coliform” bacteria likely refer to the coliform group, which consists of several genera of bacteria of the family Enterobacteriaceae (American Public Health Association et al., 1995). Four genera comprise the total coliforms (TC): Escherichia, Klebsiella, Enterobacter, and Citrobacter (Clark and Pagel, 1977). Of the four genera, only Escherichia coli is strictly fecal in homeothermic animals, with no extrafecal sources, and thus is probably the best indicator organism of fecal contamination (Cliver and Newman, 1987). The MPN is determined by the multiple tube fermentation (MTF) method as described in Standard Methods for the Examination of Water and Wastewater (American Public Health Association et al., 1995). In the context of water quality, coliform bacteria are measured as “indicator organisms” of contamination, not as waterborne pathogens. Some investigators have shown that coliform bacteria do not persist in aquatic environments (Joyce and Weiser, 1967; van Donsel et al., 1967; Geldreich, 1970; McFeters and Stuart, 1972). Moreover, their detection is affected by culture method limitations including injury or “stress” to the bacteria during collection, failure to grow in selective media, and inhibition of growth due to presence of competitive bacteria (Spotte, 1991). Similar limitations would apply to other potential “indicator organisms,” so the choice is less important. Interestingly, Grimes et al. (1986) demonstrated that enteric bacteria
0839_frame_C35 Page 784 Tuesday, May 22, 2001 1:45 PM
784
CRC Handbook of Marine Mammal Medicine
have increased survivorship in marine ecosystems, and Farwell and Hymer (1988) described increasing coliform counts in a recirculating system for sea otters. They attributed the contamination to persistence of bacteria in algal mats covering artificial rockwork.
Mechanisms of Sterilization Sterilizing agents commonly used to treat marine mammal pool waters include ultraviolet (UV) sterilization, chlorine-based oxidants, and ozone. Each of these methods can reduce microbial density, but the water is never truly sterile, as input of microorganisms from the animals and other sources is continuous (Spotte, 1991). Treatment options for marine mammal pool waters may be synergistic, whereas others are incompatible, such as chlorination and biological filtration (Sweeney and Samansky, 1995). Two methods of sterilization are used: bulk fluid sterilization and point-contact sterilization. Their relative efficacy and efficiency depend both on the agent and its behavior in water (Spotte, 1991). In bulk fluid sterilization, the sterilizing agent is added to the main area where the animals reside, but may control microbial growth throughout the entire water system. In contrast, point-contact sterilization, as the name suggests, is used to treat a portion of the water system; the effects of sterilization diminish with increasing distance from the site of application. To improve water quality, the rate of sidestream treatment must exceed that of system contamination (Spotte and Adams, 1981). UV sterilization is a point-contact process of irradiation, and is commonly used to treat influent water in a variety of systems. It has been used to treat aquarium systems (Herald et al., 1962; Herald, 1970; Spotte, 1979), hatcheries (Vlasenko, 1969; Kimura et al., 1976; Bullock and Stuckey, 1977; Blogoslawski et al., 1978; Spanier, 1978; Brown and Russo, 1979), wastewaters (Whitby et al., 1984; Qualls et al., 1985; White et al., 1986), and drinking waters (Tobin et al., 1983). UV sterilization does not sterilize the bulk fluid, however, and in closed or semiclosed systems, the microbial density nearest the resident animals may not be diminished (Spotte and Adams, 1981). Spotte and Buck (1981) concluded that UV irradiation is of limited value in the disinfection of marine mammal pool waters. Chlorine-based oxidation is associated with several chlorinated compounds, including sodium hypochlorite (NaOCl), chlorine gas (Cl2), mono- and dichloramine (NH2Cl, HNCl2), chlorine dioxide (ClO2), and their reaction products (Spotte, 1991). These oxidants are considered bulk-fluid sterilizers, because they retain sterilizing properties over some time once generated. Chlorine-based oxidants also react with bromide in natural and artificial seawater − to form hypobromous acid (HOBr) and hypobromite ion (OBr ). Chlorine is applied as a gas or as salts of hypochlorous acid, such as sodium hypochlorite or calcium hypochlorite. Chlorine dioxide is not frequently used, because it is sensitive to temperature, pressure, and light, and can be explosive even at low temperatures. Chlorine gas reacts with water to form hypochlorous and hydrochloric acid: Cl 2 + H 2 O
+
HOCl + H + Cl
(1)
Sodium hypochlorite reacts with water to form hypochlorous acid, sodium ions, and hydroxyl ions: NaOCl + H 2 O
+
HOCl + Na + OH
−
(2)
Whether chlorine is added as a gas or salt, the result is the same, because hypochlorous acid and hypochlorite establish a pH- and temperature-dependent equilibrium: HOCl
−
OCl + H
+
(3)
0839_frame_C35 Page 785 Tuesday, May 22, 2001 1:45 PM
785
Water Quality −
HOCl and OCl together make up the “free available chlorine”; they are available to act as − sterilizing agents. Below pH 7.5 at 25°C, HOCl is the dominant species; OCl is more common above pH 7.5. Hypochlorous acid is considered to have greater sterilizing capacity. Brodtmann and Russo (1979) estimated that HOCl is 150 to 300 times more effective in inactivating viruses, protozoan cysts, and enteric bacteria. Hypochlorous acid also reacts with ammonia to form chloramines (mono-, di-, and trichloramines), and the formation of each is pH and concentration dependent. Together, the amount of these chloramines is considered the combined chlorine or − combined residual chlorine. They persist longer than HOCl and OCl , but are less effective as oxidizing agents. Total chlorine, or total residual chlorine, is the sum of free plus combined chlorine. The U.S. Environmental Protection Agency (1976) recommends that long-term exposure values for total residual chlorine not exceed 0.002 mg/l for salmonid fish and 0.01 mg/l for other aquatic organisms. No limits have been identified for marine mammals, but in practice some parks try to maintain total chlorine below 1 to 1.5 ppm and free chlorine at approximately 50% of total (Reidarson, pers. comm.). For the determination of free, combined, and total residual chlorines, the American Public Health Association et al. (1995) described seven methods. The most commonly used assay is the DPD colorimetric method, and most portable test kit methods are based on this analysis (Tucker, 1993). In addition to chloride, water quality test kits are popular for determination of pH, alkalinity, dissolved oxygen, carbon dioxide, nitrate, nitrite, and total ammonia nitrogen. These kits may sacrifice accuracy and sensitivity as compared with more rigorous analytical methods, but they are portable, relatively inexpensive, and provide results quickly. In paired analyses, these kits provided estimates of water quality parameters within 80 to 120% of values obtained by standard methods (Boyd, 1977; 1980; 1981; Boyd and Hollerman, 1984; Boyd and Daniels, 1988). The use of chlorination for water sterilization in marine mammal pools has been criticized because it may destroy beneficial microflora and inactivate antimicrobial substances secreted by the skin of dolphins (Geraci et al., 1986). Alternatively, systems that rely on biological filtration and physicochemical processes for water purification without chemical sterilization have been described (van der Toorn, 1987; Dudok van Heel and van der Toorn, 1988).
Ozone Ozone is an unstable allotrope of oxygen and must be generated on site by passing a high AC voltage across a discharge gap in the presence of oxygen (Oakes et al., 1979). Ozone is commonly used as a sterilizing agent in public aquarium systems (Sander and Rosenthal, 1975; Stopka, 1975; Honn and Chavin, 1976; Honn, 1979; Spotte, 1979; Ramos and Ring, 1980) and aquaculture facilities (see Summerfelt and Hochheimer, 1997, for review). Although ozone is sometimes used as a clarifying agent (color oxidation), chlorinated waters of closed-system marine mammal pools contain organic compounds that are refractory to ozonation (Adams and Spotte, 1980). Ozone is a powerful oxidizing agent, but drawbacks include its expense because of high capital equipment and operational costs (JMM, 1990) and it poses risk to humans and aquatic life (Bablon et al., 1991). Four processes should be considered in the use of ozone: ozone gas generation, gas-toliquid absorption, contact time for reaction, and ozone residual removal (Summerfelt and Hochheimer, 1997). Ozone in solution can decompose and oxidize materials in two ways. Molecular ozone can react with oxidizable compounds directly (although this process is shortlived and directly affected by pH, bicarbonate level, TOC level, and temperature) (Bablon et al., 1991). The immediate reaction products are free radicals, hydroperoxide species, and unstable ozonide intermediates (Hoigné and Bader, 1976; Peleg, 1976; Prengle, 1983). The second pathway is the indirect action of the oxidizable compounds with radicals formed as ozone
0839_frame_C35 Page 786 Tuesday, May 22, 2001 1:45 PM
786
CRC Handbook of Marine Mammal Medicine
decomposes (Bablon et al., 1991). Once in solution, the half-life of ozone in pure water at 20°C is approximately 165 min (Rice et al., 1981). In aquatic systems with TOC, such as aquaculture facilities, the half-life may be as little as a few minutes (Glaze, 1990) or even seconds (Bullock et al., 1997). − Ozone chemistry is influenced by the presence of bromide (Br ), a component of both fresh water and seawater. It is often a contaminant of granular sodium chloride, and thus is found − in artificial seawater, as well. Ozone oxidizes Br to active bromine and finally bromate (Williams et al., 1978; Crecelius, 1979; Kosak-Channing and Helz, 1979; Kalmaz et al., 1985). Bromide − reacts with ozone to form hypobromous acid (HOBr), and hypobromite ion (OBr ) is oxidized − further to either bromate ( BrO 3 ) or bromide (Haag and Hoigné, 1983). Ozonation in the − presence of bromide is less efficient because Br is regenerated from the intermediate oxidation − − product OBr , causing the catalytic destruction of O 3 , and ultimately increasing the ozone demand (Spotte, 1991). The reactions are as follows: O 3 + Br
O 3 + OBr 2O 3 + OBr
O 2 + OBr −
−
−
2O 2 + Br
(4)
−
(5) −
2O 2 + BrO 3
(6)
Four methods for removing ozone residuals have been described: (1) increasing contact time, (2) passing the treated water through a biofilter or granular activated carbon, (3) stripping the ozone through the use of an aeration column, or (4) destruction with high-intensity UV light (Langlais et al., 1991). Ozone is harmful to aquatic organisms at very low levels. A minimum concentration reported as lethal to fish is 0.01 mg/l (Summerfelt and Hochheimer, 1997). It can oxidize many biochemical compounds, and the generation of highly reactive free radicals during ozonation can damage membrane-bound enzymes and lipids (Summerfelt and Hochheimer, 1997; Carmichael et al., 1982). For human health and safety, ozone concentration in air must be monitored, particularly in the area near the ozone generation equipment.
Conclusions System design for marine mammal pools must incorporate appropriate biological and mechanical methods for water treatment to control temperature, reduce waste, and limit pathogen introduction. Water sources may vary from open, where natural seawater flows through the system continuously, to closed, where natural or artificial seawater is recirculated within the system. Neither is inherently better, and there is a trade-off between vigilant water quality maintenance in closed systems and exposure to natural threats (e.g., pathogens and contaminants) in open systems. As facility design becomes more sophisticated, with an emphasis on naturalistic habitats, the principles of water quality remain fundamental to aquatic animal health and well-being.
Acknowledgments The author thanks Larry Dunn and Greg Lewbart for their constructive comments on earlier drafts of this chapter.
0839_frame_C35 Page 787 Tuesday, May 22, 2001 1:45 PM
Water Quality
787
References Adams, G., and Spotte, S., 1980, Effects of tertiary methods on total organic carbon removal in saline, closed-system marine mammal pools, Am. J. Vet. Res., 41: 1470–1474. American Public Health Association, American Water Works Association, and Water Pollution Control Federation, 1995, Standard Methods for the Examination of Water and Wastewater, 19th ed., American Public Health Association, Washington, D.C. Amundin, M., 1986, Breeding the bottle-nose dolphin Tursiops truncatus at the Kolmarden Dolphinarium, Int. Zoo Yearb., 24/25: 263–271. Anderson, M.C., Butler, R.C., Holdren, F.J., and Kornegay, B.H., 1981, Controlling trihalomethanes with powdered activated carbons, J. Am. Water Works Assoc., 73: 432–439. Bablon, G., and 14 coauthors, 1991, Fundamental aspects, in Ozone in Water Treatment: Application and Engineering, Langlais, B., Reckhow, D.A., and Brink, D.R. (Eds.), American Water Works Association Research Foundation, Denver, CO, 11–132. Blogoslawski, W.J., Steward, M.E., and Rhodes, E.W., 1978, Bacterial disinfection in shellfish hatchery disease control, in Proceedings of the 9th Annual Meeting of the World Mariculture Society, Avault, J.W., Jr. (Ed.), World Mariculture Society, Louisiana State University, Baton Rouge, 589–602. Boyd, C.E., 1977, Evaluation of a water analysis kit, J. Environ. Qual., 6: 381–384. Boyd, C.E., 1980, Reliability of water analysis kits, Trans. Am. Fish. Soc., 109: 239–243. Boyd, C.E., 1981, Comparisons of water analysis kits, in Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies, 34: 39–48. Boyd, C.E., and Daniels, H.V., 1988, Evaluation of Hach Fish Farmer’s Water Quality Test Kits for saline water, J. World Aquacult. Soc., 19: 21. Boyd, C.E., and Hollerman, W.D., 1984, Performance of a Water Analysis Kit, Auburn University Agricultural Experiment Station Circular 274, Auburn, AL. Brodtmann, N.V., Jr., and Russo, P.J., 1979, The use of chloramine for reduction of trihalomethanes and disinfection of drinking water, J. Am. Water Works Assoc., 71: 40–42. Brown, C., and Russo, D.J., 1979, Ultraviolet light disinfection of shellfish hatchery sea water. I. Elimination of five pathogenic bacteria, Aquaculture, 17: 17–23. Buck, J.D., and Spotte, S., 1986a, Microbiology of captive white-beaked dolphins (Lagenorhynchus albirostris) with comments on epizootics, Zoo Biol., 5: 321–329. Buck, J.D., and Spotte, S., 1986b, The occurrence of potentially pathogenic vibrios in marine mammals, Mar. Mammal Sci., 2: 319–324. Buck, J.D., Shepard, L.L., Bubucis, P.M., Spotte, S., McClave, K., and Cook, R.A., 1989, Microbiological characteristics of white whale (Delphinapterus leucas) from capture through extended captivity, Can. J. Fish. Aquat. Sci., 46: 1914–1921. Bullock, G.L., and Stuckey, H.M., 1977, Ultraviolet treatment of water for destruction of five Gramnegative bacteria pathogenic to fishes, J. Fish. Res. Board Can., 34: 1244–1249. Bullock, G.L., Summerfelt, S.T., Noble, A., Webster, A., Durant, M.D., and Hankins, J.A., 1997, Ozonation of a recirculating rainbow trout culture system: I. Effects on bacterial gill disease and heterotrophic bacteria, Aquaculture, 158(1–2): 43–55. Carmichael, N.G., Winder, C., Borges, S.H., Backhouse, B.L., and Lewis, P.D., 1982, The health implications of water treatment with ozone, Life Sci., 30: 117–129. Charfauros, V., 1986, Benefits of a natural presentation in a zoo environment, in Proceedings of the International Marine Animal Trainers Association, 49–50. Christman, R.F., and Gjessing, E.T. (Eds.), 1983, Aquatic and Terrestrial Humic Materials, Ann Arbor Science, Ann Arbor, MI, 161–179. Clark, J.A., and Pagel, J.E., 1977, Pollution indicator bacteria associated with municipal raw and drinking water supplies, Can. J. Microbiol., 23: 465–470. Cliver, D.O., and Newman, R.A. (Eds.), 1987, Drinking water microbiology, J. Environ. Pathol. Toxicol. Oncol. (Spec. Issue), 7: 115–201.
0839_frame_C35 Page 788 Tuesday, May 22, 2001 1:45 PM
788
CRC Handbook of Marine Mammal Medicine
Crecelius, E.A., 1979, Measurements of oxidants in ozonized seawater and some biological reactions, J. Fish. Res. Board Can., 36: 1006–1008. Crissey, S.D., 1998, Handling Fish Fed to Fish-Eating Animals: A Manual of Standard Operating Procedures, U.S. Department of Agriculture, Agricultural Research Service, National Agricultural Library, Washington, D.C., 21 pp. Dierauf, L.A., 1990, Pinniped husbandry, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 553–590. Dudok van Heel, W.H., and van der Toorn, J.D., 1988, A biological approach to dolphinarium water purification: II. A practical application: The Delfinaario in Tampere, Finland, Aquat. Mammals, 14(3): 92–106. Dunn, J.L., Abt, D.A., Overstrom, N.A., and St. Aubin, D.J., 1996, An epidemiologic survey to determine risk factors associated with corneal and lenticular lesions in captive harbor seals and California sea lions, in Proceedings of the 27th International Association for Aquatic Animal Medicine, 100–102. Farwell, C.J., and Hymer, J., 1988, Sea otter water quality management with regard to coliforms, in Proceedings of the American Association of Zoological Parks and Aquaria, 344–348. Faulk, E.Y., 1990, Water quality considerations for marine mammals, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 537–542. Geldreich, E.E., 1970, Applying bacteriological parameters to recreational water quality, J. Am. Water Works Assoc., 62: 113–120. Geraci, J.R., 1986, Husbandry, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 757–760. Geraci, J.R., St. Aubin, D.J., and Hicks, B.D., 1986, The epidermis of odontocetes: A view from within, in Research on Dolphins, Bryden, M.M., and Harrison, R.J. (Eds.), Clarendon Press, Oxford, U.K., 3–21. Gjessing, E.T., 1976, Physical and Chemical Characteristics of Aquatic Humus, Ann Arbor Science, Ann Arbor, MI, 120 pp. Glaze, W.H., 1990, Chemical oxidation, in Water Quality and Treatment, 4th ed., Pontius, F.W. (Ed.), American Water Works Association, Denver, CO, 747–779. Goff, M.T., 1979, Specifications for the humane handling, care, treatment and transportation of marine mammals, Fed. Regist. 44(122): 36868–36883, June 22. Gregory, J., 1989, Fundamentals of flocculation, CRC Crit. Rev. Environ. Control, 19: 185–230. Grimes, D.J., Atwell, R.W., Brayton, P.R., Palmer, L.M., Rollins, D.M., Roszak, D.B., Singleton, F.L., Tamplin, M.L., and Colwell, R.R., 1986, The fate of enteric pathogenic bacteria in estuarine and marine environments, Microbiol. Sci., 3(11): 324–329. Haag, W.R., and Hoigné, J., 1983, Ozonation of bromide-containing waters: Kinetics of formation of hypobromous acid and bromate, Environ. Sci. Technol., 17: 261–267. Harvey, G.R., Boran, D.A., Chesal, L.A., and Tokar, J.M., 1983, The structure of marine fulvic and humic acids, Mar. Chem., 12: 119–132. Herald, E.S., 1970, Ultraviolet sterilization of aquarium water, in Aquarium Design Criteria, Drum and Croaker, spec. ed., Hagen, W. (Ed.), U.S. Department of the Interior, Washington, D.C., 57–71. Herald, E.S., Dempster, R.P, Walters, C., and Hunt, M., 1962, Filtration and ultraviolet sterilization of seawater in large closed, and semi-closed aquarium systems, Bulletin of the Institute of Oceanography, spec. IB, Institute of Oceanography, Monaco, 49–61. Hewlett, S., 1986, A “new” approach to exhibiting cetaceans in captivity, in Proceedings of the American Association of Zoological Parks and Aquaria, 691–694. Hewlett, S., and Hewlett, K.G., 1986, Killer whale habitat: A new approach to cetacean exhibits, in Proceedings of the International Marine Animal Trainers Association, 11–13. Hoigné, J., and Bader, H., 1976, The role of hydroxyl radical reactions in ozonation processes in aqueous solutions, Water Res., 10: 377–386. Honn, K.V., 1979, Ozonation as a critical component of closed marine system design, Ozone Sci. Eng., 1: 11–29. Honn, K.V., and Chavin, W., 1976, Utility of ozone treatment in the maintenance of water quality in a closed marine system, Mar. Biol., 34: 201–209.
0839_frame_C35 Page 789 Tuesday, May 22, 2001 1:45 PM
Water Quality
789
JMM (James M. Montgomery Consulting Engineers), 1990, Cedar River Sockeye Project: Final Citing Report, Report to the Seattle Water Department, Seattle, WA. Jones, J., 1986, Captive pinnipeds without blue swimming pools, in Proceedings of the American Association of Zoological Parks and Aquaria, 341–347. Joyce, G., and Weiser, H.H., 1967, Survival of enteroviruses and bacteriophage in farm pond waters, J. Am. Water Works Assoc., 59: 491–501. Kalmaz, E.E., Eraslan, A.H., and Kim, K.H., 1985, A preliminary kinetics model predicting concentration variations of hypobromous acid and bromate in ozonated marine water, in Proceedings of the 1984 Annual Meeting of the North American Chapter of the International Society for Ecological Modeling, Rykiel, E.J., Jr., and Grant, W.E. (Eds.), Fort Collins, CO, 315–326. Kaneko, T., and Colwell, R.R., 1978, The annual cycle of Vibrio parahaemolyticus in Chesapeake Bay, Microb. Ecol., 4: 135–155. Kimura, T., Yoshimizu, M., Tajima, K., Ezura, Y., and Sakai, M., 1976, Disinfection of hatchery water supply by ultraviolet (U.V.) irradiation—susceptibility of some fish-pathogenic bacterium and microorganisms inhabiting pond waters, Bull. Jpn. Soc. Sci. Fish., 42: 207–211. Kosak-Channing, L., and Helz, G.R., 1979, Ozone reactivity with seawater components, Ozone Sci. Eng., 1: 39–46. Krajniak, E.F., 1987, Opening a new marine mammal exhibit, in Proceedings of the International Marine Animal Trainers Association, 63–66. Langlais, B., Reckhow, D.A., and Brink, D.R., 1991, Ozone in Water Treatment—Application and Engineering, American Water Works Association Research Foundation, Denver, CO. Loper, J.C., Tabor, M.W., Rosenblum, L., and Demarco, J., 1985, Continuous removal of both mutagens and mutagen-forming potential by an experimental full-scale granular activated carbon treatment system, Environ. Sci. Technol., 19: 333–339. McBain, J., 1999, Cetaceans in captivity: A discussion of welfare, J. Am. Vet. Med. Assoc., 214: 1170–1174. McFeters, G.A., and Stuart, D.G., 1972, Survival of coliform bacteria in natural waters: Field and laboratory studies with membrane-filter chambers, Appl. Microbiol., 24: 805–811. Murphy, K., 1984, The Living Seas Pavillion, in Proceedings of the International Marine Animal Trainers Association, 94–100. Oakes, D., Cooley, P., Edwards, L.L., Hirsch, R.W., and Miller, V.G., 1979, Ozone disinfection of fish hatchery waters: Pilot plant results, prototype design and control considerations, in Proceedings of the Annual Meeting of the World Mariculture Society, 10: 854–870. Peleg, M., 1976, Review paper: The chemistry of ozone in the treatment of water, Water Res., 10: 361–365. Prengle, H.W., Jr., 1983, Experimental rate constants and reactor considerations for the destruction of micropollutants and trihalomethane precursors by ozone with ultraviolet radiation, Environ. Sci. Technol., 17: 743–747. Proctor, S., 1986, Max Bell Marine Mammal Center at the Vancouver Aquarium, in Proceedings of the American Association of Zoological Parks and Aquaria, 334–340. Qualls, R.G., Ossoff, S.F., Chang, J.C.H., Dorfman, M.H., Dumais, C.M., Lobe, D.C., and Johnson, J.D., 1985, Factors controlling sensitivity in ultraviolet disinfection of secondary effluents, J. Water Pollut. Control Fed., 57: 1006–1011. Ramos, N.G., and Ring, J.F., 1980, The practical use of ozone in large marine aquaria, Ozone Sci. Eng., 2: 225–228. Rice, R.G., Robson, C.M., Miller, G.W., and Hill, A.G., 1981, Uses of ozone in drinking water treatment, J. Am. Water Works Assoc., 73: 1–44. Robinson, C.N., Jr., 1979, Cationic polyelectrolytes reduce organic matter in turbid surface waters, J. Am. Water Works Assoc., 71: 226–227. Sander, E., and Rosenthal, H., 1975, Application of ozone in water treatment for home aquaria, public aquaria, and for aquaculture purposes, in Aquatic Applications of Ozone, Bogoslawski, W.J., and Rice, R.G. (Eds.), International Ozone Institute, Stamford, CT, 103–114. Spanier, E., 1978, Preliminary trials with an ultraviolet liquid sterilizer, Aquaculture, 14: 75–84.
0839_frame_C35 Page 790 Tuesday, May 22, 2001 1:45 PM
790
CRC Handbook of Marine Mammal Medicine
Spotte, S., 1979, Seawater Aquariums: The Captive Environment, John Wiley & Sons, New York, 413 pp. Spotte, S., 1991, Sterilization of marine mammal pool waters: Theoretical and health considerations, U.S. Department of Agriculture, Animal Plant Health Inspection Service, Technical Bulletin 1797. Spotte, S., 1992, Captive Seawater Fishes: Science and Technology, John Wiley & Sons, New York, 942 pp. Spotte, S., and Adams, G., 1981, Pathogen reduction in closed aquaculture systems by UV radiation: Fact or artifact? Mar. Ecol. Prog. Ser., 6: 295–298. Spotte, S., and Adams, G., 1984, The type of activated carbon determines how much dissolved organic carbon is removed from artificial seawater, Aquacult. Eng., 3: 207–220. Spotte, S., and Buck, J.D., 1981, The efficacy of UV irradiation in the microbial disinfection of marine mammal water, J. Wildl. Dis., 17: 11–16. Stopka, K., 1975, European and Canadian experiences with ozone in controlled closed circuit fresh and salt water systems, in Aquatic Applications of Ozone, Blogoslawski, W.J., and Rice, R.G. (Eds.), International Ozone Institute, Syracuse, NY, 170–176. Summerfelt, S.T., and Hochheimer, J.N., 1997, Review of ozone processes and applications as an oxidizing agent in aquaculture, Prog. Fish Cult., 59: 94–105. Sweeney, J.C., 1990, Marine mammal behavioral diagnostics, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 53–72. Sweeney, J., and Samansky, T., 1995, Elements of successful facility design: Marine mammals, in Conservation of Endangered Species in Captivity: An Interdisciplinary Approach, Gibbons, E.F., Jr., Durrant, B.S., and Demarest, J. (Eds.), State University of New York Press, Albany, 465–477. Tchobanoglous, G., and Schroeder, E.D., 1985, Water Quality: Characteristics, Modeling, Modification, Addison-Wesley, Reading, MA, 768 pp. Tobin, R., Smith, D.K., Horton, A., and Armstrong, V.C., 1983, Methods for testing the efficacy of ultraviolet light disinfection devices for drinking water, J. Am. Water Works Assoc., 75: 481–484. Tucker, C.S., 1993, Water analysis, in Fish Medicine, Stoskopf, M.K. (Ed.), W.B. Saunders, Philadelphia, 166–197. USDA (U.S. Department of Agriculture), 1999, Animal Welfare Regulations, 9 CFR chapter 1, Fed. Regist., U.S. Government Printing Office, Washington, D.C., 114 pp. U.S. Environmental Protection Agency, 1976, Quality Criteria for Water, EPA-440/9-76-023. U.S. Environmental Protection Agency, Washington, D.C. van der Toorn, J.D., 1987, A biological approach to dolphinarium water purification: I. Theoretical aspects, Aquat. Mammals, 13(3): 83–92. van Donsel, D.J., Geldreich, E.E., and Clarke, N.A., 1967, Seasonal variations in survival of indicator bacteria in soil and their contribution to storm-water pollution, Appl. Microbiol., 15: 1362–1370. Vlasenko, M.I., 1969, Ultraviolet rays as a method for the control of diseases of fish eggs and young fishes, Prob. Ichthyol., 9: 697–705. Walsh, M.T., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., Dalton, L.M., and McBain, J.F., 1989, Electrolyte imbalances in cetaceans: Pathogenesis and treatment, in Proceedings of the International Association for Aquatic Animal Medicine, 20: 65. Whitby, G.E., Palmateer, G., Cook, W.G., Maarschalkerweerd, J., Huber, D., and Flood, K., 1984, Ultraviolet disinfection of secondary effluent, J. Water Pollut. Control Fed., 45: 844–850. White, J.R., and Francis-Floyd, R., 1990, Manatee biology and medicine, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 601–623. White, S.C., Jernigan, E.B., and Venosa, A.D., 1986, A study of operational ultraviolet disinfection equipment at secondary treatment plants, J. Water Pollut. Control Fed., 58: 181–192. Williams, P.M., Baldwin, R.J., and Robertson, K.J., 1978, Ozonation of seawater: Preliminary observations on the oxidation of bromide, chloride, and organic carbon, Water Res., 12: 385–388. Williams, T.D., 1990, Sea otter biology and medicine, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 625–648. Worthy, G.A.J., 1990, Nutritional energetics for marine mammals; Addendums, in Handbook of Marine Mammal Medicine, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 489–520.
0839_frame_C36.fm Page 791 Tuesday, May 22, 2001 2:07 PM
36 Nutrition and Energetics Graham A. J. Worthy
Introduction The group of mammals collectively referred to as “marine mammals” has no true basis in taxonomic reality, but is merely a convenient assembly of species that inhabit the marine environment. This group actually consists of members of three orders of mammals: (1) the seals, sea lions, walruses, sea otters, and polar bears (Carnivora); (2) the whales and dolphins (Cetacea); and (3) the manatees and dugongs (Sirenia). These orders differ in their evolutionary histories, nutritional and energy requirements, and prey preferences. Husbandry of marine mammals is often more of an art than a science, primarily due to a lack of data on the nutritional and energetic requirements of most species. No more than a handful of species have been studied adequately in the wild, and even fewer species are regularly held or studied in captivity. This lack of basic data need not impede a general understanding of proper marine mammal care in captivity. Indeed, the details of bioenergetics for a particular species are less important than the concepts that may be extended to any species. The emphasis of this chapter is on these concepts, with the hope that researchers and handlers will take the opportunity to add to the existing database on marine mammal bioenergetics, to ensure that future husbandry may be more effective. Throughout the chapter, both natural history information and requirements of free-ranging animals will be used to illustrate a nutritional approach to the care and rehabilitation of captive animals. Initially, maintenance energy requirements of an individual are considered, as well as how these requirements change on both a seasonal basis and through maturation. Next is a demonstration of how energy requirements can be translated into gross food consumption, taking into account how efficiently an animal utilizes what it ingests. The conversion of gross energy intake into actual food requirements requires knowledge of the composition of the food and how seasonal changes in composition affect the amount of energy and nutrients contained in that food. Once energy requirements have been met, the nutritional aspects of the diet are discussed in terms of potential dietary disorders and how to avoid them.
Energy Requirements The most important facet of animal husbandry is meeting the daily energetic needs of the animal. The magnitude of energy required is a function of body size, activity level, reproductive state, thermoregulatory expenses of the animal, and whether or not the animal is actively growing. These energy expenditures are collectively referred to as the daily metabolic rate.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
791
0839_frame_C36.fm Page 792 Tuesday, May 22, 2001 2:07 PM
792
CRC Handbook of Marine Mammal Medicine
This rate can be conveniently subdivided into two components: maintenance energy and production energy. Maintenance energy costs, which include basal (or standard) metabolism, thermoregulation, and the cost of locomotion, are expenses that are incurred by all animals, but do not include any investment in growth or the production of new offspring. There have been a few recent studies that have monitored food (or energy) intake in captive pinnipeds (Innes et al., 1986; Renouf and Noseworthy, 1990; Markussen et al., 1990; Krockenberger and Bryden, 1993; Rosen and Renouf, 1995; Kastelein et al., 1995) and cetaceans (Kastelein and Vaughan, 1989; Cheal and Gales, 1991; 1992; Kastelein et al., 1993; 1994). Perhaps not surprisingly, these studies all noted a relationship between food intake and factors such as proximate composition (or energy density) of food, water temperature, growth rate of juveniles, pregnancy, and activity level. An understanding of the relative costs of each of these parameters is essential to understanding energetic constraints on marine mammals.
Metabolic Rate Standard, or basal, metabolic rate is the maintenance operating metabolism of an organism, i.e., that metabolic requirement that is needed to sustain life processes of an animal in a resting state. This energy is used to maintain vital cellular activity, respiration, and blood circulation. In the 1930s and 1940s, equations relating basal metabolic rate (BMR) and body size were developed (Kleiber, 1932; Benedict, 1938; Brody, 1945). These relationships can be used to predict the BMR of an average mammal, if its body mass is known. The best known of these relationships is that of Kleiber (1932; 1975), where BMR is estimated using the equation: MR = 3.4 M
0.75
(1)
where MR is metabolic rate in watts (which is the same as J/s) and M is mass in kilograms. In describing the relationship between body mass and BMR, Kleiber emphasized that BMR determinations must meet a rigid set of four criteria. Only those metabolic rates measured (1) on mature animals, (2) within their thermoneutral zone (that range of ambient temperatures in which metabolism is lowest), (3) in the postabsorptive state (postprandial), and (4) while resting (but not asleep) are acceptable as measures of basal metabolism. The violation of any of these conditions can result in a doubling or tripling of metabolic rate. These standardized conditions are critical if one is to be able to compare data either intra- or interspecifically. In retrospect, it is unfortunate that Kleiber (1975) used the term “basal metabolic rate,” because it is not actually the lowest possible metabolic rate measurement. Metabolic rate during sleep may be as little as 40 to 60% of that while awake (Ashwell-Erikson and Elsner, 1981; Huntley, 1984; 1987; Worthy, 1987; Worthy and Lavigne, 1987; Boily and Lavigne, 1996). This has caused some confusion in the literature over the years regarding what BMR really means, and Bligh and Johnson (1972) have suggested it be renamed “standard metabolic rate” (SMR). For decades it was widely accepted that the energy requirements of marine mammals are considerably higher than those of terrestrial mammals of similar size (Irving et al., 1935; Scholander, 1940; Scholander et al., 1942). These high rates were originally thought to result from the need of the animal to cope with the thermal stresses of the cold aquatic environment. High metabolic rates require high food intakes, and these presumed high demands were the reasons cited for marine mammals having such a large impact on commercial fish stocks. The view that marine mammals have high metabolic requirements still generally persists (Irving and Hart, 1957; Hart and Irving, 1959; Kanwisher and Sundnes, 1965; 1966; Ridgway, 1972; Snyder, 1983; Schmidt-Nielsen, 1997). Several recent studies, however, have suggested
0839_frame_C36.fm Page 793 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
793
that marine mammals may have metabolic rates that are similar to those predicted by Kleiber’s equation for terrestrial mammals of similar size, if Kleiber’s four criteria are met (Øritsland and Ronald, 1975; Parsons, 1977; Gallivan and Ronald, 1979; Gaskin, 1982; Lavigne et al., 1982a; Worthy, 1987). Hence, marine mammals would not require any more food than comparably sized terrestrial mammals. Lavigne et al. (1986) reviewed the literature on the metabolic requirements of seals and whales and applied the criteria of Kleiber (1975) to those determinations, eliminating those studies that did not meet the criteria. Their conclusion was that there is no significant difference between the metabolic rates of adult phocid seals and those of terrestrial mammals when determinations meet Kleiber’s criteria. Some data for odontocetes, which meet Kleiber’s criteria, are also not significantly different from terrestrial mammals. With few exceptions, the available data for otariid and cetacean metabolic requirements did not meet the criteria for BMR (Lavigne et al., 1986). Estimates of cetacean metabolic requirements have frequently been obtained using indirect methods. One method includes estimating oxygen consumption by measuring lung volume in dead whales and then measuring respiration rate in free-ranging whales (Scholander, 1940). This can then be converted to an energy estimate, by assuming an energy equivalent for consumed oxygen. Other methods have monitored changes in fat content or changes in girth measurements and then equated these to energy requirements (Brodie, 1975; Kawamura, 1975). Other approaches have used surface area (relating heat losses to heat production) (Brodie, 1975), food intake in captive animals (Sergeant, 1969; Shapunov, 1973; Hinga, 1979; Kastelein and Vaughan, 1989; Cheal and Gales, 1991; 1992; Kastelein et al., 1993; 1994), or extrapolations of BMR estimates from smaller cetaceans to estimate BMR in large cetaceans (Kanwisher and Ridgway, 1983). Some of these extrapolations or estimates are so consistent with available or estimated intake that Brodie (1977) suggested that large cetaceans were burdened “with a [metabolic] rate so high it would appear that their greatest source of natural mortality is spontaneous combustion.” Contrary to most previous work, recent data on small odontocetes suggest that cetaceans possess normal mammalian metabolic rates (Olsen et al., 1969; Hampton et al., 1971; Hampton and Whittow, 1976; Worthy et al., 1987; Kasting et al., 1989; Williams et al., 1992a; Reed et al., 2000). In addition, there are specialized anatomical adaptations that allow cetaceans to compensate for the potentially detrimental effects of high core body temperatures on sperm viability and storage. Small odontocetes regulate the temperature of the internal testes using a countercurrent heat exchanger system (see Chapter 11, Reproduction) (Rommel et al., 1992), similar to that found in several species of phocid seals in which blood is cooled in the hind flippers (Rommel et al., 1995). Sirenians are a unique case. Data suggest that adult (Scholander and Irving, 1941; Gallivan and Best, 1980; Irvine, 1983; Miculka and Worthy, 1994; 1995) and juvenile manatees (Trichechus spp.) (Miculka and Worthy, 1995) possess metabolic rates that are only 25 to 30% of predicted values, resulting in a lack of cold tolerance. Given their herbivorous feeding habits and slow lifestyle, these low rates should not be overly surprising (McNab, 1980). Like terrestrial mammals, elevated metabolic rates are exhibited when animals are active, or forcibly restrained, or not postabsorptive, or not under thermoneutral conditions. Immature mammals, which are actively growing, have metabolic requirements approximately twice that predicted for adult mammals of similar size (Brody, 1945; Lavigne et al., 1986; Worthy, 1987; Worthy et al., 1987; Miculka and Worthy, 1995; Hansen et al., 1995). Data collected from young mammals should not be extrapolated to adults of the same, or other, species without taking this normal elevation of metabolic rate into account; this erroneous extrapolation has been made with cetaceans and pinnipeds, which has led to the common misconception that marine mammals have elevated metabolic rates (Lavigne et al., 1986).
0839_frame_C36.fm Page 794 Tuesday, May 22, 2001 2:07 PM
794
CRC Handbook of Marine Mammal Medicine
Recent results (Lavigne et al., 1985; Innes et al., 1986; Williams et al., 1992a; Hansen et al., 1995; Boily and Lavigne, 1996; Hansen and Lavigne, 1997; Reed et al., 2000) suggest that standard metabolic rates, of at least phocid seals and possibly adult odontocetes, can be estimated using Kleiber’s (1975) equation. Indirect data on metabolic expenditures of otariids suggest that their standard metabolic rates may be similar to those of terrestrial mammals (Innes et al., 1986). Collectively, all these results suggest that the standard, or basal, metabolic requirements of most commonly held marine mammals can be estimated using Equation 1.
Thermoregulation In spite of many studies concentrating on specific components of the energy budgets of marine mammals, few have studied thermoregulatory capabilities, either in air or in water. The thermoneutral zone (TNZ) is that range of temperatures where no additional metabolic energy is necessary and metabolic rates are independent of environmental temperature (Bartholomew, 1977). Its lower end is defined as the lower critical temperature ( Tlc) and is the point where physiological variations in thermal conductance are insufficient to keep heat production in balance with heat losses. Below the Tlc, metabolic rate increases linearly with decreasing environmental temperatures. At the upper critical temperature ( Tuc), metabolic rate often increases as a result of the additional work necessary to dissipate heat (Bartholomew, 1977). The range of temperatures within the TNZ is dependent on body size, degree of insulation, and surface area relationships. Younger animals of a given species have greater ratios of surface area to volume and therefore potentially experience greater rates of heat loss than adults. If animals are maintained below Tlc, metabolism will rise with a concomitant increase in food consumption (Hamilton, 1967). If animals are maintained above their Tuc, overheating and death may occur. With prolonged exposure to temperatures below the TNZ, mammals will increase their insulative layer via increased food intake and, as a result, will experience a shift in the TNZ and a decline in metabolic requirements. Studies of the TNZ of marine mammals have generally concentrated on northern phocids. The TNZ of adult harp seals (Pagophilus groenlandicus) ranges from 0 to 30°C (32 to 86°F) in water (Irving and Hart, 1957; Gallivan and Ronald, 1979; Innes, 1984). Recently weaned harp seal pups, undergoing their normal postweaning fast, showed no change in metabolic rate when in water ranging from 1 to 10°C (0 to 50°F) or in air ranging from approximately 10 to 20°C (50 to 68°F) (Worthy and Lavigne, 1987). Young gray seals (Halichoerus grypus) commencing their postweaning fast had a TNZ similar to harp seals in both air and water (Worthy and Lavigne, 1987; Boily and Lavigne, 1996). The Tlc in water for young harbor seals (Phoca vitulina) was found to be a function of season and body size, with higher Tlc in summer (approximately 20°C; 68°F) than in winter (13°C; 55°F) (Hart and Irving, 1959), when insulation was approximately 30% greater (Irving, 1973). TNZ, in air, for juvenile harbor seals ranged from −13° to +29°C (25 to 84°F) and seals became hyperthermic at air temperatures of 32.5 to 35°C (91 to 95°F) (Hansen et al., 1995; Hansen and Lavigne, 1997). These data also indicate a widening of the TNZ with age, with an 11°C (20°F) drop in the Tlc in the first year of life. Parsons (1977) examined TNZ in a single young ringed seal (Phoca hispida) and determined no change in metabolic rate over the range of 13 to 36.5°C (55 to 98°F) in water. Relatively little information exists on the thermoregulatory capabilities of any otariids. Only the metabolic responses of California sea lions (Zalophus californianus) have been described. Sea lions are generally found in temperate waters at about 15°C (59°F); and have a thinner blubber layer and a greater uninsulated flipper surface area than phocids. Measurement of TNZ
0839_frame_C36.fm Page 795 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
795
for immature California sea lions held in water suggest that the Tlc is approximately 15°C (59°F) and the Tuc is approximately 25°C (77°F) (Liao, 1990). California sea lions are apparently unable to deal with high environmental air temperatures, resorting to behavioral means of thermoregulating (Whittow et al., 1972). The Tuc of this species in air is reportedly between 22 and 30°C (72 and 86°F), with smaller animals able to cope with higher temperatures (Matsuura and Whittow, 1973; South et al., 1976). No increase in metabolic rate was observed at air temperatures as low as 10°C (50°F) (Matsuura and Whittow, 1973; South et al., 1976). There have been suggestions that movements of lactating female otariids are frequently in response to elevated air temperatures (Gentry, 1973; Campagna and Le Boeuf, 1988; Francis and Boness, 1991). Female Juan Fernández fur seals (Arctocephalus phillipii) tend to enter the water in the afternoon when solar radiation is greatest and when air temperatures exceed 20°C (68°F) (Francis and Boness, 1991). Parry (1949) inferred, from his studies on the insulation of harbor porpoise (Phocoena phocoena), that small cetaceans are obliged to remain active to maintain body temperature. Previous data for the Atlantic bottlenose dolphin (Tursiops truncatus) and the Hawaiian spinner dolphin (Stenella longirostris) showed that these species depend upon the energy produced by activity and feeding (see Heat Increment of Feeding, below), as well as marked control over peripheral blood flow, to maintain thermal balance (MacKay, 1964; Hampton et al., 1971; McGinnis et al., 1972; Hampton and Whittow, 1976). This suggests that these small odontocetes should show a strong correlation between food consumption and water temperature. Recent information on the thermal characteristics of harbor porpoise (Phocoena phocoena) and spotted dolphin (Stenella spp.) blubber suggests that another method for controlling heat loss in small cetaceans may be the quality of their insulation. Worthy and Edwards (1990) and Worthy (1991) have demonstrated that harbor porpoise blubber has 92.6 ± 3.6% lipid compared with only 54.9 ± 2.8% in spotted dolphins. This difference in lipid content, in conjunction with thicker blubber, results in harbor porpoises having insulation that is four times as effective as that of spotted dolphins (Worthy and Edwards, 1990). Worthy (1991) reported that, in addition to these two species, Pacific white-sided dolphins (Lagenorhynchus obliquidens), common dolphins (Delphinus spp.), and bottlenose dolphins showed modifications of blubber lipid content that were related to insulative quality, and that in bottlenose dolphins there were indications of seasonal shifts in blubber lipid content. These data suggest that at least harbor porpoises, and perhaps other small cetaceans, may not require elevated metabolic rates at water temperatures as low as 10 to 15°C (50 to 59°F) (Yasui and Gaskin, 1986; Worthy and Edwards, 1990; Williams et al., 1992a). Data from wild Atlantic bottlenose dolphins in Sarasota Bay, Florida, show seasonal changes in blubber depth related to changes in water temperature. In winter, when water temperatures average 20°C (68°F), blubber depths averaged 17.6 mm, whereas in summer these depths declined to 11.4 mm, as water temperatures increased to 30°C (86°F) (G.A.J. Worthy, R.S. Wells, A.J. Read, and D.P. Costa, unpubl. data, 1994). Field metabolic rates (FMR) of subadult and adult male bottlenose dolphins living in this same area were 4.2 to 5.3 times the predicted BMR in summer and 3.1 times the predicted BMR in winter months (Costa et al., 1995). FMR is a cumulative metabolic expenditure that includes all daily activities. The seemingly counterintuitive drop in energy needs in winter could be due to a number of factors, but it clearly suggests that thermoregulation is not a serious issue. Miculka and Worthy (1994; 1995; T.A. Miculka and G.A.J. Worthy, unpubl. data, 2000) collected metabolic rate data from a total of 13 manatees ranging in mass from 125 to 634 kg (275 to 1400 lb). These data suggest that manatees, weighing more than 300 kg (660 lb), exhibit the standard mammalian response to cold temperatures of increasing their metabolism. In general, this response occurred at water temperatures of approximately 19 to
0839_frame_C36.fm Page 796 Tuesday, May 22, 2001 2:07 PM
796
CRC Handbook of Marine Mammal Medicine
20°C (66 to 68°F) (consistent with Irvine, 1983), and individual animals increased their metabolic output by almost 100% when temperatures dipped to 15 °C (59°F). This suggests these animals are capable of dealing with cold, for at least some period of time, paralleling what is observed in the wild. Results for small manatees (<300 kg, or 660 lb) were very different. Younger animals were more susceptible to cold due to an apparent inability to increase their metabolic rate at low temperatures. Even at temperatures as low as 16°C (61°F), these animals showed no indication of an increase in metabolic heat production (Miculka and Worthy, 1995; T.A. Miculka and G.A.J. Worthy, unpubl. data, 2000), yet they became lethargic and began holding their pectoral flippers close to their bodies in apparent attempts to conserve body heat. This would quickly result in hypothermia and death if left even for a few hours. The apparent inability to increase their metabolic rate in response to cold temperatures is puzzling since in most mammalian species this response is independent of age. To offset these metabolic insufficiencies, manatees respond to cold weather by relocating to thermal refugia, such as natural springs or warm-water effluent from power plants or coastal industries (Reynolds and Wilcox, 1985). This response to cold weather conditions is a learned response, with mothers introducing their offspring to warm-water refugia during the prolonged period of maternal dependence common to the species (see Chapter 3, Manatee Case Study). The temperature regimen of any animal must be taken into account when calculating overall energy requirements. Ideally, the animal is kept within its TNZ and therefore requires no additional energy. When this is not possible, or if an animal has recently been relocated, energy needs may be several times what might otherwise be expected if based strictly on body size and activity. With time, the insulative layer will increase and a reassessment of metabolic needs may be required to prevent obesity.
Locomotion Locomotion, like any activity, requires energy, and therefore results in an increase in the animal’s metabolic rate. The absolute amount of energy required is a function of body mass, velocity, time spent traveling, distance traveled, and mode of locomotion (Costello and Whittow, 1975; Innes, 1984; Davis et al., 1985; Williams and Kooyman, 1985; Feldkamp, 1988). Costs of locomotion are usually determined by training an animal to exercise on a treadmill or in a water flume. The animal is forced to travel at a certain velocity and its metabolic rate is measured. These experiments yield data on the moving metabolic rate (watts or J/s) at a given velocity (m/s) and can be used to calculate the cost of transport (J/km) (Davis et al., 1985; Williams and Kooyman, 1985; Feldkamp, 1988). Moving metabolic rate (J/s) increases with speed, but cost of transport (J/km) while swimming decreases curvilinearly with velocity (Figure 1). Feldkamp (1988) measured the cost of swimming in immature California sea lions, finding metabolic expenditure to increase with velocity (Figure 2): MR = 2.1 e
0.48U
(2)
where U is velocity (m/s) and MR is metabolic rate (J/s/kg or W/kg). Davis et al. (1985) performed a similar series of measurements on harbor seals. They found the relationship for metabolic rate and swimming velocity to be: For yearling seals:
MR = 1.7 + 2.09 U
1.42
(3)
0839_frame_C36.fm Page 797 Tuesday, May 22, 2001 2:07 PM
797
14
0.7
12
0.6
10
0.5
8
0.4
6
0.3
4
0.2
2
0.1
0 0
1
2
Velocity
3
(ms-1)
14
0.7
12
0.6
10
0.5
8
0.4
6
0.3
4
0.2
2
0.1
Cost of transport (ml O2 kg-1m-1)
Cost of transport (J kg-1m-1)
Nutrition and Energetics
0 0
0.5
1.0
1.5
2.0
2.5
-1
Velocity (ms )
FIGURE 1 Cost of transport plotted against swimming speed. (From Feldkamp, S.D., J. Exp. Biol., 131, 117, 1988. With permission.)
For adults:
MR = 1.5 + 1.04 U
1.42
(4)
where MR is measured in J/s/kg. The intercepts for these two relationships are equivalent to resting or SMR, and therefore the net cost of swimming is the difference between the two figures (Figure 3).
0839_frame_C36.fm Page 798 Wednesday, May 23, 2001 11:57 AM
798
CRC Handbook of Marine Mammal Medicine
35
Vo2 (ml O2 min-1 kg-1)
30 25 20 15 10 5 0 0
1
2 Velocity
(ms-1
3
)
FIGURE 2 Mass-specific metabolic rate of three sea lions as a function of swimming speed. (From Feldkamp, S.D., J. Exp. Biol., 131, 117, 1988. With permission.)
km.hr-1 1
18
2
3
4
5
yearling seals (33kg) adult seals (63kg)
(5)
6
16 14
(6)
12 10
5
(4)
(3)
4
W. kg-1
Metabolic Rate (ml O2.min-1.kg-1)
(3)
(8) 3
(3) 8 6
(3)
2
4 1 2 0
0.5
1.0
1.4
Swimming Velocity (m.sec-1)
FIGURE 3 Mass-specific metabolic rate as a function of swimming velocity for one adult and two yearling harbor seals. The number of swimming sessions at each speed is shown in parentheses. Metabolic rates at zero swimming velocity are minimum values for seals resting in the water and are equivalent to BMR. (From Davis, R.W. et al., Physiol. Zool., 58, 590, 1985. With permission.)
0839_frame_C36.fm Page 799 Tuesday, May 22, 2001 2:07 PM
799
Nutrition and Energetics
Example 1: Calculate the metabolic rate needed for a 100-kg harbor seal to swim at 1.4 m/s. Using Equation 4, the total metabolic expenditure can be calculated for a 100-kg seal swimming at a speed of 1.4 m/s. This would equate to 3.18 J/s/kg. However, do not forget that this includes an SMR of 1.08 J/s/kg (from Equation 1) and therefore the net cost of swimming at this speed is 2.10 J/s/kg.
Another way of expressing this expenditure is as a net cost of transport (COT), with units of J/kg/m. This is calculated by simply dividing the cost of swimming (2.10 J/s/kg) by swimming velocity (1.4 m/s). The adult harbor seal in the example swimming at 1.4 m/s (the lowest cost of transport speed) (Davis et al., 1985) would have a net COT of 1.5 J/kg/m or 0.0015 MJ/kg/km. This net cost of transport is very similar to that of sea lions, and indeed there appears to be very little difference between the energetic cost of foreflipper propulsion of sea lions and the hind limb propulsion of phocids (Innes, 1984; Feldkamp, 1988). The major difference between otariids and phocids is the amount of time each group spends swimming, with otariids generally being more active. Only limited data are available for swimming cetaceans, but Williams et al. (1992) estimated that for bottlenose dolphins the cost of transport was 1.29 J/kg/m at a swimming speed of 2.1 m/s, increasing to 2.85 J/kg/m at 2.9 m/s. Despite the exponential rise in energy expenditure with speed, swimming marine mammals expend less energy for locomotion than running terrestrial mammals of similar size (Taylor et al., 1982; Williams et al., 1992b; Williams et al., 1993). These reduced costs are possibly due to several factors, including neutral buoyancy and streamlining. Williams (1999) examined total transport costs (COT) in relation to body size for a wide range of marine mammals, including phocids, otariids, large and small odontocetes, and a baleen whale. She found that, over the mass range of 21 to 15,000 kg (46 to 33,000 lb), total transport costs could be estimated by the equation: COT = 7.94 M
−0.28
(5)
where COT is measured in J/kg/m and mass (M) is in kg.
Summary: Average Daily Metabolic Rate Average daily metabolic rate (ADMR) is the total daily energy requirement and includes SMR, thermoregulatory costs, and cost of locomotion. This is the value that is actually important in the determination of energy intake. Most studies of ADMR, or FMR, in other mammals suggest values that are between 1.7 and 3 times SMR (Kooyman et al., 1973; Moen, 1973; McNab, 1980; 1986; Lavigne et al., 1982a; Worthy, 1985). Studies performed on captive and wild marine mammals, using a variety of techniques, suggest rates that fall within this same range (Kooyman et al., 1973; Lavigne et al., 1982a; Worthy, 1987; Murie, 1987). Calculation of ADMR includes the use of three formulae: an estimate of BMR (using Equation 1), an evaluation of whether or not the animal is in its thermoneutral zone, and an assessment of its cost of locomotion (using Equation 2, 3, or 4). This energy requirement is not directly converted into an energy intake. The amount of food that the animal needs to ingest to attain this energetic value is dependent on a variety of other factors, which are further discussed under the Bioenergetic Scheme.
Water Requirements Marine mammals occupy a number of different habitats, primarily marine, but in some instances freshwater rivers, e.g., Amazon River dolphins (Inia geoffrensis) or manatees, or lakes,
0839_frame_C36.fm Page 800 Tuesday, May 22, 2001 2:07 PM
800
CRC Handbook of Marine Mammal Medicine
e.g., some harbor seals. These latter species obviously have a source of fresh water if they need it. Seals have been observed to chew on ice or snow, and captive ones have been observed to drink from a hose or trough (Irving et al., 1935; Rand, 1955; Tarasoff and Toews, 1972; Ridgway, 1972; Renouf et al., 1990), but do they need to drink water under natural conditions? Under normal circumstances there are three sources of water that an animal can use. The most obvious of these is free water that the animal can drink. Less obvious, but of great importance to a number of animals, is the pre-formed and metabolic water in their food. Preformed water is that which is a direct component of food. Since most fish and invertebrates consist of 60 to 80% water, this can supply a considerable amount of free water. Metabolic water is that derived from the metabolism of fat, protein, or carbohydrate. An animal can derive 1.07 g of water from each gram of fat, and 0.4 g of water from each gram of protein that is catabolized. The fattier the fish, the more water and energy are available. When fasting, marine mammals derive not only energy, but also water from their fat reserves. Pilson (1970) maintained a sea lion without access to fresh or salt water for 45 days without adverse nutritional effects. Fasting elephant seal (Mirounga angustirostris) pups reduced urine output by 84% after 10 weeks of fasting (Adams and Costa, 1993) complementing other water-sparing mechanisms (Huntley et al., 1984), a critical adaptation for a species that fasts without access to alternative water sources. Routes of water loss from the body are either by evaporation or via urine or feces. Much of the anatomy and physiology of marine mammals is adapted for reducing water loss. Marine mammals possess either very few (pinnipeds) or no (cetaceans and sirenians) sweat glands (see Chapter 9, Anatomy), and therefore surface evaporation is a minor route of water loss. Similarly, evaporation from the respiratory tract is low because of the presence of countercurrent exchangers, similar to those found in desert species, which help retain moisture (Gallivan and Ronald, 1979; Huntley et al., 1984). Urinary water losses vary depending on the level of protein catabolism. Protein breakdown results in the formation of urea, which is subsequently lost in the urine. Since kidney function dictates how concentrated the urine can become, increased protein catabolism results in increased water losses. Mariposia, or seawater drinking, may be beneficial to animals on a high-protein diet, since seawater can provide urinary osmotic space for urea (Wolf et al., 1959; Hui, 1981; Costa, 1982). Sea otters (Enhydra lutris) have one of the highest reported rates of seawater consumption for any marine mammal, averaging 62 ± 27 ml/kg/day, with a range of 0 to 124 ml/kg/day (Costa, 1982). Incidental ingestion of seawater while swallowing prey is unlikely, since otters consume their prey while floating on their backs (Kenyon, 1969). Instead, sea otters may actively consume seawater to aid in dealing with high rates of urea production (Costa, 1982). Seawater ingestion has been reported in a number of other marine mammals. Captive northern fur seals (Callorhinus ursinus) have been shown to consume very small quantities of seawater (1.8 ml/kg/day) (Fadely, 1988). Similarly, harbor seals have been reported to consume small amounts (4.8 ml/kg/day) of seawater incidental to feeding (Depocas et al., 1972). Common dolphins drank 12 to 13 ml/kg/day of seawater while not feeding, and additionally took in approximately 73 ml/kg/day (70% of total influx) across the skin surface (Hui, 1981). This is similar to what has been reported for harbor porpoises (Anderson and Neilsen, 1973). Feeding Atlantic bottlenose dolphins had a water flux of 42.1 to 71.3 ml/kg/day, 31% from preformed and metabolic water and 69% from drinking water and/or water crossing the skin surface (D.P. Costa and G.A.J. Worthy, unpubl. data, 1990). Seals, sea lions, and porpoises have been shown to either ingest seawater or at least be capable of it (Pilson, 1970; Ridgway, 1972; Bester, 1975; Telfer et al., 1975; Gentry, 1981; Hong et al., 1982). Little is known about the ability of manatees to osmoregulate and maintain water balance, but their anatomy suggests an ability to concentrate their urine (Maluf, 1989; Hill and Reynolds,
0839_frame_C36.fm Page 801 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
801
1989). Manatee osmoregulation and water balance has been examined (Ortiz, 1994; Ortiz et al., 1998; 1999) with animals held under various conditions, including some that were maintained in salt water (34 ppt), with a freshwater source available, and fed seagrasses, beets, and lettuce. When these saltwater animals were fed only seagrass for a period of 9 days, with no access to fresh water, plasma osmolarity and sodium and chloride concentrations increased significantly. The manatees eventually refused to eat the high-salt seagrass, suggesting that wild manatees may require regular access to fresh, or perhaps brackish, water to meet water balance needs. In captive situations, this need is met by drinking fresh water, or by eating food that is high in free water (e.g., lettuce: ∼94% water). Manatees living in fresh water and consuming lettuce had the highest rates of water intake (145 ± 12 ml/kg/day), animals held in salt water, eating lettuce, had significantly lower intakes (45 ± 3 ml/kg/day), while manatees chronically exposed to salt water and eating seagrasses had the lowest rate of intake (21 ± 3 ml/kg/day) (Ortiz et al., 1999).
Fasting and Starvation Fasting is a normal part of the life cycle of many pinnipeds and cetaceans. Indeed, in some species, such as northern elephant seals, adult territorial males may not feed for 5 to 6 months of each year. Prolonged periods of fasting can be associated with lactation, migration, molting, or defending a territory, and may occur in captive situations. Fasting is something marine mammals are physiologically adapted to, both in the wild and in captivity, and occurs whenever an animal has a more important activity to perform, even in the presence of available food (Mrosovsky and Sherry, 1980; Rea, 1995). Starvation differs from fasting, in that it is commonly associated with a decreased or nonexistent food supply. Fasting animals are “adapted to maintain a level of metabolic homeostasis so that critical organ function is maintained,” whereas in starvation “homeostatic control is lost and critical organ function becomes compromised” (Castellini and Rea, 1992). Starving and fasting animals undergo a predictable series of responses. During periods of food deprivation, animals depend on endogenous reserves to maintain homeostasis, maintain glucose availability for the central nervous system (CNS), and meet energy demands. Initially, the body utilizes hepatic glycogen reserves, but these are typically exhausted within days (Phase I). During Phase I, BMR decreases, and in humans can fall to 50% of the nonstarving rate (Castellini and Rea, 1992). Subsequently, the body switches to using proteins (skeletal muscle and internal organs) to act as precursors for producing glucose, and eventually the body begins to mobilize lipids, to spare proteins, and to prolong the body’s ability to go without food. Phase II involves the increased oxidation of lipids, the production of ketone bodies, and the partial sparing of proteins. During this phase, glucose from proteins and ketone bodies supply the CNS, and fat supplies the rest of the body. After a long Phase II period, the body’s protein stores can become seriously depleted. One of the first sources of protein to be depleted is cardiac muscle, and therefore animals with reasonable fat stores remaining can die due to protein wasting. Phase III is entered when 30 to 50% of the body’s proteins have been utilized; recovery from this phase is difficult. Many species of pinnipeds, penguins, and bears, which routinely fast as part of their life cycle, prolong Phase II, and spare body protein for as long as possible (Castellini and Rea, 1992; Rea, 1995). Overall, the basic biochemical pattern associated with food deprivation is similar between fasting and starving (nonfasting-adapted) animals but differs greatly in the duration of the different phases (Castellini and Rea, 1992), in particular that of Phase II. Many species of pinnipeds fast during the annual molt. To allow for proper molt and subsequent hair growth, the animal must be out of the water for prolonged periods of time,
0839_frame_C36.fm Page 802 Tuesday, May 22, 2001 2:07 PM
802
CRC Handbook of Marine Mammal Medicine
Phocid Strategy: nursing on-shore
at-sea feeding
at-sea feeding
Otariid Strategy: nursing on-shore
at-sea feeding
at-sea feeding
FIGURE 4 Phocid and otariid lactation strategies. Phocids nurse their pups for a period of time ranging from 4 days (hooded seals) to several weeks, while fasting on-shore. Otariids nurse their pups for several days during the perinatal period and then go to sea to feed. They subsequently alternate nursing their pups with feeding trips to sea for the duration of the lactation period, which may last for more than a year.
presumably to allow skin temperatures to rise above ambient water temperatures. This requires that the animal haul-out on a beach or on ice to molt, which obviously does not allow for feeding. In captivity, these animals may have the opportunity, but not the inclination, to feed (Mrosovsky and Sherry, 1980). Lactating phocid seals fast for periods of up to 4 weeks (Figure 4), while transferring enormous amounts of energy to their pups (Laws, 1959a; Stewart and Lavigne, 1980; Fedak and Anderson, 1982; Costa et al., 1986). Territorial male phocids may fast for considerably longer periods, of up to 4 months, while they vigorously defend their territories (LeBoeuf and Condit, 1983). Newly weaned phocid pups generally undergo a postweaning fast that may last up to 10 weeks (Ortiz et al., 1978; Pernia et al., 1978; Brodie and Pasche, 1982; Worthy and Lavigne, 1983; 1987; Nordøy and Blix, 1985; Nordøy et al., 1990; 1993a; Reilly, 1991). Some species fast on land (e.g., gray seals or elephant seals), whereas others fast in the water (e.g., harp seals) (Worthy and Lavigne, 1987). As a consequence of the different thermal stresses resulting from fasting either on land or in water, different species have evolved different energy utilization strategies (Worthy, 1985; Worthy and Lavigne, 1987; Nordøy et al., 1990; 1993a; Rea and Costa, 1992; Rea, 1995). These strategies involve the differential usage of blubber and core energy stores to meet energetic demands. The preferential consumption of core reserves of fat and protein (muscle and visceral stores) in harp seals allows this species to conserve its insulative blubber layer to prevent excessive heat loss to the cold aquatic environment (Worthy and Lavigne, 1983; 1987; Nordøy et al., 1993a). Species such as northern elephant seals or gray seals, which fast on land, use blubber almost exclusively as their energy source (Ortiz et al., 1978; Pernia et al., 1978; Worthy and Lavigne, 1987; Nordøy et al., 1990; 1993a; Rea and Costa, 1992), while conserving protein (which provides <4% of MR) (Pernia et al., 1978; Adams and Costa, 1993). These pinniped species commonly exhibit a significant decrease (up to 45%) in resting metabolism during the fast (Worthy and Lavigne, 1987; Nordøy et al., 1990; 1993a; Rea and Costa, 1992; Adams and Costa, 1993; Boily and Lavigne, 1995). Interestingly, harbor seals (which do not generally undergo prolonged fasts) showed only a 20% decline in metabolism during a forced fast and obtained only 77% of their energy from fat (Markussen et al., 1992), suggesting they rely on a relatively high rate of protein utilization.
0839_frame_C36.fm Page 803 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
803
Otariid females and their pups undergo intermittent periods of fasting during lactation (see Figure 4), albeit for considerably shorter periods than do phocids. An otariid female usually nurses the pup for a period of 2 to 3 days during which time the female does not feed. The mother then goes to sea to feed for as little as 1 day in the California sea lion (Gentry and Kooyman, 1986) to as many as 12 days in the Juan Fernández fur seal (Francis and Boness, 1991), during which time the pup fasts. These alternating periods of feeding and fasting continue throughout the nursing period, which may last 6 to 9 months (Gentry and Kooyman, 1986). Male otariids, when holding territories, may not feed for periods of up to 3 months during the breeding season (Gentry and Kooyman, 1986). Apparently, young otariids do not fast once weaned, although they do remain on the beach for some time after weaning, and there is no data on how soon after entering the sea they begin to forage efficiently (Gentry and Kooyman, 1986). Rosen and Trites (1999) found that Steller sea lions (Eumetopias jubatus) reduced BMR by 20% in response to a decrease in energy intake resulting from a switch from a herring (Clupea harengus) to a squid (Loligo spp.) diet. Rice and Wolman (1971) reported that weight losses in fasting, migrating gray whales (Eschrichtius robustus) were due more to usage of internal fat stores, rather than the blubber layer. Similarly, Bryden (1969) noted that in southern elephant seals (Mirounga leonina), fat, muscle, and viscera all declined significantly in mass during the postweaning fast, whereas bone and skin changed very little. Of the mass accumulated during lactation, 34% of blubber and 45% of muscle were consumed during fasting (Bryden, 1969). Other investigations indicate that tissue protein may actually be used extensively during fasting in several species of marine mammals to meet their energy demands. Fasting bottlenose dolphins and Pacific white-sided dolphins lose muscle as rapidly as body fat, and apparently meet their glucose requirements through catabolism of protein (Ridgway, 1972). Animals that maintain blood glucose levels by utilizing protein as a precursor, or alternatively using protein directly as an energy source, must rid their bodies of nitrogenous wastes, potentially resulting in marked water losses. Because muscle tissue is approximately 72% water (Worthy, 1985), it can serve not only as an energy source, but also as a source of pre-formed water.
The Bioenergetic Scheme Using estimates of standard or basal metabolism, cost of locomotion, and thermoregulation, one can now estimate how much energy an individual marine mammal may require. The next question is how much food does this equate to? The bioenergetic scheme is a useful framework to describe the flow of energy through an individual or through a population. This scheme takes into account how efficient the body is at absorbing and processing the energy and nutrients it ingests, and how it allocates whatever energy and nutrients are absorbed. Not all of the energy and nutrients an animal ingests are available to it for maintenance, growth, and reproduction. A relatively large amount is lost as waste. There are basically two schemes in use today. One was suggested by the International Biological Programme (Humphreys, 1979) and the other by the National Research Council (NRC, 1981). Both of these schemes share many features, differing only in how the various losses of energy are arranged (Figure 5). Although variations that are more accurate representations of the physiological processes operating within the animal have been suggested, it is more practical to quantify the various components using simplified schemes. Ingested energy (IE) basically passes through two stages, digestible energy and metabolizable energy, before being retained as either new tissue or reproduction (recovered energy, RE). Energy is lost at various stages in the form of fecal energy (FE), urinary energy (UE), and as body heat
0839_frame_C36.fm Page 804 Tuesday, May 22, 2001 2:07 PM
804
CRC Handbook of Marine Mammal Medicine
Intake of Energy in Food (IE)
Fecal Energy (FE)
Digestible Energy (DE)
Total Heat Production (HE) Metabolizable Energy (ME)
Waste Energy Urine (UE)
Recovered Energy (RE) (useful product)
a. Basal Metabolism b. Voluntary Activity
a. Tissue
c. Product Formation
b. Lactation
d. Digestion and Absorption
c. Ovum (Egg)
e. Thermal Regulation
d. Conceptus e. Wool, Hair, Feathers
*Under some circumstances the energy contained could be considered to be a useful product for fuel.
FIGURE 5 The idealized flow of energy through an animal. (From National Research Council, Nutritional Energetics of Domestic Animals and Glossary of Energy Terms, 2nd ed., National Academy Press, 1981. With permission.)
(HE). According to the first law of thermodynamics, the law of conservation of energy may be stated as: IE = FE + UE + HE + RE
(6)
Since the goal of this exercise is to calculate gross energy intake, it is logical to consider the bioenergetic scheme from the bottom to the top and to backcalculate from maintenance (BMR, thermoregulation, and locomotion) and production (growth and reproduction) costs. To do this requires some knowledge of each of the variables listed in Figure 5.
Maintenance Energy In an animal that is not actively growing or reproducing, BMR, thermoregulation, and locomotion costs equal total energy requirements. These components of the bioenergetic scheme have been discussed above.
Production Energy Reproduction
Marine mammal females typically produce only one offspring per season, except polar bears (Ursus maritimus), which produce two. In the few reported cases where twins are born, one
0839_frame_C36.fm Page 805 Tuesday, May 22, 2001 2:07 PM
805
Nutrition and Energetics
pup usually dies (Spotte, 1982). Pinnipeds also undergo “delayed” or “suspended” implantation (Daniel, 1981), whereby embryonic development is arrested at the blastocyst stage. Further development and implantation are delayed for approximately 3 months (see Chapter 11, Reproduction) (Fisher, 1952). This process allows for parturition to occur at the same time each year, even though gestation is only 9 months long. It may also allow the female time to rebuild her energy reserves after nursing the previous pup. This is especially true for phocid seals, which fast during lactation. There have been suggestions that pinnipeds can also delay parturition until a suitable breeding haul-out site has been located. Species in this category include the harbor seal (Fisher, 1952), the ice-breeding Baltic gray seal (Lockley, 1966), the Steller sea lion (Sandegren, 1970), and the harp seal (Sergeant, 1976; Hammill, 1982). The costs of pregnancy include the production of fetal tissue and associated maternal tissues (such as the placenta) and an increase in maternal metabolism to meet the metabolic demands of these new tissues (QG ; cost of pregnancy). These demands are actually unlikely to make great energy demands until about the fifth month of development (Laws, 1959b), when fetal growth becomes more rapid (Laws, 1959b; Hewer and Backhouse, 1968). Lockyer (1987) calculated that the energy invested into fetal fin whales (Balaenoptera physalus) was minimal during the first 5 months and represented very little energy drain on the female. Average energy density of fetal fin whale tissue at the end of 5 months of gestation was 0.75 kcal/g, compared with 2.94 kcal/g at the end of gestation (Lockyer, 1987). These values are relatively similar to those of other species and suggest that most fetal fat deposition occurs late in pregnancy (Lockyer, 1987). Concurrent with the direct energy investment in terms of fetal growth, the female must expend additional metabolic energy to maintain the metabolic requirements of the growing fetus. This cost of pregnancy can be estimated using a relationship derived for terrestrial mammals (Brody, 1945): QG = 18.48M
1.2
(7)
where M is maternal mass in kilograms and QG is in megajoules (MJ) per day. Lactation costs are considerably greater than those associated with fetal growth. Perez and Mooney (1986) estimated that lactating northern fur seals consumed 160% more energy during nursing than nonlactating animals. These estimates are similar to those for sperm whales (Physeter macrocephalus) (132 to 163%) and minke whales (Balaenoptera acutirostrata) (175 to 186%) (Lockyer, 1981). Preliminary data for California sea lions suggest a similar increase (175%) (G.A.J. Worthy, unpubl. data, 1990). Data from bottlenose dolphins are inconsistent. Spotte and Babus (1980) did not note an increase in food consumption during lactation, whereas Perez and Mooney (1986) found a lactating bottlenose dolphin to consume 170% more food than when not lactating. Ronald and Thomson (1981) reported mass intake of a harbor seal during late pregnancy and through lactation. During the 8 months prior to pregnancy, the average daily intake of the female was 2.1 kg (5 lb) (energy density = 7.7 MJ/kg) (Worthy, 1985). This rate of intake continued for the first 5 months of term, but declined during the last 4 months to between 1.3 and 2.0 kg (5 lb)/day (Ronald and Thomson, 1981). Despite the decline in intake, the female continued to gain weight throughout the last trimester. This continued growth could possibly be due to a decrease in activity and/or an investment of a large portion of ingested protein into the fetus, while the female was utilizing ingested fat and fat reserves as the energy source. A relative increase in lean tissue, which is heavier than fat, could account for this apparent discrepancy.
0839_frame_C36.fm Page 806 Tuesday, May 22, 2001 2:07 PM
806
CRC Handbook of Marine Mammal Medicine
The female harbor seal lost 14 kg (31 lb) at parturition (9 kg or 20 lb pup) and lost a further 20 kg (44 lb) during the 28-day lactation period, returning to the approximate prepregnancy mass. During this period the pup gained 18 kg (40 lb). During the first 12 days of lactation, the adult female ingested 1.6 kg/day of herring, increasing to 4 kg (9 lb)/day by day 20 when her weight had returned to normal nonreproductive mass (Ronald and Thomson, 1981). Unlike the majority of mammals, most phocid seals do not consume food while lactating. Phocids, such as gray, harp, hooded (Cystophora cristata), and elephant seals (Lavigne et al., 1982; Fedak and Anderson, 1982; Stewart and Lavigne, 1984; Bowen et al., 1985; 1987; Costa et al., 1986), produce an energy-rich milk with a high fat content (up to 60% in hooded seals; Oftedal et al., 1988) during a relatively short nursing period (generally less than 4 weeks), while they completely abstain from food and water. Milk composition varies both with species and over the course of the lactation period (see Oftedal et al., 1987; Oftedal, in press). The milk of fasting phocids generally exhibits increasing lipid and concurrent decreasing water content as lactation proceeds (Kooyman and Drabek, 1968; Reidmann and Ortiz, 1979; Lavigne et al., 1982b). This phenomenon has been suggested to be related to water conservation on the part of the female. The initial high water content may also provide the pup with additional free water at a time when it is potentially dependent on protein and carbohydrate metabolism to meet its energy demands, and therefore needs to rid its body of nitrogenous wastes such as urea. Later, as blubber is deposited, the pup presumably switches to a lipid-based metabolism, with the opportunity to use metabolic water (Reidmann and Ortiz, 1979; Lavigne et al., 1982). Otariids and walruses (Odobenus spp.) have considerably longer lactation periods than phocids, ranging from 4 months (northern fur seals) to 2 to 3 years (walrus and Galapagos fur seals, Arctocephalus galapagoensis) (Gentry and Kooyman, 1986). The major difference between otariid and phocid lactation strategies is that, whereas phocids fast continuously for the duration of lactation, otariid females periodically leave their pups and go to sea to feed (see Figure 4). This means that the mother and pup undergo periodic fasts throughout the nursing period. Composition of the otariid milk is adapted to satisfy the pup’s need for sustaining normal activity during maternal absences, with higher fat content correlating with longer periods of maternal absence (Arnold and Trillmich, 1985; Trillmich and Lechner, 1986). For example, Galapagos sea lions (Zalophus californianus wollebacki) visit their young almost daily and have 17% fat in their milk (Trillmich and Lechner, 1986), whereas California sea lions, which are absent for up to 3 days, have 35% milk fat (Trillmich and Lechner, 1986), and northern fur seals, which exhibit feeding trips of 6 to 7 days (Gentry and Holt, 1986; Loughlin et al., 1987), have milk with 50% fat (Ashworth et al., 1966; Dosako et al., 1982; Gentry and Holt, 1986). Milk composition data for manatees (G.A.J. Worthy and O.T. Oftedal, unpubl. data) indicates that the fat content of milk declines from an initial value of ∼21%, to a midlactation level of 18%, and then ultimately to a level of 10% in late lactation after 11 to 12 months. Water content of milk increased during the same time period. Concurrent with this decline in fat content was an increase in the rate of consumption of solid food. Although few data have been available on milk composition for this species, these values are consistent with previously published values (Bachman and Irvine, 1979; Pervaiz and Brew, 1986). These results, based on measurements of one captive manatee, suggest that the calf was consuming approximately 4.0 l of milk per day during the first 2 months of lactation, increasing to 12.2 l/day (day 245) and eventually declining to 2.4 l/day shortly before weaning (day 550). The calf suckled hourly for 1 to 2 min per session and maintained a growth rate of 0.7 kg (2 lb)/day for 550 days. During midlactation, lettuce intake commenced and increased from 2.1 kg (5 lb)/day at age 4 months to 18 kg (40 lb)/ day by 18 months (G.A.J. Worthy and O.T. Oftedal, unpubl. data).
0839_frame_C36.fm Page 807 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
807
Sea otter milk is more similar to milk of other marine mammals than to other mustelids (Jeness et al., 1981). Sea otter milk has 62% water, 23% fat, 13% “protein,” 1% lactose, and 0.8% ash (Jeness et al., 1981). Polar bear milk had a fat content ranging from a high of 38.5% when the cubs emerged from the den in the spring to 20.6% 1 year later (Derocher et al., 1993). Marine mammal milks, in general, are very low in carbohydrate compared with those of other mammals (Jeness, 1974; Derocher et al., 1993). Bottlenose dolphin milk has less than 2.5% neutral sugars with 9 to 12% protein and 10 to 37% fat (Pervaiz and Brew, 1986; Peddemors et al., 1989). Lactose content of harp seal milk was 0.89 ± 0.14 g of lactose per 100 g of whole milk (i.e., <1%) (see Chapter 37, Hand-Rearing) (Stewart et al., 1983). Molt
Molt usually occurs after the mating season for phocid seals and involves the replacement of the entire pelage. Since free-ranging animals generally remain ashore during the molt, which may last 4 to 6 weeks, they are presumably fasting. A lack of interest in feeding has been noted for several captive species during this period (Ronald et al., 1969). Some indoor facilities have found improved progression of the molt under natural photoperiod conditions (Ronald et al., 1969). This is also a period of reorganization of existing protein stores. Since the animals are fasting, all protein required for new hair growth must be of endogenous origin. Commencement of molting in phocids coincides with a decrease in plasma thyroxine and an increase in plasma cortisol (Riviere et al., 1977; Englehardt, 1977; 1979; Ashwell-Erikson and Elsner, 1981) (see Chapter 10, Endocrinology). These hormonal changes are also correlated with a decrease in BMR (Hoch, 1971), which for harbor seals declined to 83% of premolt levels (Ashwell-Erikson and Elsner, 1981). A decline in BMR, in addition to the general lack of activity, suggests that the molt period is one of low energy requirement. Two studies have been completed on molting in free-ranging phocids: the northern (Worthy et al., 1992) and southern elephant seals (Boyd et al., 1993). In both cases, average daily metabolic rate was 2.0 ± 0.6 to 2.4 ± 0.2 times predicted basal metabolic rates, for northern and southern animals, respectively. Overall, elephant seals lost total mass at a rate of 3.0 kg (6.6 lb)/day, with approximately 3.5% of total mass loss associated with shedding of hair and skin (13.5 kg, or 30 lb) (Worthy et al., 1992). Molting for otariids is less of a traumatic experience and does not involve total replacement of the pelage (Scheffer, 1962). Molt in fur seals is an annual event and can take 4 to 5 months. It is gradual, with old hairs being shed singly as new ones erupt. Thus, the fur seal always has a coat that is both an effective insulator and a waterproofer (Scheffer, 1962). The molt is not accompanied by major changes in the epidermis, as is seen in monachine phocids, such as Weddell seals (Leptonychtes weddellii), elephant seals, or monk seals (Monachus spp.), where large sheets of epidermis are lost with the hair.
Heat Increment of Feeding The mechanical and biochemical processes of digestion cause an increase in metabolic rate, which has been termed the heat increment of feeding (HIF) (Brody, 1945; Webster, 1976; Tandler and Beamish, 1979; NRC, 1981), also referred to as specific dynamic action (SDA) (Kleiber, 1975), and is distinct from the cost of physically ingesting the food. There has been some debate over the years about how this term should be defined and indeed whether it should continue to be used (IUPS, 1987). The difficulty with this measurement is that it cannot be distinguished precisely from other factors that stimulate metabolic energy transformations in the hours following food intake. IUPS (1987) suggested that the term be replaced with “post-prandial (excess) heat production,” but the term HIF has remained in common usage.
0839_frame_C36.fm Page 808 Tuesday, May 22, 2001 2:07 PM
808
CRC Handbook of Marine Mammal Medicine
The magnitude and duration of this increase varies with the size and composition of the diet. Carbohydrates increase metabolism by 4 to 30% for 2 to 5 hours after ingestion, lipids from 4 to 15% for 7 to 9 hours, and proteins from 30 to 70% for as long as 12 hours after ingestion (Hoch, 1971). Mixed diets produce lower HIF values than those predicted from the individual components (Forbes and Swift, 1944). Elevation in metabolic rate has been associated with an increase in excretion of urinary nitrogen after intake of amino acids, suggesting that deamination and urea formation may account for at least some of the HIF. HIF is generally considered to be waste energy, unless the animal is experiencing environmental temperatures below its TNZ. Under these conditions this energy could be conserved and used to offset some of the necessary increase in metabolism. HIF has been measured in only a few species of marine mammals, including sea otters (Costa and Kooyman, 1984), harp seals (Gallivan and Ronald, 1981), harbor seals (Ashwell-Erikson and Elsner, 1981; Markussen et al., 1994), ringed seals (Parsons, 1977), northern elephant seals (Barbour, 1993), and Steller sea lions (Rosen and Trites, 1997). There are currently no data available for cetaceans. It has been suggested that sirenians, because of their hindgut fermentation and prolonged food passage times, do not experience a heat increment of feeding, as is also true for other nonruminant herbivores (Gallivan and Best, 1986). Gallivan and Best (1986) did, however, measure the actual cost of feeding, which accounted for 3.4 ± 0.3% of ingested energy for grass (Brachiaria mutica) and 5.4 ± 0.8% of IE for water hyacinth (Eichhornia crassiceps). HIF measured in harbor seals accounted for 4.7 to 9.0% of ingested energy on herring (Clupea harengus) and pollock (Pollachius virens) diets (Ashwell-Erikson and Elsner, 1981; Markussen et al., 1994). HIF in elephant seals ranged from 9 to 11% of ingested energy when consuming herring (8.7% fat) and 11 to 13% when eating capelin (Mallotus villosus) (5.2% fat). Harp seals, on a herring diet, expended up to 17% of ingested energy on HIF (Gallivan and Ronald, 1981). In general, the absolute magnitude of HIF increased with increasing quantity of fish fed. Rosen and Trites (1997) found that HIF in Steller sea lions averaged 12.4 ± 0.9% of energy intake for meals of 4 kg (9 lb), but that the magnitude of HIF dropped to 9.9 ± 0.9% for 2-kg (4.4-lb) meals. Meal size also appeared to affect the duration of HIF, consistent with other studies, ranging from 6 to 8 hours (2-kg meal) to 8 to 10 hours (4-kg meal). Sleep tends to depress overall metabolism, resulting in a metabolic expenditure that can be balanced by increasing HIF and which is approximately the same as standard metabolic rate (Ashwell-Erikson and Elsner, 1981; Worthy, 1985). When harbor seals remained awake, the measured HIF for 1.8 kg (4 lb) of herring was 28.2% of BMR for a 10-hour duration (Ashwell-Erikson and Elsner, 1981). This is comparable with that of ringed seals (30% of BMR for 12 to 13 hours) (Parsons, 1977) and for dogs on a raw meat diet (30% of BMR) (Brody, 1945). Gallivan and Ronald (1981) reported increases in harp seal BMR of 40% for 1 kg (2.2 lb) of herring and 67% for 2 kg (4.4 lb) of herring over the 10-hour period following feeding. Costa and Kooyman (1984) discussed the phenomenon of HIF in sea otters from the perspective of using HIF to offset metabolic expenditures. HIF in otters accounted for 10 to 13% of ingested energy on clam (Spisula) and squid (Loligo spp.) diets, respectively (Costa and Kooyman, 1984). This resulted in a 54% increase over BMR that lasted 4 to 5 hours (0.8 kg, or 2 lb, meal of either diet). Sea otters may use their short and intense HIF increase over BMR to offset heat loss during rest (with its decreased metabolism) and to maintain body temperature (Costa and Kooyman, 1984). Otters also achieve thermoneutrality by augmenting BMR with periodic bursts of activity.
0839_frame_C36.fm Page 809 Tuesday, May 22, 2001 2:07 PM
809
Nutrition and Energetics
Fecal and Urinary Energy Losses Although fecal and urinary energy losses are separate mechanisms (see Figure 5), in terms of backcalculating through the bioenergetic scheme, it is easier to discuss them together. Since the efficiency of assimilation (AE) is not 100%, not all of the food ingested is available to the animal. The efficiency with which animals absorb the different components of a meal varies as a function of the length and morphology of the digestive tract, food type, seasonality, and nutritional state (Petrusewicz and MacFayden, 1970; Moors, 1977; Ashwell-Erikson and Elsner, 1981; Helm, 1984; Sibly and Calow, 1987). Assimilation efficiency also declines under ad libitum conditions (Petrusewicz and MacFayden, 1970). The subtraction of fecal energy from ingested energy gives the “apparent” digestible energy (DE), i.e., that which is absorbed and enters the bloodstream of the animal; “apparent” because not all of the energy that appears to be lost in the feces is of food origin. Digestive enzymes, sloughed intestinal lining, and bacteria add to the energy value of the feces and therefore underestimates the true DE. Measurements performed on fur seals indicate that AE is consistently high, with 88% of ingested capelin being assimilated, 90% of pollock, 92% of squid, and 90 to 93% of herring (Miller, 1978; Fadely et al., 1990) (Table 1). These values are consistent with data collected for TABLE 1 Digestive Efficiencies for Various Pinniped Species Species
n
Prey
Odobenus rosmarus Adult walrus Female walrus Male walrus
4 2 2
Herring Herring Herring
92.7 ± 2.1 94.4 ± 1.2 91.0 ± 0.8
Fisher et al., 1992 Fisher et al., 1992 Fisher et al., 1992
Atlantic cod Arctic cod Halibut Capelin Atlantic herring Parathemisto libellula Thysanoessa spp. Capelin Herring Pandalus borealis
93.2 ± 1.1 93.5 ± 1.7 94.7 ± 1.6 95.7 ± 1.5 96.6 ± 0.9 80.0 ± 3.0 82.0 ± 5.0 92.0 ± 5.0 92.5 – 95.0 72.7 ± 2.3
Lawson et al., 1997b Lawson et al., 1997b Lawson et al., 1997b Lawson et al., 1997b Lawson et al., 1997b Mårtensson et al., 1994 Mårtensson et al., 1994 Mårtensson et al., 1994 Keiver et al., 1984 Keiver et al., 1984
2 2 2 2 2 3
Redfish Capelin Arctic cod Atlantic herring Herring/shrimp Herring
83.2 ± 5.1 86.6 ± 4.4 88.3 ± 1.3 93.8 ± 2.0 92.1 ± 3.7 97.0 ± 0.8
Lawson et al., 1997a Lawson et al., 1997a Lawson et al., 1997a Lawson et al., 1997a Lawson et al., 1997a Parsons, 1977
1 1 2 3
Capelin Pollock Squid Herring
88.0 90.0 92.0 ± 1.1 90.0 ± 1.2
Miller, 1978 Miller, 1978 Miller, 1978 Fadely et al., 1990
5
Herring
92.6 ± 2.1
Ronald et al., 1984
Pagophilus groenlandicus Adult harp seal
Subadult harp seal
Adult harp seal Phoca hispida Adult ringed seal
Callorhinus ursinus Northern fur seal
Halichoerus grypus Adult gray seal
12 12 12 12 12 6 6 6 4 4
%DE
Reference
0839_frame_C36.fm Page 810 Tuesday, May 22, 2001 2:07 PM
810
CRC Handbook of Marine Mammal Medicine
harp seals, gray seals, ringed seals, and walrus where between 92.1 and 97.0% of ingested herring was assimilated (Gallivan and Ronald, 1979; Ashwell-Erikson and Elsner, 1981; Keiver et al., 1984; Ronald et al., 1984; Fisher et al., 1992; Lawson et al., 1997a,b) (see Table 1). AE does vary when other species are consumed (see Table 1) presumably as a result of varying proportions of fat and protein. Harp seals maintained on a shrimp (Pandalus borealis) diet had lower efficiencies (73 to 82%), likely due to the large component of indigestible chitin in shrimp (Keiver et al., 1984; Mårtensson et al., 1994) (see Table 1). Recently there have been attempts to gain a better understanding of digestion in cetaceans. These studies have relied on in vitro determinations of digestibility where the multicompartmental nature of the cetacean stomach is simulated (Nordøy et al., 1993b). These studies suggest that baleen whales have digestive efficiencies comparable with pinnipeds, with an efficiency of 92.1% for herring and 83.4% for krill (Nordøy et al., 1993b). Olsen et al. (1994) have suggested that symbiotic chitinolytic bacteria in the forestomach may enhance some of this efficiency. Sea otters have been shown to have average AE varying from 76.6% for squid, 79.6% for crab (Cancer antennarius), 80.3% for clams (Spisula solidissima), to 87.6% for abalone (Haliotis cracheriodii) (Fausett, 1976). Variation between individuals and seasons resulted in an actual range of AE from a low of 66.3% for a female eating crab in the winter, to a high of 96.5% for a female eating abalone in the winter (Fausett, 1976). AE of West Indian manatees (Trichechus manatus) has been determined for both lettuce (90%) and water hyacinth (80%) diets. For herbivores, AE is inversely correlated with the crude fiber content of the food. Consistently, AE for a lettuce diet (10 to 12% crude fiber) was greater than for water hyacinths (12 to 17% crude fiber) (Lomolino and Ewel, 1984). These measured AE values for manatees are higher than for most herbivores, especially nonruminants, and have been attributed to the extremely slow food passage time (5 to 6 days) (Best, 1981; Lomolino and Ewel, 1984) and extensive digestive tract (18 m) in this species (Quiring and Harlan, 1953; Lomolino and Ewel, 1984). Manatees also have one of the highest digestibility coefficients for cellulose (80%) of any known mammalian herbivore (Burn, 1986). UE losses are directly proportional to nitrogen intake, and animals fed higher-nitrogen diets experience higher UE losses. Urinary losses are usually expressed as a percentage of DE, because the losses are proportional to absorbed nitrogen, and not that which was lost in the feces. Mean energy excreted in urine of harp and gray seals was 6.9 to 9.5% of DE intake on a herring diet (Keiver et al., 1984; Ronald et al., 1984) and for ringed seals was 8.6% (Parsons, 1977). This left 82.7 to 88.7% of IE available as metabolizable energy (accounting for UE and FE losses). These values are similar to those found for ringed seals (Parsons, 1977) and harbor seals (AshwellErikson and Elsner, 1981). Metabolizable energy available to California sea lions is also a function of diet, varying from 78.3% for squid, 88.2% for herring, and 91.6% for anchovy (Engraulis mordax) and 91.4% for mackerel (Scomber japonicus) (Costa, 1986). Similar results have been found for bottlenose dolphins with ME values of 89.2% for mullet (Mugil spp.) and 90.4% for mackerel (Shapunov, 1973) and 88.6% for ringed seals consuming herring (Parsons, 1977). Neither of these latter studies partitioned these energy losses between urinary and fecal routes, and determinations were performed on combined urine and feces.
Calculation of Gross Energy Requirements With information either available or at least estimatable, for each parameter in the bioenergetic scheme, an appraisal of gross energy intake can be made. Assuming, initially, that the animal is not pregnant or actively growing, energy requirements consist of standard metabolic rate,
0839_frame_C36.fm Page 811 Tuesday, May 22, 2001 2:07 PM
811
Nutrition and Energetics
the cost of activity, and thermoregulation. IE can then be calculated by adding HIF and FE and UE losses to energy expenditures. Example 2: Calculate the amount of ingested energy required for a 100-kg adult male phocid seal eating herring under thermoneutral conditions, and swimming at 1 m/s for 6 hours/day. Standard metabolic rate (SMR) can be estimated using Equation 1: SMR = 3.4(100)
0.75
= 107.5 W
Expressing metabolic requirements in watts is not very convenient for the calculation of energy intake, and therefore should be converted to megajoules per day (MJ/day) (1 W = 0.0864 MJ/day). For the purposes of this example, the calculated SMR (107.5 W) is equivalent to 9.29 MJ/day. The net cost of transport for a 100-kg phocid can be estimated (see Example 1) to be 0.15 MJ/ km (0.0015 MJ/kg/km × 100 kg). If it is swimming 21.6 km/day [(1 m/s × 21600 s)/1000 m/km] the total expenditure would be: 0.15 MJ/km × 21.6 km/day = 3.24 MJ/day But not everything the seal eats is available to meet these costs. Energy losses result from the HIF (0.17 × IE) and combined UE and FE (0.15 × IE). The estimated IE is then equivalent to: IE = FE + UE + HIF + ( BMR + locomotion ) IE = ( 0.15 × IE ) + ( 0.17 × IE ) + ( 9.29 + 3.24 ) IE = 18.4 MJ/day This energy requirement could be met by consuming 2.4 kg of herring assuming an energy density of 7.8 MJ/kg (Worthy, 1985). This value compares favorably with intakes ranging from 1.5 to 2.7 kg/day for a 100-kg harp seal feeding on herring of this energy density (Innes, 1984). Example 3: Calculate the amount of fish required to meet the calculated ingested energy requirements. One may not always know the energy content of the fish but if one knows the lipid content, the appropriate energy equivalent value can be roughly estimated. Each gram of lipid in a fish contributes approximately 0.039 MJ of energy. Therefore, 1000 g of fish, that are 20% lipid, would contain approximately 7.8 MJ of energy (0.2 × 1000 g × 0.039 MJ/kg).
Prey Species That Marine Mammals Consume in Captivity and in the Wild Captive marine mammals are generally fed diets that are predominantly commercial fish species, although many marine mammals in the wild may not eat those particular species. Usually, marine mammal diets are dictated by a combination of economics and food species availability. This often results in diets that consist of one, or at best a few, species (Table 2). Results of field studies have consistently shown that marine mammals consume highly varied diets, which change both seasonally and by region (Mathisen et al., 1962; Perez and Bigg, 1986). This variety in the diet allows for potential nutritional shortcomings of one species to be made up for by ingestion of other species. The offering of only one or two food species to a captive mammal may lead to possible nutritional deficiencies.
0839_frame_C36.fm Page 812 Tuesday, May 22, 2001 2:07 PM
812
CRC Handbook of Marine Mammal Medicine
TABLE 2 Prey Species Commonly Fed to Captive Marine Mammals Atlantic herring (Clupea harengus harengus) Pacific herring (Clupea harengus pallasi) Smelt (Osmerus mordax) Atlantic mackerel (Scomber scombrus) Spanish mackerel (Scomberomorus maculatus) Squid (Loligo spp.) Capelin (Mallotus villosus) Butterfish (Peprilus spp.) Scad (Decapterus maruadsi) Whiting (Gadus merlangus) Walleye pollock (Theragra chalcogramma) Cod (Gadus morhua) Clams (Spisula spp.)
TABLE 3 Proximate Composition (±SD) of Species Commonly Fed to Captive Marine Mammals Species
Capture Season
% H2O
% Protein
% Fat
% Ash
Energy (MJ/kg)
Ref.
Pollock
Summer
78.8 (1.3)
19.2 (1.4)
0.8 (0.2)
1.6 (0.2)
4.59 (0.3)
1
Smelt
Winter
74.5 (2.0)
12.1 (0.7)
8.8 (2.4)
1.7 (0.1)
6.80 (1.0)
2
Capelin
Summer
80.3 (1.2)
12.9 (1.8)
3.1 (0.7)
2.4 (0.2)
4.16 (0.7)
2
68.2 (10.1)
16.1 (5.0)
13.7 (7.9)
2.1 (0.4)
8.4 (3.3)
3
Summer
65.8 (1.8)
15.5 (1.0)
13.6 (2.3)
2.3 (0.2)
9.74 (0.6)
2
Autumn
69.7 (1.3)
16.5 (1.4)
8.9 (2.0)
2.3 (0.4)
6.93 (1.6)
2
68.2 (2.8)
17.5 (0.8)
9.9 (3.6)
2.6 (0.2)
8.15 (1.6)
2
Winter
65.0 (3.4)
20.1 (1.3)
13.7 (3.9)
0.8 (0.1)
9.4 (1.4)
3
Mackerel
Summer
74.9
18.8
3.2
3.5
5.59
2
Squid
Winter
76.8 (1.2)
16.7 (1.2)
2.2 (0.9)
1.5 (0.5)
4.58 (0.3)
2
73.5 (5.6)
13.7 (2.7)
10.9 (3.8)
1.9 (0.6)
6.90 (1.8)
3
75.1 (1.4)
17.0 (0.7)
6.6 (1.6)
1.4 (0.1)
5.90 (0.6)
3
Herring
Summer
Note: All values are based on analysis of whole fish (values in parentheses = standard deviation). References: 1. Ashwell-Erikson and Elsner, 1981; 2. Worthy, G.A.J., unpublished data, 1990; 3. Lawson et al., 1998.
0839_frame_C36.fm Page 813 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
813
Seasonal Changes in Prey Composition A sound nutritional program in any captive situation must rely on a good understanding of the composition and quality of the food being offered. All food can be classified with respect to the amount of moisture, protein, and fat it contains. As studies of these components are generally limited to the edible portions (for humans) of commercially important species, there are few data available on the fish species commonly used in marine mammal diets. Knowledge of each of the three major components is of importance in assessing the true value of a diet. The water content provides information regarding the amount of preformed water that is available to the animal, the protein content will influence the amount of waste heat generated as the HIF and UE losses, and the fat content is the major determinant of the energy value of the food. Many prey species show age-related and seasonal changes in proximate composition, and therefore energy content (Table 3). These changes are usually related to reproductive condition, with gravid females being exceptionally high in lipid content. These seasonal changes can be extreme, as in the case of herring where fat content can range from 2 to 4% during early spring to 15 to 20% in the winter (Stoddard, 1988). Other species such as capelin show equally impressive changes in composition (Jangaard, 1974). Mårtensson et al. (1996) noted that the energy density of krill can vary seasonally from a low of 2.3 MJ/kg to a high of 8.2 MJ/kg and that herring show a range of 3.9 to 13.0 MJ/kg. Changes in energy content can lead to a great disparity in energy intake with similar amounts of different fish being consumed. For example, 2 kg (4.4 lb) of herring, with 18% lipid, would have an energy value of 21.7 MJ, whereas 2 kg of capelin, with 3% lipid, would only yield 9.5 MJ of energy. These seasonal and species differences in energy density must be taken into account. These two example species do not differ very much in their protein content, but do differ in the amount of heat lost as the HIF and also in the amount of pre-formed and potential metabolic water. As the concentration of fat increases, the protein to calorie and water to calorie ratios decrease. It also follows that the fat-soluble vitamins will increase and water-soluble vitamins will decrease with increasing fat content.
Major Nutritional Disorders Thiamine Deficiency Thiamine deficiency can be induced by an animal feeding on one or more varieties of fish that contain thiaminase (Table 4). Thiaminase is not a single compound, but a mixture of heatlabile and heat-stable compounds (Fujita, 1954) that are widely found in clupeid (herring) and osmerid (smelt) fishes. In marine mammals, the condition has been tentatively diagnosed in captive gray seals (Myers, 1955), California sea lions (Rigdon and Dragger, 1955), and an Atlantic bottlenose dolphin (White, 1970). Geraci (1972a,b) induced thiamine deficiency in harp seals and showed that clinical signs of the disorder result from vitamin deprivation combined with hyponatremia. In the wild this disorder is likely very rare since animals would be feeding on a variety of fish species, most of which would not contain thiaminase. Thiamine-deprived harp seals developed altered behavior within 40 to 60 days (Geraci, 1972a,b). Seals refuse to eat and become passive and unresponsive to touch, noise, and light stimulation. Breathing may also become irregular and rapid (Geraci, 1972c). This is followed progressively within 2 to 3 days by mild to severe tremors, fore flipper spasms, head shaking,
0839_frame_C36.fm Page 814 Wednesday, May 23, 2001 11:02 AM
814
CRC Handbook of Marine Mammal Medicine
TABLE 4 Presence or Absence of Thiaminase in Fish Commonly Used in Marine Diets Common Name Alewife Anchovy Capelin Atlantic herring Pacific herring Lake herring Atlantic mackerel Pacific mackerel Spanish mackerel Smelt Lake whitefish Whiting a
Scientific Name Alosa pseudoharengus Engraulus mordax Mallotus villosus Clupea harengus harengus Clupea harengus pallasi Coregonus artedii Scomber scombrus Scomber japonicus Scomberomorus maculatus Osmerus mordax Coregonus clupeaformus Gadus merlangus
a
Location M, F M M M M F M M M M, F M, F M
Thiaminase Present Yes Yes Possibly Yes Yes No Possibly Yes No Yes Possibly No
M = marine; F = fresh water.
Source: Geraci, J.R., J. Am. Vet. Med. Assoc., 165, 801, 1974. With permission.
muscle quivering, and death (Geraci, 1972a,b). The most significant clinical diagnostic feature is the finding of abnormal red cell transketolase activity, which requires the apoenzyme thiamine pyrophosphate (Fujita, 1954; Jangaard, 1974; Stoddard, 1988). In the early stages of thiamine deficiency, the progression of the disorder can be prevented by oral or parenteral therapy. Geraci (1972a,b,c; 1974) suggests that an intramuscular injection of 100 mg of thiamine produces a noticeable effect within 1 hour. If the same diet continues to be used, vitamin therapy must continue or the problem will recur. If fish are known to contain thiaminase (see Table 4), one of two vitamin schedules is recommended. Either 5 mg of thiamine for every 4.2 MJ of fish is given 2 hours before feeding or 25 mg/kg of fish is given at the time of the meal (Geraci, 1972c). The former schedule assures that the vitamin is absorbed before the bulk of the enzyme containing fish is eaten. The second compensates for the presence of the “enzyme” by providing surplus thiamine.
Hyponatremia Hyponatremia, or low blood sodium, is characterized by a gradual or sudden decrease in plasma sodium and an equivalent decrease in chloride, but not necessarily a change in potassium (Geraci, 1972a). The condition is manifested by anorexia, followed by uncoordinated or spastic movements progressing to a generalized muscle quivering over the entire body, especially the flippers (Geraci, 1972a,c). The main diagnostic features are low plasma sodium and chloride levels. Sodium values in phocid seals are generally in the range of 147 to 160 mEq/l (Worthy and Lavigne, 1982; Costa and Ortiz, 1982) and in sea lions from 143 to 148 mEq/l (Medway and Geraci, 1978). Plasma levels of below 143 mEq/l are suspect, but levels as low as 120 mEq/l have been reported (Geraci, 1972a). However, some data for apparently healthy harp seal young of the year, undergoing their postweaning fast, indicate that these animals can maintain levels in this range with no apparent debilitating effects, possibly suggesting a greater tolerance in at least young animals (Geraci, 1974; Worthy and Lavigne, 1982). In captivity, the only source of salt is from food, chiefly in the form of thawed whole or cut fish. These fish generally contain salt concentrations that are low enough for human patients requiring a low-salt diet (Thurston, 1958). These levels are further reduced by husbandry practices of thawing fish in water. Geraci (1972a) showed that nearly 25% of sodium is lost
0839_frame_C36.fm Page 815 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
815
after 3 hours of immersion. In the wild, marine mammals consume a varied diet, including invertebrates, which contain considerably more salt. In addition, they receive some salt from incidental ingestion of salt water. Seals that are hyponatremic respond well to parenteral NaCl replacement therapy (Geraci, 1972a). The preferred route of administration is an intraperitoneal injection of 100 to 200 mg NaCl/kg body weight as a saline solution containing 9 to 12 g NaCl/l. This can be either as a slow drip or bolus injection. Treatment may have to be repeated twice daily for the first few days (Geraci, 1972a,c). A daily food supplement of 3 g NaCl/kg fish helps to maintain the electrolyte balance in most seals maintained in fresh water. Seals in seawater maintain a normal electrolyte balance and need not be given a salt-supplemented diet.
Vitamins A, D, and E The fat-soluble vitamins, notably vitamins A, D, and E, are abundant in marine organisms (Sugii and Kinumaki, 1968; Keiver et al., 1988; Dierenfeld et al., 1991): 1 kg (2.2 lb) of herring may provide on the order of 2000 IU of vitamin A, 8000 IU of vitamin D, and 40 to 60 mg (IU) of tocopherol or vitamin E. Therefore, as long as the fish is fresh, adequate levels of these vitamins are present, but they break down quickly in bad fish. Marine and coldwater fish store energy as polyunsaturated fats that remain fluid at low temperatures. These polyunsaturates are also unstable in the presence of oxygen, leading quickly to peroxidation and rancidity. Peroxidation consumes vitamin E in the fish, as well as affecting the marine mammal eating it by increasing its vitamin E requirements. It has been shown that hooded seals, and presumably other marine mammals, have a high capacity for vitamin D storage in their large blubber mass (Keiver et al., 1988), reducing the likelihood of vitamin D deficiency. Vitamin E deficiency results in the accumulation of peroxides and causes steatitis, muscular degeneration, liver necrosis, and anemia (Lanneck, 1973). There is no characteristic manifestation of this problem in marine mammals, and as long as fish are stored at −2°C (30°F) (for maximum periods of 4 months for mackerel, 6 to 7 months for herring, and 9 months for smelt or capelin), minimal supplementation should be required. Ackman (1967) did, however, show a complete loss of tocopherol from fresh frozen cod in only 4 months of shelf life. Englehardt and Geraci (1978) studied the effects of experimental vitamin E deficiency by feeding harp seals headless, eviscerated herring for an 18-month period. The most significant findings were electrolyte imbalances characterized by abnormally low plasma sodium levels (112 to 146 mEq/l), lowered plasma tocopherol levels, and molt irregularities. Plasma enzyme levels were not diagnostic, and food consumption and weight gain were the same as supplemented control animals (Englehardt and Geraci, 1978). A 1-year-old California sea lion that died, potentially due to vitamin E deficiency, was diagnosed as being hypocalcemic, hyperphosphatemic, hyponatremic, with increased blood urea nitrogen (BUN), lactate dehydrogenase, aspartate transaminase (AST), alanine transaminase (ALT), and creatine kinase (Citino et al., 1985). This sea lion exhibited clinical signs of myopathy (i.e., pain, reluctance to move, dyspnea) and gross and microscopic lesions of myopathy and/or steatitis (Citino et al., 1985). Plasma vitamin E levels in deprived harp seals were initially 20 to 42 µg/ml, declining to 15 to 27 µg/ml until the start of the ninth month of deprivation, at which time they rose to predeprivation levels (Englehardt and Geraci, 1978). This latter rise was associated with the molt period and may have been due to mobilization from blubber reserves due to the normal molt fast. Molting seals showed aberrant and incomplete molting patterns, suggesting a potential regulatory function for vitamin E (Englehardt and Geraci, 1978). Concentrations declined again after the molt to levels of 10 to 25 µg/ml.
0839_frame_C36.fm Page 816 Tuesday, May 22, 2001 2:07 PM
816
CRC Handbook of Marine Mammal Medicine
It appears that diets consisting of fish that have been stored for more than 4 months or diets where eviscerated fish are used need supplementation of 100 IU of vitamin E/kg of fish daily (Englehardt and Geraci, 1978). If fish are stored for shorter periods, then a similar sized supplement need only be administered weekly (Geraci, 1972c). There is general agreement that vitamins A and E should be supplemented, but there have been questions about proper dosage because of evidence that large amounts of vitamin A can result in decreases in serum and tissue vitamin E (see Mazzaro et al., 1995, for a review). Mazzaro et al. (1995) examined whether the vitamin E status of northern fur seals was affected by the levels of vitamin A supplementation commonly seen in captive facilities. They concluded that there was indeed a negative effect in some cases and that many institutions may be giving excessive vitamin A. They found that supplementation of 50,000 IU/day of vitamin A had a detrimental effect on vitamin E status in northern fur seals and recommended that lower levels of supplementation be used.
Vitamin C Accidental ascorbic acid deficiency has been reported in bottlenose dolphins, where severe necrotic stomatitis, anorexia, and weight loss responded to a combination of antibiotics and therapeutic vitamins (Miller and Ridgway, 1963). Similar problems were later found to be resolved by provision of 1000 mg ascorbic acid alone, and to be prevented by 200 to 250 mg ascorbic acid daily (Barck Moore, 1980). The stomatitis was considered to be a form of scurvy (Miller and Ridgway, 1963). In this same report, a white-sided dolphin with gingivitis, glossitis, pharyngitis, and large necrotic areas around the teeth is also described (Miller and Ridgway, 1963). Similar cases were cured by the addition of 1000 mg ascorbic acid/day to the diet and the feeding of small, rather than large, fish (Miller and Ridgway, 1963). Pinniped species appear to differ in their ability to synthesize ascorbic acid (Barck Moore, 1980). Northern fur seals and bearded seals (Erignathus barbatus) appear capable of in vivo synthesis, whereas California sea lions, Steller sea lions, ringed, ribbon (Phoca fasciata), and harbor seals do not (Barck Moore, 1980). Dugong (Dugong dugon) and sea otters were also capable of synthesis (Barck Moore, 1980). Pygmy sperm whales (Kogia breuiceps), common dolphins, pilot whales (Globocephala spp.), bottlenose dolphins, and false killer whales (Pseudorca crassidens) were all apparently incapable of synthesis (Barck Moore, 1980). Tissue distribution patterns of ascorbic acid in marine mammal tissues are similar to those reported for humans and guinea pigs (St. Aubin and Geraci, 1980). Adrenal glands contained the highest levels (up to 158 mg/100 g), and muscle and blubber contained the lowest (St. Aubin and Geraci, 1980). Levels in stranded harbor seals, a pilot whale, and a sperm whale, which were emaciated and obviously had not been feeding for some time, did not differ appreciably from levels in healthy animals (St. Aubin and Geraci, 1980). This may suggest some in vivo synthesis, contrary to Barck Moore’s (1980) findings.
Scombroid Poisoning Scombroid poisoning is a potential health hazard whenever poorly preserved scrombroid fish, i.e., mackerel or tuna, are eaten over an extended period. The clinical signs in humans are intense headache, dizziness, abdominal pain, thirst, cardiac palpitation, nausea, vomiting, and diarrhea. These symptoms resemble those produced by histamine or related compounds (Geraci and St. Aubin, 1980). Scombroid fish have high levels of histamine, up to 2 mg/g, and when inadequately preserved or handled, histidine is decarboxylated to histamine (Geraci and St. Aubin, 1980). The toxic effects can be quite high even without visible evidence of putrefaction.
0839_frame_C36.fm Page 817 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
817
There is as yet no direct evidence of scombroid poisoning in marine mammals, but this is probably due to a failure to recognize the problem rather than a resistance of marine mammals to histamine. There have been reports of suspected clinical cases in captive bottlenose dolphins, ringed seals, California sea lions, and killer whales (Orcinus orca) (Geraci and St. Aubin, 1980). These symptoms range from recurring episodes of “sore throat,” to respiratory congestion, and refusal to perform (Geraci and St. Aubin, 1980). The best prevention for scombroid poisoning is to avoid using scombroid fish species (mackerel or tuna) that have been stored beyond their safe shelf life (4 months).
Conclusions The philosophy of this chapter is to encourage the transition of marine mammal husbandry from an art to a science. The care and rehabilitation of marine mammals has often been approached by trial and error over many years of experience with a particular species or group of individual animals. A lack of knowledge regarding the husbandry and specific requirements of many species of marine mammals need not impede estimation of their basic requirements based on knowledge of other species. A general understanding of the bioenergetic scheme and some information on natural history can allow one to make predictions about the metabolic requirements of any animal. Many perceived problems, such as refusing to eat, may be related to normal aspects of the life history of the species. Mass loss may be related to an increase in needs due to obvious changes, such as reproduction or increased activity, or to less obvious ones, such as changes in fish composition or efficiency of assimilation. When basic nutritional needs are not met, animals may divert nutrients from production to maintenance and develop changes in function, related to changes in normal physiological processes, to compensate for stress, leading to decreased resistance to infectious agents and ultimately to disease. The understanding and application of nutritional energetics can prevent many such problems from developing.
Acknowledgments The author thanks Tamara Miculka and Lisa Mazzaro for their comments on this chapter. Some of the unpublished results in this chapter were funded by grants from the Florida Department of Environmental Protection, the Save the Manatee Club, Mote Marine Laboratory, and Hubbs-SeaWorld Research Institute.
References Ackman, R.G., 1967, Influence of lipids on fish quality, J. Food Technol., 2: 169–181. Adams, S.H., and Costa, D.P., 1993, Water conservation and protein metabolism in northern elephant seal pups during the postweaning fast, J. Comp. Physiol. B, 163: 367–373. Anderson, S.H., and Nielsen, E., 1973, Exchange of water between the harbor porpoise, Phocoena phocoena, and the environment, Experientia, 39: 52–53. Arnold, W., and Trillmich, F., 1985, Time budget in Galapagos fur seal pups: The influence of the mother’s presence and absence on pup activity and play, Behaviour, 92: 302–321. Ashwell-Erikson, S., and Elsner, R., 1981, The energy cost of free existence for Bering Sea harbor and spotted seals, in The Eastern Bering Sea Shelf: Oceanography and Resources, Hood, D.W., and Calder, J.A. (Eds.), University of Washington Press, Seattle, 869–899.
0839_frame_C36.fm Page 818 Tuesday, May 22, 2001 2:07 PM
818
CRC Handbook of Marine Mammal Medicine
Ashworth, U.S., Ramaiah, G.D., and Keyes, M.C., 1966, Species differences in the composition of milk with special reference to the northern fur seal, J. Dairy Sci., 49: 1206–1211. Bachman, K.C., and Irvine, A.B., 1979, Composition of milk from the Florida manatee, Trichechus manatus latirostris, Comp. Biochem. Physiol., 62A: 873–878. Barbour, A.S., 1993, Heat Increment of Feeding in Juvenile Northern Elephant Seals, M.Sc. thesis, University of California, Santa Cruz, 47 pp. Barck Moore, L., 1980, Ascorbic Acid Biosynthesis in Certain Species of Marine Mammals, Ph.D. thesis, University of California, Berkeley, 116 pp. Bartholomew, G.A., 1977, Body temperature and energy metabolism, in Animal Physiology: Principles and Adaptations, Gordon, M.S. (Ed.), Macmillan, New York, 364 pp. Benedict, F.G., 1938, Vital Energetics: A Study in Comparative Basal Metabolism, No. 503, Carnegie Institute of Washington, Washington, D.C., 232 pp. Best, R.C., 1981, Food and feeding habits of wild and captive sirenia, Mamm. Rev., 11: 3–29. Bester, M.N., 1975, The functional morphology of the kidney of the Cape fur seal, Arctocephalus pusillus (Schreber), Modoqua Ser. II, 4: 69–92. Bligh, J., and Johnson, K.G., 1972, Glossary of terms for thermal physiology, J. Appl. Physiol., 35: 941–961. Boily, P., and Lavigne, D.M., 1995, Resting metabolic rates and respiratory quotients of gray seals (Halichoerus grypus) in relation to time of day and duration of food deprivation, Physiol. Zool., 68: 1181–1193. Boily, P., and Lavigne, D.M., 1996, Thermoregulation of juvenile grey seals, Halichoerus grypus, in air, Can. J. Zool., 74: 201–208. Bowen, W.D., Oftedal, O.T., and Boness, D.J., 1985, Birth to weaning in 4 days: Remarkable growth in the hooded seal, Cystophora cristata, Can. J. Zool., 63: 2841–2846. Bowen, W.D., Boness, D.J., and Oftedal, O.T., 1987, Mass transfer from mother to pup and subsequent mass loss by the weaned pup in the hooded seal, Cystophora cristata, Can. J. Zool., 65: 1–8. Boyd, I., Arnbom, T., and Fedak, M., 1993, Water flux, body composition, and metabolic rate during molt in female southern elephant seals (Mirounga leonina), Physiol. Zool., 66: 43–60. Brodie, P.F., 1975, Cetacean energetics, an overview of intraspecific size variation, Ecology, 56: 152–161. Brodie, P.F., 1977, Form, function and energetics of cetacea: A discussion, in Functional Anatomy of Marine Mammals, Harrison, R.J. (Ed.), Academic Press, New York, 45–56. Brodie, P.F., and Pasche, A.J., 1982, Density dependent condition and energetics of marine mammal populations in multispecies fisheries management, Can. Spec. Publ. Fish. Aquat. Sci., 59: 35–38. Brody, S., 1945, Bioenergetics and Growth, Hafner Publishing, New York, 236 pp. Bryden, M.M., 1969, Relative growth of the major body components of the southern elephant seal, Mirounga leonina (Linn.), Physiol. Zool., 17: 153–177. Burn, D.M., 1986, The digestive strategy and efficiency of the West Indian manatee, Trichechus manatus, Comp. Biochem. Physiol., 85A: 139–142. Campagna, C., and Le Boeuf, B.J., 1988, Thermoregulatory behaviour of southern sea lions and its effect on mating strategies, Behaviour, 107: 73–90. Castellini, M.A., and Rea, L.D., 1992, The biochemistry of natural fasting at its limits, Experientia, 48: 575–582. Cheal, A.J., and Gales, N.J., 1991, Body mass and food intake in captive, breeding bottlenose dolphins, Tursiops truncatus, Zoo Biol., 10: 451–456. Cheal, A.J., and Gales, N.J., 1992, Growth, sexual maturity and food intake of Australian Indian Ocean bottlenose dolphins, Tursiops truncatus, in captivity, Aust. J. Zool., 40: 215–223. Citino, S.B., Montali, R.J., Bush, M., and Phillips, L.G., 1985, Nutritional myopathy in captive California sea lions, J. Am. Vet. Med. Assoc., 187: 1232–1233. Costa, D.P., 1982, Energy, nitrogen and electrolyte flux and sea water drinking in the sea otter, Enhydra lutris, Physiol. Zool., 55: 35–44.
0839_frame_C36.fm Page 819 Tuesday, April 9, 2002 2:15 PM
Nutrition and Energetics
819
Costa, D.P., 1986, Assessment of the impact of the California sea lion and northern elephant seal on commercial fisheries, California Sea Grant College Program Final Report No. R/F-92, Institute of Marine Sciences, University of California, San Diego, 12 pp. Costa, D.P., and Kooyman, G.L., 1984, Contribution of specific dynamic action to heat balance and thermoregulation in the sea otter, Enhydra lutris, Physiol. Zool., 57: 199–203. Costa, D.P., and Ortiz, C.L., 1982, Blood chemistry homeostasis during prolonged fasting in the northern elephant seal, Am. J. Physiol., 242: R591–R595. Costa, D.P., Le Boeuf, B.J., Huntley, A.C., and Ortiz, C.L., 1986, The energetics of lactation in the northern elephant seal, Mirounga angustirostris, J. Zool. London, 209: 21–33. Costa, D.P., Worthy, G.A.J., Wells, R.S., Read, A.J., Waples, D., Scott, A.B., and Irvine, A.B., 1995, Freeranging energetics of the bottlenose dolphin, presented at 11th Biennial Conference on the Biology of Marine Mammals, Orlando, FL, 14–18 December, 25. Costello, R.R., and Whittow, G.C., 1975, Oxygen cost of swimming in a trained California sea lion, Comp. Biochem. Physiol., 50A: 645–647. Daniel, J.C., 1981, Delayed implantation in the northern fur seal (Callorhinus ursinus) and other pinnipeds, J. Reprod. Fertil., Suppl., 29: 35–50. Davis, R.W., Williams, T.M., and Kooyman, G.L., 1985, Swimming metabolism of yearling and adult harbor seals, Phoca vitulina, Physiol. Zool., 58: 590–596. Depocas, F., Hart, J., and Fisher, H.D., 1972, Sea water drinking and water flux in starved and in fed harbour seals, Phoca vitulina, Can. J. Physiol. Pharmacol., 49: 53–62. Derocher, A.E., Andriashek, D., and Arnould, J.P.Y., 1993, Aspects of milk composition and lactation in polar bears, Can. J. Zool., 71: 561–567. Dierenfeld, E., Katz, N., Pearson, J., Murro, F., and Asper, E., 1991, Retinol and alpha-tocopherol concentrations in whole fish commonly fed in zoos and aquariums, Zoo Biol., 10: 119–125. Dosako, S., Taneya, S., Kimura, A., Ohmori, T., Daikoku, H., Suzuki, N., Sawa, J., Kano, K., and Katayama, S., 1982, Milk of the northern fur seal: Composition, especially carbohydrates and protein, J. Dairy Sci., 66: 2076–2083. Englehardt, F.R., 1977, Changes in physical and chemical characteristics of blood in phocid seals as related to molt, presented at 2nd Biennial Conference on the Biology of Marine Mammals, San Diego, CA, 12–15 December, 42. Englehardt, F.R., 1979, Functional roles of cortisol in phocid seals, presented at 3rd Biennial Conference on the Biology of Marine Mammals, Seattle, WA, 7–11 October, 17. Englehardt, F.R., and Geraci, J.R., 1978, Effects of experimental vitamin E deprivation in the harp seal, Phoca groenlandica, Can. J. Zool., 56: 2186–2193. Fadely, B.S., 1989, Determination of Seawater Ingestion and Food Intake by Water and Sodium Turnover in Juvenile Northern Fur Seals (Callorhinus ursinus), M.Sc. thesis, University of California, Santa Cruz, 55 pp. Fadely, B.S., Worthy, G.A.J., and Costa, D.P., 1990, Assimilation efficiency of northern fur seals determined using dietary manganese, J. Wildl. Manage., 54: 246–251. Fausett, L.L., 1976, Assimilation Efficiency of Captive Sea Otters, Enhydra lutris (Carnivora: Mustelidae), M.A. thesis, California State University, Long Beach, 39 pp. Fedak, M.A., and Anderson, S.S., 1982, The energetics of lactation: Accurate measurements for a large wild mammal, the grey seal (Halichoerus grypus), J. Zool. London, 198: 473–479. Feldkamp, S.D., 1987, Swimming in the California sea lion: Morphometrics, drag and energetics, J. Exp. Biol., 131: 117–135. Fisher, H.D., 1952, The status of the harbour seal in British Columbia, with particular reference to the Skeena River, Bull. Fish. Res. Board Can., 93: 1–58. Fisher, K.I., Stewart, R.E.A., Kastelein, R.A., and Campbell, L.D., 1992, Apparent digestive efficiencies in walruses (Odobenus rosmarus) fed herring (Clupea harengus) and clams (Spisula sp.), Can. J. Zool., 70: 30–36. Forbes, E.B., and Swift, R.W., 1944, Associative dynamic effect of protein, carbohydrate and fat, J. Nutr., 27: 453–468.
0839_frame_C36.fm Page 820 Tuesday, May 22, 2001 2:07 PM
820
CRC Handbook of Marine Mammal Medicine
Francis, J.M., and Boness, D.J., 1991, The effect of thermoregulatory behaviour on the mating system of the Juan Fernández fur seal, Arctocephalus phillipii, Behaviour, 119: 104–126. Fujita, A., 1954, Thiaminase, in Advances in Enzymology, Vol. 15, Nord, F.F. (Ed.), Interscience, New York, 389–421. Gallivan, G.J., and Best, R.C., 1980, Metabolism and respiration of the Amazonian manatee (Trichechus inunguis), Physiol. Zool., 53: 245–253. Gallivan, G.J., and Best, R.C., 1986, The influence of feeding and fasting on the metabolic rate and ventilation of the Amazonian manatee (Trichechus inunguis), Physiol. Zool., 59: 552–557. Gallivan, G.J., and Ronald, K., 1979, Temperature regulation in freely diving harp seals (Phoca groenlandica), Can. J. Zool., 57: 2256–2263. Gallivan, G.J., and Ronald, K., 1981, Apparent specific dynamic action in the harp seal (Phoca groenlandica), Comp. Biochem. Physiol., 69A: 579–581. Gaskin, D.E., 1982, The Ecology of Whales and Dolphins, Heinemann, London, 459 pp. Gentry, R.L., 1973, Thermoregulatory behaviour of eared seals, Behaviour, 46: 73–93. Gentry, R.L., 1981, Seawater drinking in eared seals, Comp. Biochem. Physiol., 68A: 81–86. Gentry, R.L., and Holt, J.R., 1986, Attendance behavior of northern fur seals, in Fur Seals, Maternal Strategies on Land and at Sea, Gentry, R.L., and Kooyman, G.L. (Eds.), Princeton University Press, Princeton, NJ, 41–60. Gentry, R.L., and Kooyman, G.L. (Eds.), 1986, Fur Seals, Maternal Strategies on Land and at Sea, Princeton University Press, Princeton, NJ, 291 pp. Geraci, J.R., 1972a, Hyponatremia and the need for dietary salt supplementation in captive pinnipeds, J. Am. Vet. Med. Assoc., 161: 618–623. Geraci, J.R., 1972b, Experimental thiamine deficiency in captive harp seals, Phoca groenlandica, induced by eating herring, Clupea harengus, and smelts, Osmerus mordax, Can. J. Zool., 50: 179–195. Geraci, J.R., 1972c, Dietary disorders in marine mammals: Synthesis and new findings, J. Am. Vet. Med. Assoc., 179: 1183–1189. Geraci, J.R., 1974, Thiamine deficiency in seals and recommendations for its prevention, J. Am. Vet. Med. Assoc., 165: 801–803. Geraci, J.R., and St. Aubin, D.J., 1980, Nutritional disorders of captive fish-eating animals, in Pathology of Zoo Animals, Montali, R.J., and Migaki, G. (Eds.), Smithsonian Institution Press, Washington, D.C., 41–49. Hamilton, C.L., 1967, Food and temperature, in Handbook of Physiology, Vol. 1, Code, C.F. (Ed.), American Physiological Society, Bethesda, MD, 238 pp. Hammill, M.O., 1982, Late pupping by the harp seal, Pagophilus groenlandicus, in the eastern Canadian Arctic, Can. Field Nat., 96: 216–217. Hampton, I.F.G., and Whittow, G.C., 1976, Body temperature and heat transfer in the Hawaiian spinner dolphin, Stenella longirostris, Comp. Biochem. Physiol., 55A: 195–197. Hampton, I.F.G., Whittow, G.C., Szekerczes, J., and Rutherford, S., 1971, Heat transfer and body temperature in the Atlantic bottlenose dolphin, Tursiops truncatus, Int. J. Biochem. Biometab., 15: 247–253. Hansen, S., and Lavigne, D.M., 1997, Ontogeny of the thermal limits in the harbor seal (Phoca vitulina), Physiol. Zool., 70: 85–92. Hansen, S., Lavigne, D.M., and Innes, S., 1995, Energy metabolism and thermoregulation in juvenile harbor seals (Phoca vitulina) in air, Physiol. Zool., 68: 290–315. Hart, J.S., and Irving, L., 1959, The energetics of harbor seals in air and in water with special consideration of seasonal changes, Can. J. Zool., 37: 447–457. Helm, R.C., 1984, Rate of digestion in three species of pinnipeds, Can. J. Zool., 62: 1751–1756. Hewer, H.R., and Backhouse, K.M., 1968, Embryology and foetal growth of the grey seal, Halichoerus grypus, J. Zool. London, 155: 507–533. Hill, D.A., and Reynolds, J.E., 1989, Gross and microscopic anatomy of the kidney of the West Indian manatee, Trichechus manatus (Mammalia: Sirenia), Acta Anat., 135: 53–56.
0839_frame_C36.fm Page 821 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
821
Hinga, K.R., 1979, The food requirements of whales in the southern hemisphere, Deep Sea Res., 26A: 569–577. Hoch, F., 1971, Energy Transformations in Mammals. Regulatory Mechanisms, W.B. Saunders, Philadelphia, PA, 217 pp. Hong, S.K., Elsner, R., Clayburgh, J.R., and Ronald, K., 1982, Renal functions of the Baikal seal, Pusa sibirica, and ringed seal, Pusa hispida, Physiol. Zool., 55: 289–299. Hui, C.A., 1981, Seawater consumption and water flux in the common dolphin, Delphinus delphis, Physiol. Zool., 54: 430–440. Humphreys, W.F., 1979, Production and respiration in animal populations, J. Anim. Ecol., 48: 427–453. Huntley, A.C., 1984, Relationships between Metabolism, Respiration, Heart Rate and Arousal States in the Northern Elephant Seal, Ph.D. thesis, University of California, Santa Cruz, 193 pp. Huntley, A.C., 1987, Determination of basal metabolic rate by extrapolation in mammals that breathe periodically, in Approaches to Marine Mammal Energetics, Huntley, A.C., Costa, D.P., Worthy, G.A.J., and Castellini, M.A. (Eds.), Society for Marine Mammalogy, Special Publication 1: 97–108. Huntley, A.C., Costa, D.P., and Rubin, R.D., 1984, The contribution of nasal countercurrent heat exchange to water balance in the northern elephant seal, Mirounga angustirostris, J. Exp. Biol., 113: 447–454. Innes, S., 1984, Swimming Energetics, Metabolic Rates and Hind Limb Muscle Anatomy of Some Phocid Seals, Ph.D. thesis, University of Guelph, Ontario, Canada, 198 pp. Innes, S., Lavigne, D.M., Earle, W.M., and Kovacs, K.M., 1986, Feeding rates of seals and whales, J. Anim. Ecol., 56: 115–130. Irvine, A.B., 1983, Manatee metabolism and its influence on distribution in Florida, Biol. Conserv., 25: 315–334. Irving, L., 1973, Aquatic Mammals, in Comparative Physiology of Thermoregulation, Whittow, G.C. (Ed.), Academic Press, New York, 47–96. Irving, L., and Hart, J.S., 1957, The metabolism and insulation of seals as bare skinned mammals in cold water, Can. J. Zool., 35: 497–511. Irving, L., Fisher, K.C., and McIntosh, F.C., 1935, The water balance of a marine mammal, the seal, J. Cell. Comp. Physiol., 6: 387–391. IUPS (International Union of Physiological Sciences) Thermal Commission, 1987, Glossary of terms for thermal physiology, 2nd ed., Pflugers Arch., 410: 567–587. Jangaard, P.M., 1974, The capelin (Mallotus villosus): Biology, distribution, exploitation, utilization, and composition, Bull. Fish. Res. Board Can., 186: 1–70. Jeness, R., 1974, The composition of milk in lactation, in Lactation: A Comprehensive Treatise, Vol. 3, Larson, B.L., and Smith, V.R. (Eds.), Academic Press, New York, 3. Jeness, R., Williams, T.D., and Mullin, R.J., 1981, Composition of milk of the sea otter (Enhydra lutris), Comp. Biochem. Physiol., 70A: 375–379. Kanwisher, J.W., and Ridgway, S.H., 1983, The physiological ecology of whales and porpoises, Sci. Am., 248: 110–120. Kanwisher, J., and Sundnes, G., 1965, Physiology of a small cetacean, Hvalradets Skr., 48: 45–53. Kanwisher, J., and Sundnes, G., 1966, Thermal regulation of cetaceans, in Whales, Dolphins and Porpoises, Norris, K. (Ed.), University of California Press, Berkeley, 397–407. Kastelein, R.A., and Vaughan, N., 1989, Food consumption, body measurements, and weight changes of a female killer whale (Orcinus orca), Aquat. Mammals, 15: 18–21. Kastelein, R.A., McBain, J., Neurohr, B., Mohri, M., Saijo, S., Wakabayashi, I., and Wiepkema, P.R., 1993, The food consumption of Commerson’s dolphins (Cephalorhynchus commersonii), Aquat. Mammals, 19: 99–121. Kastelein, R.A., Ford, J., Berghout, E., Wiepkema, P.R., and van Boxsel, M., 1994, Food consumption, growth, and reproduction of belugas (Delphinapterus leucas) in human care, Aquat. Mammals, 20: 81–97.
0839_frame_C36.fm Page 822 Tuesday, May 22, 2001 2:07 PM
822
CRC Handbook of Marine Mammal Medicine
Kastelein, R.A., Kershaw, J., Berghout, E., and Wiepkema, P.R., 1995, The food consumption of South American sea lions (Otaria flavescens), Aquat. Mammals, 21: 43–53. Kasting, N.W., Adderley, S.A.L., Safford, T., and Hewlett, K.G., 1989, Thermoregulation in beluga (Delphinapterus leucas) and killer (Orcinus orca) whales, Physiol. Zool., 62: 687–701. Kawamura, A., 1975, A consideration of an available source of energy and its cost for locomotion in fin whales with special reference to their seasonal migrations, Sci. Rep. Whales Res. Inst., 27: 61–79. Keiver, K.M., Draper, H.H., and Ronald, K., 1988, Vitamin D metabolism in the hooded seal (Cystophora cristata), J. Nutr., 118: 332–341. Keiver, K.M., Ronald, K., and Beamish, F.W.H., 1984, Metabolizable energy requirements for maintenance and faecal and urinary losses of juvenile harp seals (Phoca groenlandica), Can. J. Zool., 62: 769–776. Kenyon, K., 1969, The sea otter in the eastern Pacific Ocean, U.S. Fish and Wildlife Service, North American Fauna Series 68: 1–352. Kleiber, M., 1932, Body size and metabolism, Hilgardia, 6: 315–353. Kleiber, M., 1975, The Fire of Life: An Introduction to Animal Energetics, R.E. Kreiger, Huntington, NY, 453 pp. Kooyman, G.L., and Drabek, C.M., 1968, Observations on milk, blood and urine constituents of the Weddell seal, Physiol. Zool., 41: 187–194. Kooyman, G.L., Kerem, D.H., Campbell, W.B., and Wright, J.J., 1973, Pulmonary gas exchange in a freely diving Weddell seal, Respir. Physiol., 17: 283–290. Krockenberger, M.B., and Bryden, M.M., 1993, Energy intake of immature southern elephant seals, Mirounga leonina (Carnivora: Phocidae), Aust. J. Zool., 41: 589–597. Lanneck, N., 1973, The importance of vitamin E for domestic animals in sickness and in health, Acta Agric. Scand. Suppl., 19: 1–19. Lavigne, D.M., Barchard, W., Innes, S., and Øritsland, N.A., 1982a, Pinniped bioenergetics, FAO Fish. Serv., 5: 191–235. Lavigne, D.M., Stewart, R.E.A., and Fletcher, F., 1982b, Changes in composition and energy content of harp seal milk during lactation, Physiol. Zool., 55: 1–9. Lavigne, D.M., Innes, S., Worthy, G.A.J., Kovacs, K.M., Schmitz, O.J., and Hickie, J.P., 1986, Metabolic rates of seals and whales, Can. J. Zool., 64: 279–284. Laws, R.M., 1959a, Foetal growth rates of whales with special reference to the fin whale, Balaenoptera physalus (L.), Discovery Rep., 29: 281–308. Laws, R.M., 1959b, Accelerated growth in seals, with special reference to the Phocidae, Nor. HvalfangstTid., 9: 425–452. Lawson, J.W., Hare, J.A., Noseworthy, E., and Friel, J.K., 1997a, Assimilation efficiency of captive ringed seals (Phoca hispida) fed different diets, Polar Biol., 18: 107–111. Lawson, J.W., Miller, E.H., and Noseworthy, E., 1997b, Variation in assimilation efficiency and digestive efficiency of captive harp seals (Phoca groenlandica) on different diets, Can. J. Zool., 75: 1285–1291. Lawson, J.W., Magalhães, A.M., and Miller, E.H., 1998, Important prey species of marine vertebrate predators in the northwest Atlantic: Proximate composition and energy density, Mar. Ecol. Prog. Ser., 164: 13–20. Le Boeuf, B.J., and Condit, R.S., 1983, The high cost of living on the beach, Pac. Discovery, 36: 12–18. Liao, J., 1990, An Investigation on the Effect of Water Temperature on the Metabolic Rate of the California Sea Lion (Zalophus californianus), M.Sc. thesis, University of California, Santa Cruz, 55 pp. Lockley, R.M., 1966, Grey Seal, Common Seal, Andre Deutsch, London, 59 pp. Lockyer, C., 1981, Growth and energy budgets of large baleen whales from the southern hemisphere, FAO Fish. Serv., 5: 379–487. Lockyer, C., 1987, Evaluation of the role of fat reserves in relation to the ecology of north Atlantic fin and sei whales, in Approaches to Marine Mammal Energetics, Huntley, A.C., Costa, D.P., Worthy, G.A.J., and Castellini, M.A. (Eds.), Society for Marine Mammalogy, Special Publication 1: 183–203.
0839_frame_C36.fm Page 823 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
823
Lomolino, M.V., and Ewel, K.C., 1984, Digestive efficiencies of the West Indian manatee (Trichechus manatus), Fla. Sci., 47: 176–179. Loughlin, T.R., Bengston, J.L., and Merrick, R.L., 1987, Characteristics of feeding trips of female northern fur seals, Can. J. Zool., 65: 2079–2084. MacKay, R.S., 1964, Deep body temperature of untethered dolphin recorded by ingested radio transmitter, Science, 144: 864–865. Maluf, N.S.R., 1989, Renal anatomy of the manatee, Trichechus manatus, Linneaus, Am. J. Anat., 184: 269–286. Markussen, N.H., Ryg, M., and Øritsland, N.A., 1990, Energy requirements for maintenance and growth of captive harbour seals, Phoca vitulina, Can. J. Zool., 68: 423–426. Markussen, N.H., Ryg, M., and Øritsland, N.A., 1992, Metabolic rate and body composition of harbour seals, Phoca vitulina, during starvation and refeeding, Can. J. Zool., 70: 220–224. Markussen, N.H., Ryg, M., and Øritsland, N.A., 1994, The effect of feeding on the metabolic rate in harbour seals (Phoca vitulina), J. Comp. Physiol. B, 164: 89–93. Mårtensson, P.-E., Nordøy, E.S., and Blix, A.S., 1994, Digestibility of crustaceans and capelin in harp seals (Phoca groenlandica), Mar. Mammal Sci., 10: 325–331. Mårtensson, P.-E., Lager Gotas, A.R., Nordøy, E.S., and Blix, A.S., 1996, Seasonal changes in energy density of prey of northeast Atlantic seals and whales, Mar. Mammal Sci., 12: 635–640. Mathisen, O.A., Baade, R.T., and Lopp, R.J., 1962, Breeding habits, growth and stomach contents of the Steller sea lion in Alaska, J. Mammal., 43: 469–477. Matsuura, D.T., and Whittow, G.C., 1973, Oxygen uptake of the California sea lion and harbor seal during exposure to heat, Am. J. Physiol., 225: 711–715. Mazzaro, L.M., Dunn, J.L., Furr, H.C., and Clark, R.M., 1995, Study of vitamin A supplementation in captive northern fur seals (Callorhinus ursinus) and its effect on serum vitamin E, Mar. Mammal Sci., 11: 545–553. McGinnis, S.M., Whittow, G.C., Ohata, C.A., and Huber, H., 1972, Body heat dissipation and conservation in two species of dolphins, Comp. Biochem. Physiol., 43A: 417–423. McNab, B.K., 1980, Food habits, energetics and the population biology of mammals, Am. Nat., 116: 106–124. McNab, B.K., 1986, The influence of feeding habits on the energetics of eutherian mammals, Ecol. Monogr., 56: 1–29. Medway, W., and Geraci, J.R., 1978, Clinical pathology of marine mammals, in Zoo and Wild Animal Medicine, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 604–610. Miculka, T.A., and Worthy, G.A.J., 1994, Peripheral tissue composition and its impact on heat loss in the west Indian manatee (Trichechus manatus), presented at First International Manatee and Dugong Research Conference, Gainesville, FL, 11–13 March, 91–92. Miculka, T.A., and Worthy, G.A.J., 1995, Metabolic capabilities and the limits to thermoneutrality in juvenile and adult West Indian manatees (Trichechus manatus), presented at 11th Biennial Conference on the Biology of Marine Mammals, Orlando, FL, 14–18 December, 77. Miller, L.K., 1978, Energetics of the northern fur seal in relation to climate and food resources of the Bering Sea, Final Report to the U.S. Marine Mammal Commission, Report MMC-75/08, 27 pp. Miller, R.M., and Ridgway, S.H., 1963, Clinical experiences with dolphins and whales, Small Anim. Clin., 3: 189–193. Moen, A.N., 1973, Wildlife Ecology: An Analytical Approach, W.H. Freeman, San Francisco, 126 pp. Mooney, E.E., 1981, Unpublished data cited in Perez, M.A., and Mooney, E.E., 1986, Increased food and energy consumption of lactating northern fur seals, Callorhinus ursinus, Fish. Bull., 84: 371–381. Moors, P., 1977, Studies of the metabolism, food composition and assimilation efficiency of a small carnivore, Oecologia, 27: 185–202. Mrosovsky, N., and Sherry, D.F., 1980, Animal anorexias, Science, 207: 837–842.
0839_frame_C36.fm Page 824 Tuesday, May 22, 2001 2:07 PM
824
CRC Handbook of Marine Mammal Medicine
Murie, D.J., 1987, Experimental approaches to stomach content analyses of piscivorous marine mammals, in Approaches to Marine Mammal Energetics, Huntley, A.C., Costa, D.P., Worthy, G.A.J., and Castellini, M.A. (Eds.), Society for Marine Mammalogy, Special Publication 1: 147–164. Myers, B.J., 1955, The rearing of a grey seal in captivity, Can. Field Nat., 69: 151–153. Nordøy, E.S., and Blix, A.S., 1985, Energy sources in fasting grey seal pups evaluated with computed tomography, Am. J. Physiol., 249: R471–R476. Nordøy, E.S., Ingebretsen, O.C., and Blix, A.S., 1990, Depressed metabolism and low protein catabolism in fasting grey seal pups, Acta Physiol. Scand., 139: 361–369. Nordøy, E.S., Aakvaag, A., and Larsen, T.S., 1993a, Metabolic adaptations to fasting in harp seal pups, Physiol. Zool., 66: 926–945. Nordøy, E.S., Sørmo, W., and Blix, A.S., 1993b, In vitro digestibility of different prey species of minke whales (Balaenoptera acutorostrata), Br. J. Nutr., 70: 485–489. NRC (National Research Council), 1981, Nutritional Energetics of Domestic Animals and Glossary of Energy Terms, 2nd ed., National Academy Press, Washington, D.C., 54 pp. Oftedal, O.T., in press, Lactation in whales and dolphins: Evidence of divergence between baleen and toothed species. Oftedal, O.T., Boness, D.J., and Tedman, R.A., 1987, The behavior, physiology, and anatomy of lactation in the Pinnipedia, Current Mammal., 1: 175–245. Oftedal, O.T., Boness, D.J., and Bowen, W.D., 1988, The composition of hooded seal (Cystophora cristata) milk: An adaptation for postnatal fattening, Can. J. Zool., 66: 318–322. Olsen, C.R., Hall, F.C., and Elsner, R., 1969, Mechanics of ventilation in the pilot whale, Respir. Physiol., 7: 7–12. Olsen, M.A., Aagnes, T.H., and Mathiesen, S.D., 1994, Digestion of herring by indigenous bacteria in the minke whale forestomach, Appl. Environ. Microbiol., 60: 4445–4455. Øritsland, N.A., and Ronald, K., 1975, Energetics of the freely diving harp seal (Pagophilus groenlandicus), Rapp. P.-V. Reun. Cons. Int. Explor. Mer, 169: 451–455. Ortiz, C.L., Costa, D.P., and Le Boeuf, B.J., 1978, Water and energy flux in elephant seal pups fasting under natural conditions, Physiol. Zool., 51: 166–178. Ortiz, R.M., 1994, Water Flux and Osmoregulatory Physiology of the West Indian Manatee (Trichechus manatus), M.Sc. thesis, Texas A&M University, College Station, 87 pp. Ortiz, R.M., Worthy, G.A.J., and MacKenzie, D.S., 1998, Osmoregulation in wild and captive West Indian manatees (Trichechus manatus), Physiol. Zool., 71: 449–457. Ortiz, R.M., Worthy, G.A.J., and Byers, F.M., 1999, Estimation of water turnover rates of captive West Indian manatees (Trichechus manatus) held in fresh and salt water, J. Exp. Biol., 202: 33–38. Parry, D.A., 1949, The structure of whale blubber and a discussion of its thermal properties, Q. J. Microbiol. Sci., 90: 13–26. Parsons, J.L., 1977, Metabolic Studies on Ringed Seals (Phoca hispida), M.S. thesis, University of Guelph, Ontario, Canada, 82 pp. Peddemors, V.M., de Muelenaere, H.J.H., and Devchand, K., 1989, Comparative milk composition of the bottlenosed dolphin (Tursiops truncatus), humpback dolphin (Sousa plumbea) and common dolphin (Delphinus delphis) from Southern African waters, Comp. Biochem. Physiol., 94A: 639–641. Perez, M.A., and Bigg, M.A., 1986, Diet of northern fur seals, Callorhinus ursinus, off western North America, Fish. Bull., 84: 957–971. Perez, M.A., and Mooney, E.E., 1986, Increased food and energy consumption of lactating northern fur seals, Callorhinus ursinus, Fish. Bull., 84: 371–381. Pernia, S.D., Ortiz, C.L., and Hill, A., 1978, Urea turnover during prolonged fasting in the northern elephant seal, Comp. Biochem. Physiol., 65B: 731–734. Pervaiz, S., and Brew, K., 1986, Composition of the milks of the bottlenose dolphin (Tursiops truncatus) and the Florida manatee (Trichechus manatus latirostris), Comp. Biochem. Physiol., 84A: 357–360. Petrusewicz, K., and MacFayden, A., 1970, Productivity of terrestrial mammals, in Principles and Methods, International Biological Program Handbook, No. 13, Blackwell Scientific, Oxford, U.K., 190 pp.
0839_frame_C36.fm Page 825 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
825
Pilson, E.E.Q., 1970, Water balance in California sea lions, Physiol. Zool., 43: 257–269. Quiring, D.P., and Harlan, C.F., 1953, On the anatomy of the manatee, J. Mammal., 34: 192–203. Rand, R., 1955, Reproduction in the female Cape fur seal, Arctocephalus pusillus, Proc. Zool. Soc. London, 124: 717–740. Rea, L.D., 1995, Prolonged Fasting in Pinnipeds, Ph.D. thesis, University of Alaska, Fairbanks, 135 pp. Rea, L.D., and Costa, D.P., 1992, Changes in standard metabolism during long-term fasting in northern elephant sea pups (Mirounga angustirostris), Physiol. Zool., 65: 97–111. Reed, J.Z., Chambers, C., Hunter, C.J., Lockyer, C., Kastelein, R., Fedak, M.A., and Boutilier, R.G., 2000, Gas exchange and heart rate in the harbour porpoise, Phocoena phocoena, J. Comp. Physiol. B, 170: 1–10. Reidmann, M., and Ortiz, C.L., 1979, Changes in milk composition during lactation in the northern elephant seal, Physiol. Zool., 52: 240–249. Reilly, J.J., 1991, Adaptations to prolonged fasting in free-living weaned gray seal pups, Am. J. Physiol., 260: R267–R272. Renouf, D., and Noseworthy, E., 1990, Feeding cycles in captive harbour seals (Phoca vitulina): Weight gain in spite of reduced food intake and increased thermal demands, Mar. Behav. Physiol., 17: 203–212. Renouf, D., Noseworthy, E., and Scott, M.C., 1990, Daily fresh water consumption by captive harp seals (Phoca groenlandica), Mar. Mammal Sci., 6: 253–257. Reynolds, J.E., and Wilcox, J.R., 1985, Abundance of West Indian manatees (Trichechus manatus) around selected Florida power plants following winter cold fronts, 1982–1983, Bull. Mar. Sci., 36: 413–422. Rice, D.W., and Wolman, A.A., 1971, The life history and ecology of the grey whale (Eschrichtius robustus), American Society Mammalology, Special Publication No. 3, 142 pp. Ridgway, S.H., 1972, Homeostasis in the aquatic environment, in Mammals of the Sea, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 590–747. Rigdon, R.H., and Dragger, G.A., 1955, Thiamine deficiency in sea lions (Otaria californiana) fed only frozen fish, J. Am. Vet. Med. Assoc., 127: 453–455. Riviere, J.E., Englehardt, F.R., and Solomon, J., 1977, The relationship of thyroxine and cortisol to the moult of the harbour seal, Phoca vitulina, Gen. Comp. Endocrinol., 31: 398–401. Rommel, S.A., Pabst, D.A., McLellan, W.A., Mead, J.G., and Potter, C.W., 1992, Anatomical evidence for a countercurrent heat exchanger associated with dolphin testes, Anat. Rec., 232: 150–156. Rommel, S.A., Early, G.A., Matassa, K.A., Pabst, D.A., and McLellan, W.A., 1995, Venous structures associated with thermoregulation of phocid seal reproductive organs, Anat. Rec., 243: 390–402. Ronald, K., and Thomson, C.A., 1981, Parturition and postpartum behavior of a captive harbour seal, Phoca vitulina, Aquat. Mammals, 8: 79–84. Ronald, K., Johnson, E., Foster, M., and Van der Pol, D., 1969, The harp seal, Pagophilus groenlandicus (Erxleben, 1777). I. Methods of handling, molt, and diseases in captivity, Can. J. Zool., 48: 1035–1040. Ronald, K., Keiver, K.M., Beamish, F.W.H., and Frank, R., 1984, Energy requirements for maintenance and faecal and urinary losses of the grey seal (Halichoerus grypus), Can. J. Zool., 62: 1101–1105. Rosen, D.A.S., and Renouf, D., 1995, Variation in the metabolic rates of captive harbour seals, in Whales, Seals, Fish, and Man, Blix, A.S., Walløe, L., and Ulltang, Ø. (Eds.), Elsevier Science, Amsterdam, the Netherlands, 393–399. Rosen, D.A.S., and Trites, A.W., 1997, Heat increment of feeding in Steller sea lions, Eumetopias jubatus, Comp. Biochem. Physiol., 118A: 877–881. Rosen, D.A.S., and Trites, A.W., 1999, Metabolic effects of low-energy diet on Steller sea lions, Eumetopias jubatus, Physiol. Biochem. Zool., 72: 723–731. St. Aubin, D.J., and Geraci, J.R., 1980, Tissue levels of ascorbic acid in marine mammals, Comp. Biochem. Physiol., 66A: 605–609. Sandegren, F.E., 1970, Breeding and Maternal Behavior of the Steller Sea Lion (Eumetopias jubatus) in Alaska, M.Sc. thesis, University of Alaska, Fairbanks, 138 pp.
0839_frame_C36.fm Page 826 Tuesday, May 22, 2001 2:07 PM
826
CRC Handbook of Marine Mammal Medicine
Scheffer, V.B., 1962, Pelage and surface topography of the northern fur seal, U.S. Fish and Wildlife Service, North American Fauna Series, 64: 1–206. Schmidt-Nielsen, K., 1997, Animal Physiology, 5th ed., Cambridge University Press, London, 607 pp. Scholander, P.F., 1940, Experimental investigations on the respiratory function in diving mammals and birds, Hvalradets Skr., 22: 1–131. Scholander, P.F., and Irving, L., 1941, Experimental investigations on the respiration and diving of the Florida manatee, J. Cell. Comp. Physiol., 17: 169–191. Scholander, P.F., Irving, L., and Grinnell, S.W., 1942, On the temperature and metabolism of the seal during diving, J. Cell. Comp. Physiol., 19: 67–78. Sergeant, D.E., 1969, Feeding rates of Cetacea, Fiskeridir. Skr. Ser. Havunders., 15: 246–258. Sergeant, D.E., 1976, History and present status of harp and hooded seals, Biol. Conserv., 10: 95–117. Shapunov, V.M., 1973, Evaluation of the economy and effectiveness of external respiration in the dolphin, Phocoena phocoena, J. Biochem. Biophysiol., 7: 331–336. Sibly, R.M., and Calow, P., 1987, Physiological Ecology of Animals, Blackwell Scientific, Palo Alto, CA, 179 pp. Snyder, G.K., 1983, Respiratory adaptations in diving mammals, Respir. Physiol., 54: 269–294. South, F.E., Luecke, R.H., Zatzman, M.L., and Shanklin, M.D., 1976, Air temperature and direct partitional calorimetry of the California sea lion (Zalophus californianus), Comp. Biochem. Physiol., 54: 27–30. Spotte, S., 1982, The incidence of twins in pinnipeds, Can. J. Zool., 60: 2226–2233. Spotte, S., and Babus, B., 1980, Does a pregnant dolphin (Tursiops truncatus) eat more? Cetology, 39: 1–7. Stewart, R.E.A., and Lavigne, D.M., 1980, Neonatal growth in northwest Atlantic harp seals (Pagophilus groenlandicus), J. Mammal., 60: 670–680. Stewart, R.E.A., and Lavigne, D.M., 1984, Energy transfer and female condition in nursing harp seals, Phoca groenlandica, Holarctic Ecol., 7: 182–194. Stewart, R.E.A., Webb, B.E., and Lavigne, D.M., 1983, Determining lactose content of harp seal milk, Can. J. Zool., 61: 1094–1100. Stoddard, J.H., 1988, Fat contents of Canadian Atlantic herring, Technical Report, Fish. Res. Brd. Can., 79: 1–23. Sugii, K., and Kinumaki, T., 1968, Distribution of vitamin E in a few species of fish, Bull. Jpn. Soc. Fish., 34: 420–428. Tandler, A., and Beamish, F.W.H., 1979, Mechanical and biochemical components of apparent specific dynamic action in largemouth bass, Micropterus salmoides Lacepede, J. Fish. Biol., 14: 343–351. Tarasoff, F., and Toews, D., 1972, The osmotic and ionic regulatory capabilities of the kidney of the harbor seal, Phoca vitulina, J. Comp. Physiol., 81: 121–132. Taylor, C.R., Heglund, N.C., and Maloiy, G.M.D., 1982, Energetics and mechanics of terrestrial locomotion. I. Metabolic energy consumption as a function of speed and body size in birds and mammals, J. Exp. Biol., 97: 1–21. Telfer, N., Cornell, L.H., and Prescott, J.H., 1975, Do dolphins drink water? J. Am. Vet. Med. Assoc., 157: 555–558. Thurston, C.E., 1958, Sodium and potassium content of 34 species of fish, J. Am. Diet. Assoc., 34: 396–399. Trillmich, F., and Lechner, E., 1986, Milk of the Galapagos fur seal and sea lion with a comparison of the milk of eared seals (Otariidae), J. Zool. London, 209: 271–277. Webster, A.J., 1976, Efficiencies of energy utilization during growth, in Meat Animals: Growth and Productivity, Lister, D., Rhodes, D.N., Fowler, V.R., and Fuller, M.F. (Eds.), Plenum Press, New York, 234 pp. White, J.R., 1970, Thiamine deficiency in an Atlantic bottle-nosed dolphin (Tursiops truncatus) on a diet of raw fish, J. Am. Vet. Med. Assoc., 157: 559–562. Whittow, G.C., Matsuura, D.T., and Lin, Y.C., 1972, Temperature regulation in the California sea lion (Zalophus californianus), Physiol. Zool., 45: 68–77. Williams, T.M., 1999, The evolution of cost efficient swimming in marine mammals: Limits to energetic optimization, J. Am. Vet. Med. Assoc., 354: 193–201.
0839_frame_C36.fm Page 827 Tuesday, May 22, 2001 2:07 PM
Nutrition and Energetics
827
Williams, T.M., and Kooyman, G.L., 1985, Swimming performance and hydrodynamic characteristics of harbor seals, Phoca vitulina, Physiol. Zool., 58: 576–589. Williams, T.M., Haun, J.E., Friedl, W.A., Hall, R.W., and Bivens, L.W., 1992a, Assessing the thermal limits of bottlenose dolphins: A cooperative study by trainers, scientists and animals, Int. Mar. Anim. Trainers Assoc. (IMATA) Soundings, Fall: 16–17. Williams, T.M., Friedl, W.A., Fong, M.L., Yamada, R.M., Sedivy, P., and Haun, J.E., 1992b, Travel at low energetic cost by swimming and wave-riding bottlenose dolphins, Nature, 355: 821–823. Williams, T.M., Friedl, W.A., and Haun, J.E., 1993, The physiology of bottlenose dolphins (Tursiops truncatus): Heart rate, metabolic rate and plasma lactate concentration before exercise, J. Exp. Biol., 179: 31–46. Wolf, A.V., Prentiss, G.L., Douglass, L.G., and Swett, R.S., 1959, Potability of sea water with special reference to the cat, Am. J. Physiol., 196: 663–641. Worthy, G.A.J., 1985, Thermoregulation of Young Phocid Seals, Ph.D. thesis, University of Guelph, Ontario, Canada, 254 pp. Worthy, G.A.J., 1987, Metabolism and growth of young harp and grey seals, Can. J. Zool., 65: 1377–1382. Worthy, G.A.J., 1991, Thermoregulatory implications of the interspecific variation in blubber composition of odontocete cetaceans, presented at 9th Biennial Conference on the Biology of Marine Mammals, Chicago, IL, 5–9 December, 95. Worthy, G.A.J., and Edwards, E.F., 1990, Morphometric and biochemical factors affecting heat loss in a small temperate cetacean (Phocoena phocoena) and a small tropical cetacean (Stenella attenuata), Physiol. Zool., 63: 432–442. Worthy, G.A.J., and Lavigne, D.M., 1982, Changes in blood properties of fasting and feeding harp seal pups, Phoca groenlandica, after weaning, Can. J. Zool., 60: 586–592. Worthy, G.A.J., and Lavigne, D.M., 1983, Energetics of fasting and subsequent growth in weaned harp seal pups, Phoca groenlandica, Can. J. Zool., 61: 447–456. Worthy, G.A.J., and Lavigne, D.M., 1987, Mass loss, metabolic rate, and energy utilization by harp and grey seal pups during the postweaning fast, Physiol. Zool., 60: 352–364. Worthy, G.A.J., Innes, S., Braune, B.M., and Stewart, R.E.A., 1987, Rapid acclimation of cetaceans to an open-system respirometer, in Approaches to Marine Mammal Energetics, Huntley, A.C., Costa, D.P., Worthy, G.A.J., and Castellini, M.A. (Eds.), Society for Marine Mammalogy, Special Publication 1: 115–112. Worthy, G.A.J., Morris, P.A., Costa, D.P., and Le Boeuf, B.J., 1992, Moult energetics of the northern elephant seal (Mirounga angustirostris), J. Zool. London, 227: 257–265. Yasui, W.Y., and Gaskin, D.E., 1986, Energy budget of a small cetacean, the harbour porpoise, Phocoena phocoena (L.), Ophelia, 25: 183–197.
0839_frame_C36.fm Page 828 Tuesday, May 22, 2001 2:07 PM
0839_frame_37.fm Page 829 Tuesday, May 22, 2001 1:47 PM
37 Hand-Rearing and Artificial Milk Formulas Forrest I. Townsend, Jr. and Laurie J. Gage
Introduction Methods for hand-rearing marine mammals have improved over the last decade, especially for cetaceans (Townsend, 1999). Species from common dolphins (Delphinus delphis) to gray whales (Eschrichtius robustus) have been successfully hand-reared on artificial formulas. With some of the more commonly hand-reared species, such as harbor seals (Phoca vitulina), comparative studies of growth following use of different formulas have been conducted (Sanderson, 1999). In species infrequently hand-reared, such as cetaceans and polar bears (Ursus maritimus), data are gathered from the successful, usually individual, case reports. In recent years, detailed analyses of formulas have been utilized to develop optimal diets for each species. This chapter includes suggestions for formula selection and preparation, practical tips for hand rearing, and advice on weaning procedures to help facilitate a successful outcome. However, for each species, the reader is advised to seek advice from colleagues with experience hand-rearing the species of concern, since hand-rearing is as much an art as a science.
Cetaceans Formula Herring filets plus viscera Zoologic® Milk Matrix 30/55 (Pet-Ag, Inc., Hampshire, IL) Salmon, sardine, menhaden, or safflower oil Lecithin Lactobacillus tablets Osteoform powder (Veta-mix, Shenandoah, IA) Multiple vitamin with zinc Taurine Bottled water
750 ml 2.5 cups 50 ml 1 tbsp 3 1 tbsp 1 capsule 250 mg 1100 ml
Blend herring filets and viscera in a commercial blender. Add bottled water. Blend in milk substitute. Add salmon oil, Osteoform powder, lecithin, lactobacillus, taurine, and vitamin supplement, blending to mix well. Label formula container with date and time of preparation, and refrigerate until used. Warm the formula to 36°C (97°F) before feeding. Discard unused formula after 24 hours. 0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
829
0839_frame_37.fm Page 830 Wednesday, May 23, 2001 11:05 AM
830
CRC Handbook of Marine Mammal Medicine
Delivery Methods and Techniques If the calf will accept a nipple and bottle, and nurse on its own, this is the preferred method, because both labor and the possibility of aspiration pneumonia are reduced. Nipple selection is critical; a lamb’s nipple is a good choice for bottlenose dolphin (Tursiops truncatus) and harbor porpoise (Phocoena phocoena) calves. The hole in the nipple should be large enough to allow formula to be squeezed from the bottle, and depends upon the consistency of the formula, which can be adjusted. Nipples may also be made from pieces of rubber tubing, and gradually modified as the suckling behavior of the calf improves. Initially, start working with the calf in waist-deep water, with one handler holding the calf and another handler placing a hand under the calf ’s lower jaw for support. Lift the rostrum slightly out of the water allowing only formula to be ingested and not seawater. Squeeze the bottle to assist the calf ’s own suckling. With this method, several ounces of formula can be delivered in a short time. Feeding may be messy at first, but soon the calf will need no support and can nurse satisfactorily at the side of the pool. Control the feeding time, allowing 10 to 15 min for nursing. Calves may get very playful and forget it is time to eat. Remove any pool toys prior to feeding; then return toys and give tactile reinforcement after successful nursing. Another method found useful when hand-raising harbor porpoises is the tube-feeding technique. Use a soft rubber stomach tube (1 cm diameter) with a rounded tip and a hole near the end. With bottlenose dolphin calves (or other small cetaceans), a slightly larger foal-feeding tube is utilized. The tube is marked at a length equivalent to the distance from the tip of the rostrum to the leading edge of the dorsal fin. This distance will allow the tube tip to be placed in the forestomach of the porpoise. Insert the lubricated stomach tube to the stomach (marked length) and use a funnel to gravity-feed formula slowly (or a large syringe for harbor porpoises). This allows for expansion of the forestomach. Administering formula too rapidly may result in vomiting and possible aspiration pneumonia.
Feeding Frequency and Daily Requirements In display facilities, dolphin calves have been observed nursing for brief periods two to three times per hour. This schedule is impractical in a hand-rearing situation. Feeding frequency depends upon caloric density of the formula, daily caloric requirements, and maximum volume of formula per feeding (see Chapter 36, Nutrition). For the bottlenose dolphin calf nursing from a bottle, hourly feeding is not unreasonable. However, tube-feeding the same calf would be labor intensive, impractical, and stressful to the animal. It is important to know the approximate caloric density of the formula being used. This information, combined with knowledge of the caloric requirements for the species, allows the total daily volume of formula to be calculated. The total daily volume is divided by maximum volume per feeding that can be safely given to a calf to determine the estimated daily number of feedings. This calculation is only a guideline, and the actual daily formula required will ultimately depend on weight gain (or loss). There are situations in which daily weighing is not feasible. In these cases, careful measurements of the calf ’s girth at the anterior edge of the dorsal fin will give useful information on the trends in body weight. Guidelines to caloric requirements for some species are as follows: Bottlenose dolphin (Tursiops truncatus) Spotted dolphin (Stenella attenuata) Pygmy sperm whale (Kogia breviceps) Harbor porpoise (Phocoena phocoena) Killer whale (Orcinus orca) Common dolphin (Delphinus delphis)
150 kcal/kg/day 200 kcal/kg/day 80 kcal/kg/day 200 kcal/kg/day 125 kcal/kg/day 150 kcal/kg/day
0839_frame_37.fm Page 831 Tuesday, May 22, 2001 1:47 PM
Hand-Rearing and Artificial Milk Formulas
831
Monitoring Neonates Initial blood tests should include compete blood cell counts, serum chemistries, and electrolytes. Twice weekly blood sampling is recommended until the calf is medically stable. Weigh the calf daily for 2 weeks, then every other day up to 2 months of age. After this time, weekly weighing is recommended. Target weight gain for a bottlenose dolphin should be 0.25 kg/day. In smaller species (Stenella spp. and Delphinus spp.) expect a weight gain of approximately 0.125 kg/day, and in larger species (Pseudorca spp. and Kogia spp.) 0.5 kg/day should be the daily goal (Townsend, 1999).
Weaning Procedures There are several ways to wean a calf. The introduction of live fish has in the past stimulated hunting behavior in harbor porpoise calves (Spotte et al., 1978; Kastelein et al., 1990). Small dead fish may also be offered. Calves will often play with the fish, carrying them around and sometimes shredding them. The presence of a surrogate mother or other pool mates may initiate the transition to eating fish. Start offering fish at 10 weeks of age and gradually wean off formula by 8 months of age for bottle-fed calves. If tube-feeding, try to wean at an earlier age. Initiate the weaning process by force-feeding at 4 months of age.
Other Practical Information In display situations, it is advisable to collect and store frozen dolphin milk for hand rearing. Frozen milk may be stored for a year after collection. Use this milk to supplement the formula for the first few days (Sweeney, pers. comm.). The use of immunoglobulins has been recommended (Dalton et al., 1993). Dolphin IgG given parenterally at 1 mg/kg body weight is approximately equal to 10 µg of IgG/ml of the calf ’s blood volume. This treatment may be repeated every 2 to 3 weeks (McBain, 2000).
References and Suggested Further Reading Ackman, R.G., Eaton, C.A., and Hooper, S.N., 1968, Lipids of the fin whale (Balaenoptera physalus) from North Atlantic waters, IV, Fin whale milk, Can. J. Biochem., 46: 197–203. Ackman, R.G., Eaton, C.A., and Mitchell, E.D., 1971, The bottle-nosed dolphin Tursiops truncatus: Fatty acid composition of milk triglyceride, Can. J. Biochem., 49: 1172–1174. Dalton, L.M., Schwertner, H.A., and McBain, J.F., 1993, The use of immunoglobulin concentrate in a beluga whale calf, in Proceedings of the 24th International Association for Aquatic Animal Medicine, Chicago, IL, 110. Harms, C.A., 1993, Composition of prepartum mammary secretions of two bowhead whales (Balaena mysticetus L.), J. Wildl. Dis., 29: 94–97. Kastelein, R.A., Bakker, M.J., and Dokter, T., 1990, The medical treatment of three stranded harbour porpoises (Phocoena phocoena), Aquat. Mammals, 15: 181–202. Kastelein, R.A., Schooneman, N.M., Staal, C., and Boer, H., 1997, A method for tube-feeding juvenile harbour porpoises (Phocoena phocoena), in The Biology of the Harbour Porpoise, Read, A.J., Wiepkema, P.R., and Nachtigall, P.E. (Eds.), De Spil Publishers, Woerden, the Netherlands, 63–83. Lauer, B.H., and Baker, B.E., 1969, Whale milk, I. Fin whale (Balaenoptera physalus) and beluga whale (Delphinapterus leucas) milk: Gross composition and fatty acid constitution, Can. J. Zool., 47: 95–97. McBain, J.F., 2000, Early neonatal diseases and therapeutics in bottlenose dolphins, including a brief discussion of obstetrical manipulations, Bottlenose Dolphin Reproduction Workshop Report, AZA Marine Mammal Taxonomy Advisory Group, Silver Spring, MD, 317. Oftedal, O.T., 1993, The adaptation of milk secretion to the constraints of fasting in bears, seals, and baleen whales, J. Dairy Sci., 76: 3234–3246.
0839_frame_37.fm Page 832 Tuesday, May 22, 2001 1:47 PM
832
CRC Handbook of Marine Mammal Medicine
Peddemors, V.M., de Muelenaere, H.J., and Devchand, K., 1989, Comparative milk composition of the bottlenosed dolphin (Tursiops truncatus), humpback dolphin (Sousa plumbea) and common dolphin (Delphinus delphis) from southern African waters, Comp. Biochem. Physiol. A, 94: 639–641. Pervaiz, S., and Brew, K., 1986a, Purification and characterization of the major whey proteins from the milks of the bottlenose dolphin (Tursiops truncatus), the Florida manatee (Trichechus manatus latirostris), and the beagle (Canis familiaris), Arch. Biochem. Biophys., 246: 846–854. Pervaiz, S., and Brew, K., 1986b, Composition of the milks of the bottlenose dolphin (Tursiops trucatus) and the Florida manatee (Trichechus manatus latirostris), Comp. Biochem. Physiol. A, 84: 357–360. Pilson, M.E., and Waller, D.W., 1970, Composition of milk from spotted and spinner porpoises, J. Mammal., 51: 74–79. Reidarson, T.H., Innis, S., Dyer, R., and McBain, J., 2000, Ingredients of an “ideal” cetacean milk formula, Bottlenose Dolphin Reproduction Workshop Report, AZA Marine Mammal Taxon Advisory Group, Silver Spring, MD, 297–310. Reidarson, T.H., McBain, J., and Yochem, P., in press, Medical and nutritional aspects of a California gray whale rehabilitation, Aquat. Mammals. Ridgway, S., Kamolinck, T., Reddy, M., and Curry, C., 1995, Orphan-induced lactation in (Tursiops) and analysis of collected milk, Mar. Mammal Sci., 11: 172–182. Sanderson, S., 1999, Evaluation of two widely used milk replacers for the rearing of orphaned grey seals (Halichoerus grypus) and harbor seals (Phoca vitulina) with a view to determining the limiting factor to growth in rehabilitation, Master’s Degree Project Report, University of London. Shaw, D.H., 1971, Neutral carbohydrates in the milk of the bottlenose dolphin (Tursiops truncatus), Carbohydr. Res., 19: 419–422. Spotte, S., Dunn, J.L., Kezer, L.E., and Heard, F.M., 1978, Notes on the care of a beach-stranded harbour porpoise (Phocoena phocoena), Cetology, 32: 1–5. Townsend, F., 1999, Hand-rearing techniques for neonate cetaceans, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 493–497. Ullrey, D.E., Schwartz, C.C., Whetter, P.A., Rajeshwar Rao, T., Euber, J.R., Cheng, S.G., and Brunner, J.R., 1984, Blue-green color and composition of Stejneger’s beaked whale (Mesoplodon stejnegeri) milk, Comp. Biochem. Physiol. B, 79: 349–352. Walsh, M.T., and Quinton, R.R., 1995, Taurine levels in cetaceans, a preliminary investigation, in Proceedings of the International Association for Aquatic Animal Medicine, Mystic, CT, 115.
Pinnipeds Harbor Seals Harbor seals are the most common species of phocid to be hand-reared, yet relatively little has been published about the composition of harbor seal milk (Boness and Bowen, 1996). Rearing of other species is less well documented, and although it is likely that similar methods would be successful, readers are referred to further literature on the biology of these species.
Formula (Harbor Seal Formula, HSF) Zoologic® Milk Matix 30/55 (Pet-Ag, Inc., Hampshire, IL) Filtered water Fish oil (salmon oil, menhaden oil) Lecithin granules Pinniped multivitamin (Mazuri®, Purina Mills, Inc., St. Louis, MO)
450 ml (dry) 450 ml 350 ml 1 tsp 1 tablet
Do not use a blender to mix this formula. Scoop dry ingredients into a large bowl, add water, and blend slowly with a wire whisk until powder is dissolved. Mix in oil and lecithin granules.
0839_frame_37.fm Page 833 Tuesday, May 22, 2001 1:47 PM
833
Hand-Rearing and Artificial Milk Formulas
Be careful not to overmix the formula, as it tends to thicken and turn pasty. The formula should be smooth with no lumps, ideally the consistency of pancake batter. Warm to 25 to 30°C (77 to 86°F) prior to feeding. Although the formula can be kept refrigerated and used for 24 hours, this is not recommended as the formula tends to thicken over time and may not pass through the stomach tube. It is simpler and less aggravating to make the amount required just prior to feeding.
Delivery Methods and Techniques A clear vinyl stomach tube (1 cm outside diameter) is preferred. Estimate the length of tubing required by measuring from the animal’s snout to the last rib and clearly mark this distance on the tube for future reference. Pass the tube to this depth each time. A 400-ml dose syringe or several 60- or 140-ml catheter-tipped syringes will need to be prefilled with the appropriate mixture and used to deliver the formula slowly. Bottles with lamb or human baby nipples have been used successfully, but are more labor intensive, and tend to result in pups being more used to human handling; thus, the choice of method will be influenced by plans for the pup’s future release possibility.
Feeding Frequency and Daily Requirements The tube-fed pups are fed approximately every 4 hours. The following schedule is used to rehydrate the pups and work them up slowly from an electrolyte solution to full-strength formula. Electrolyte formula (EL) may be any balanced electrolyte solution, such as Pedialyte® (Ross Products Division, Abbott Laboratories, Columbus, OH), or lactated Ringer’s.
Weight at Admission <6 kg (<13 lb)
6–7 kg (13–15 lb)
7–8.5 kg (15–19 lb)
>8.5 kg (>19 lb)
Formula Mixture 150 ml EL 110 ml EL + 40 ml HSF 75 ml EL + 75 ml HSF 40 ml EL + 110 ml HSF 150 ml HSF 170 ml EL 125 ml EL + 45 ml HSF 85 ml EL + 85 ml HSF 45 ml EL + 125 ml HSF 170 ml HSF 190 ml EL 140 ml EL + 50 ml HSF 95 ml EL + 95 ml HSF 50 ml EL + 140 ml HSF 190 ml HSF 220 ml EL 165 ml EL + 55 ml HSF 110 ml EL + 110 ml HSF 55 ml EL + 165 ml HSF 220 ml HSF
Frequency/Number of Feedings q4h × 3 tubings q4h × 1 tubing q4h × 2 tubings q4h × 2 tubings q4h q4h × 3 tubings q4h × 1 tubing q4h × 2 tubings q4h × 2 tubings q4h q4h × 3 tubings q4h × 1 tubing q4h × 2 tubings q4h × 2 tubings q4h q4h × 3 tubings q4h × 1 tubing q4h × 2 tubings q4h × 2 tubings q4h
Once the pups have been fed full-strength formula for 24 hours, the volume may be increased by 20 to 25 ml each day.
0839_frame_37.fm Page 834 Tuesday, May 22, 2001 1:47 PM
834
CRC Handbook of Marine Mammal Medicine
Weaning Procedures Once a pup has reached 1 month of age, is clinically healthy, gaining satisfactory weight, and the teeth have erupted, it is time to introduce fish to the diet. Initially, the pup is shown a fish by placing the pup into a pool and offering the fish held with forceps. The preferred fish for pups is small herring, but smelt may be used. Smaller fish are preferred for the fish introduction process whenever available. Floating fish in the water or offering live fish are other alternatives to fish introduction. This is repeated on a daily basis until the pup exhibits a response. Fish pieces and/or whole fish may be offered at this point. Harbor seals tend to start eating fish with little encouragement, although swallowing fish may be difficult, if the fish offered is too large. Pups tend to gnaw and chew larger fish prior to actually swallowing. Force feeding is necessary if the pup has not responded to the above methods of offering fish. Force feeding involves restraining the pup, placing the fish in the animal’s mouth, and assisting the swallowing reflex by gently pushing the fish past the gag point. It is best to use long, slender fish, preferably herring. The fish should be firm, but not frozen. Dipping the fish in water before placing into the mouth is helpful. During the force-feeding process, the number of stomach tubings is decreased to twice daily. Animals that are still not eating after attempting force feeds may be fasted to stimulate appetite.
Other Practical Information Upon admission, each pup is given a physical examination, which includes auscultation of heart and lungs, weight, standard length, and sex determination. Pups with lanugo coats have a patch of hair clipped over the venipuncture site and around the umbilicus to reduce the risk of contamination at the sites. The presence of an umbilical cord and/or icteric gums should be noted and the umbilicus treated with topical povidone iodine or 0.5% chlorhexidine solution. Blood is drawn every other week. Pups that show clinical illness or abnormalities in serum chemistry or hematological values have additional blood drawn as deemed necessary. Most stranded harbor seal pups hand-reared during rehabilitation have not suckled from their dams and are very young, so tend to have low leukocyte counts (Gulland, pers. comm.). Strict hygiene is recommended, until the pup’s leukocyte counts start to increase as they mature (Ross et al., 1993; 1994). All pups are weighed twice a week, or more frequently if medical problems are noted. Neonatal pups are often hyperbilirubinemic with serum levels up to 16 mg/dl (Gage, 1990). This is presumed due to lysis of fetal red blood cells, and usually resolves without specific therapy. Hypoglycemia in young pups is a common problem. Adding 5 ml of 50% dextrose to each feeding during the first week may help prevent this problem (Gage, 1990). Although access to water at all times is beneficial to older pups, access may be restricted for pups with lanugo coats and those that are severely emaciated. Provision of child-safe heating pads for these pups to lie on further reduces calorie consumption. If the harbor seal pup is to be rehabilitated for release, keep handling and human interaction to a minimum. The recommended release weight is 35 kg (77 lb) (Sanderson, 1999), but pups may be released at 20 kg (44 lb), if the girth is adequate and body shape is more stout than long (Gulland, pers. comm.).
References and Suggested Further Reading Arnould, J.P.Y., Boyd, I.L., and Clarke, A., 1995, A simplified method for determining the gross chemical composition of pinniped milk samples, Can. J. Zool., 73: 404–410. Baker, J.R., 1990, Grey seal (Halichoerus grypus) milk composition and its variation over lactation, Br. Vet. J., 146: 233–238. Boness, D.J., and Bowen, W.D., 1996, The evolution of maternal care in pinnipeds, BioScience, 46: 645–654.
0839_frame_37.fm Page 835 Tuesday, May 22, 2001 1:47 PM
Hand-Rearing and Artificial Milk Formulas
835
Davis, T.A., Nguyen, H.V., Costa, D.P., and Reeds, P.J., 1995, Amino acid composition of pinniped milk, Comp. Biochem. Physiol. B, 110: 633–639. Gage, L.J., 1990, Hand-rearing pinniped pups, in Handbook of Marine Mammal Medicine, Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 533–535. Gage, L.J., 1993, Hand rearing pinnipeds, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 413–415. Hindell, M.A., Bryden, M.M., and Burton, H.R., 1994, Early growth and milk composition in southern elephant seals (Mirounga leonina), Aust. J. Zool., 42: 723–732. Iverson, S.J., Hamosh, M., and Bowen, W.D., 1995a, Lipoprotein lipase activity and its relationship to high milk fat transfer during lactation in grey seals, J. Comp. Physiol. B, 165: 384–395. Iverson, S.J., Oftedal, O.T., Bowen, W.D., Boness, D.J., and Sampugna, J., 1995b, Prenatal and postnatal transfer of fatty acids from mother to pup in the hooded seal, J. Comp. Physiol. B, 165: 1–12. Kovacs, K.M., and Lavigne, D.M., 1986, Maternal investment and neonatal growth in phocid seals, J. Anim. Ecol., 55: 1035–1051. Lavigne, D., Stewart, R.E.A., and Fletcher, F., 1982, Changes in composition and energy content of harp seal milk during lactation, Physiol. Zool., 55: 1. Lydersen, C., and Hammill, M.O., 1993, Activity, milk intake and energy consumption in free-living ringed seal (Phoca hispida) pups, J. Comp. Physiol. B, 163: 433–438. Lydersen, C., and Kovacs, K.M., 1996, Energetics of lactation in harp seals (Phoca groenlandica) from the Gulf of St. Lawrence, Canada, J. Comp. Physiol. B, 166: 295–304. Lydersen, C., Hammill, M.O., and Kovacs, K.M., 1995, Milk intake, growth and energy consumption in pups of ice-breeding grey seals (Halichoerus grypus) from the Gulf of St. Lawrence, Canada, J. Comp. Physiol. B, 164: 585–592. Lydersen, C., Kovacs, K.M., Hammill, M.O., and Gjertz, I., 1996, Energy intake and utilisation by nursing bearded seal (Erignathus barbatus) pups from Svalbard, Norway, J. Comp. Physiol. B, 166: 405–411. Lydersen, C., Kovacs, K.M., and Hammill, M.O., 1997, Energetics during nursing and early postweaning fasting in hooded seal (Cystophora cristata) pups from the Gulf of St. Lawrence, Canada, J. Comp. Physiol. B, 167: 81–88. Messer, M., Crisp, E.A., and Newgrain, K., 1988, Studies on the carbohydrate content of milk of the crabeater seal (Lobodon carcinophagus), Comp. Biochem. Physiol., 90: 367–370. Nordoy, E.S., and Blix, A.S., 1985, Energy sources in fasting grey seal pups evaluated with computed tomography, Am. J. Physiol., 249: R471–R476. Nordoy, E.S., Ingebretsen, O.C., and Blix, A.S., 1990, Depressed metabolism and low protein catabolism in fasting grey seal pups, Acta Physiol. Scand., 139: 361–369. Oftedal, O.T., Boness, D.J., and Bowen, W.D., 1988, The composition of hooded seal (Cystophora cristata) milk: An adaptation for postnatal fattening, Can. J. Zool., 66: 318–322. Reilly, J.J., 1991, Adaptations to prolonged fasting in free-living weaned gray seal pups, Am. J. Physiol., 260: R267–R272. Ross, P.S., Pohajdak, B., Bowen, W.D., and Addison, R.F., 1993, Immune function in free-ranging harbor seal (Phoca vitulina) mothers and their pups during lactation, J. Wildl. Dis., 29: 21–29. Ross, P.S., de Swart, R.L., Visser, I.K., Vedder, L.J., Murk, W., Bowen, W.D., and Osterhaus, A.D., 1994, Relative immunocompetence of the newborn harbour seal, Phoca vitulina, Vet. Immunol. Immunopathol., 2: 331–348. Ryg, M., and Oritsland, N.A., 1991, Estimates of energy expenditure and energy consumption of ringed seals (Phoca hispida) throughout the year, Polar Res., 10: 595–601. Sanderson, S., 1999, Evaluation of two widely used milk replaces for the rearing of orphaned grey seals (Halichoerus grypus) and harbor seals (Phoca vitrulina) with a view to determining the limiting factor to growth in rehabilitation, Master’s degree project report, University of London. Schweigert, F.J., 1993a, Effects of energy mobilization during fasting and lactation on plasma metabolites in the grey seal (Halichoerus grypus), Comp. Biochem. Physiol., 105: 347–352.
0839_frame_37.fm Page 836 Tuesday, May 22, 2001 1:47 PM
836
CRC Handbook of Marine Mammal Medicine
Schweigert, F.J., 1993b, Effects of fasting and lactation on blood chemistry and urine composition in the grey seal (Halichoerus grypus), Comp. Biochem. Physiol., 105: 353–357. Tedman, R.A., 1980, Lactation in the Weddell Seal, Leptonychotes weddellii, Ph.D. dissertation, University of Queensland, Australia. Tedman, R.A., and Green, B., 1987, Water and sodium fluxes and lactational energetics in suckling pups of Weddell seals (Leptonychotes weddellii), J. Zool., 212: 2942. Van Horn, D.R., and Baker, B.E., 1971, Seal milk. II. Harp seal (Pagophilus groenlandicus) milk: Effects of stage of lactation on the composition of the milk, Can. J. Zool., 49: 1085–1088. Worthy, G.A., 1991, Insulation and thermal balance of fasting harp and grey seal pups, Comp. Biochem. Physiol., 100: 845–851.
Elephant Seals Very young northern elephant seals (Mirounga angustirostris) with lanugo coats (“blackcoats”) are reared on harbor seal formula (HSF, above). Older pups that are weaned, but have not started to feed on live prey in the wild, are reared on a different formula (elephant seal formula, ESF). However, as elephant seals are adapted to prolonged periods of fasting, the physiological and biochemical sequences that occur in healthy wild animals may be difficult to mimic when hand-rearing pups (Ortiz et al., 1978; 1984; Riedman, 1990). Many pups strand as emaciated weaned pups, and are rehabilitated by increasing their body weights to a level similar to those of their cohorts prior to release. Thus, methods used to hand-rear these pups are very different from the natural sequence of nutritional events for this species.
Formulas Fish Mash (FM; used with ESF 50–50 and ESF 75–25) Fish pieces (preferably herring) 1 kg Bottled or filtered water 450 ml
Grind fish and mix with water in a blender until smooth. Elephant Seal Formula (ESF) Ground fish (preferably herring) Water Fish oil (salmon or menhaden oil) Lecithin Whipping cream
0.75 kg 225 ml 5 ml 1 tablet 250 ml
Note: The whipping cream must be treated with the 0.75 ml lactase 24 hours prior to use. Grind whole fish in a meat grinder. Place ground fish, water, oil, and lecithin into blender. Blend on low for 10 to 20 s. When ingredients are thoroughly mixed, pour into large bowl. Do not mix cream into the formula in the blender; as the formula becomes too thick. Fold the whipping cream into the formula with a spatula. ESF 50–50 Fish mash Elephant Seal Formula Fish oil
50% volume 50% volume 10 ml
Mix ingredients in a bowl. Do not mix in a blender, since the formula becomes too thick.
0839_frame_37.fm Page 837 Tuesday, May 22, 2001 1:47 PM
837
Hand-Rearing and Artificial Milk Formulas
ESF 75–25 Fish mash 25% volume Elephant Seal Formula 75% volume Fish oil 10 ml
Mix ingredients in a bowl. Do not mix in blender as formula becomes too thick. These three formulas above may be made and kept refrigerated for 24 hours.
Feeding Frequency and Daily Requirements Any pup admitted with a full or partial blackcoat (lanugo) will be tubed HSF. Upon admission, the following schedule is used to rehydrate animals, which are slowly introduced to full-strength formula. Weight at Admission <30 kg (<66 lb) >30 kg (>66 lb)
Formula Mixture 300 ml EL 150 ml EL + 150 ml HSF 100 ml EL + 200 ml HSF 300 ml HSF 500 ml EL 250 ml EL + 250 ml HSF 125 ml EL + 375 ml HSF 500 ml HSF
Frequency/Number of Tubings q4h × 4 tubings q4h × 2 tubings q4h × 2 tubings q4h q4h × 4 tubings q4h × 2 tubings q4h × 2 tubings q4h
Once pups are on full-strength formula, the volume may be increased 50 ml/day. Pups are tubed every 4 hours, with the first tubing at 0800 hours and the last at 2400 hours. Each animal is supplemented with one marine mammal vitamin (Mazuri, Purina Mills, Inc., St. Louis, MO), 200 IU vitamin E, and 2 g NaCl, BID. Weaners are pups that are approximately 1 month old (or older), have teeth erupting, and have lost their black lanugo coat. Most weaner pups are started on ESF 50–50 and worked slowly up to ESF 75–25 and eventually to full ESF. Upon admission, the following schedule is used to rehydrate weaned animals and work them slowly up to full-strength formula. <30 kg (<66 lb) 30–35 kg (66–77 lb) >35 kg (>77 lb)
300 ml EL 150 ml EL + 150 ml ESF 50–50 300 ml ESF 50–50 500 ml EL 250 ml EL + 250 ml ESF 50–50 500 ml ESF 50–50 750 ml EL 325 ml EL + 325 ml ESF 50–50 750 ml ESF 50–50
q4h × 2 tubings q4h × 2 tubings q4h q4h × 2 tubings q4h × 2 tubings q4h q4h × 2 tubings q4h × 2 tubings q4h
Once they are on full-strength formula, the volume can be increased by 50 ml/day. Pups are tubed every 4 hours. Each animal’s diet is supplemented with 1 Mazuri Pinniped Vitamin, 200 IU vitamin E, and 2 g NaCl BID.
0839_frame_37.fm Page 838 Tuesday, May 22, 2001 1:47 PM
838
CRC Handbook of Marine Mammal Medicine
Delivery Methods and Techniques All formulas are delivered through a stomach tube using a 400-ml dose syringe. A clear vinyl tubing with 1/2 in. inside diameter and 3/4 in. outside diameter cut in 36-in. lengths is ideal for the stomach tube.
Weaning Procedures Once the animal’s teeth have erupted, the blackcoat has been shed, and the pup appears clinically healthy, fish may be offered. Initially, with the pup in a pool, offer it floating fish, live fish, or fish held in forceps. This exercise is repeated on a daily basis. If the pup accepts the floating fish, pieces or whole fish may be offered. Force feeding may be required if the pup refuses to take fish offers after several daily attempts. Techniques similar to those described in the harbor seal section are recommended.
Other Practical Information Since elephant seals usually undergo a marked fast postweaning in the wild, a reduction in body weight of up to 10% may be needed to trigger eating behavior in tube-fed animals. Other medical routines (weighing, blood sampling) are similar to those used for harbor seals.
References and Suggested Further Reading Adams, S.H., and Costa, D.P., 1993, Water conservation and protein metabolism in northern elephant seal pups during the postweaning fast, J. Comp. Physiol. B, 163: 367–373. Carlini, A.R., Marquez, M.E.I., Soave, G., Vergani, D.F., and Ronayne de Ferrer, P.A., 1994, Southern elephant seal, Mirounga leonina: Composition of milk during lactation, Polar Biol., 14: 37–42. Castellini, M.A., and Costa, D.P., 1990, Relationships between plasma ketones and fasting duration in neonatal elephant seals, Am. J. Physiol., 259 (5 Pt 2): R1086–1089. Castellini, M.A., Costa, D.P., and Huntley, A.C., 1987, Fatty acid metabolism in fasting elephant seal pups, J. Comp. Physiol. B, 157: 445–449. Castellini, J.M., Castellini, M.A., and Kretzmann, M.B., 1990, Circulatory water concentration in suckling and fasting northern elephant seal pups, J. Comp. Physiol. B, 160: 537–542. Cook, H.W., and Baker, B.E., 1969, Seal milk. I. Harp seal (Pagophilus groenlandicus) milk: Composition and pesticide residue content, Can. J. Zool., 47: 1129–1132. Gage, L.J., 1993, Hand rearing pinnipeds, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 413–415. Gage, L.J., 1994, Handrearing northern elephant seal pups (Mirounga angustirostris), in Proceedings American Association of Zoo Veterinarians and Association of Reptilian and Amphibian Veterinarians, 190. Le Boeuf, B.J., and Ortiz, C.L., 1977, Composition of elephant seal milk, J. Mammal., 58: 683–685. Marquez, M.E., Slobodianik, N.H., Ronayne de Ferrer, P.A., Carlini, A.R., Vergani, D.F., and Daneri, G.A., 1995, Immunoglobulin A levels in southern elephant seal (Mirounga leonina) milk during the suckling period, Comp. Biochem. Physiol. B, 112: 569–572. Mayer, S.J., and Hutchinson, A.J., 1990, Rearing and rehabilitation of common seal pups (Phoca vitulina), Vet. Rec., 127: 614–616. Ortiz, C.L., Costa, D., and Le Boeuf, B.J., 1978, Water and energy flux in elephant seal pups fasting under natural conditions, Physiol. Zool., 51: 166–178. Ortiz, C.L., LeBoeuf, B.J., and Costa, D.P., 1984, Milk intake of elephant seal pups: An index of parental investment, Am. Nat., 124: 416–422. Patterson-Buckendahl, P., Adams, S.H., Morales, R., Jee, W.S., Cann, C.E., and Ortiz, C.L., 1994, Skeletal development in newborn and weanling northern elephant seals, Am. J. Physiol., 267: R726–R734.
0839_frame_37.fm Page 839 Tuesday, May 22, 2001 1:47 PM
Hand-Rearing and Artificial Milk Formulas
839
Puppione, D.L., Kuehlthau, C.M., Jandacek, R.J., and Costa, D.P., 1996, Chylomicron triacylglycerol fatty acids in suckling northern elephant seals (Mirounga angustirostris) resemble the composition and the distribution of fatty acids in milk fat, Comp. Biochem. Physiol. B, 114: 53–57. Riedman, M., 1990, The Pinnipeds, Seal, Sea Lions and Walruses, University of California Press, Berkeley, 439 pp. Shaughnessy, P.D., 1974, An electrophoretic study of blood and milk proteins of the southern elephant seal, Mirounga leonina, J. Mammal., 55: 796–780.
Sea Lions California sea lions (Zalophus californianus) are the species most commonly reared by the method below, but Steller sea lions (Eumatopias jubatus) and northern fur seals (Callorhinus ursinus) have also been reared successfully using it.
Formula Fish paste (see below) Pedialyte Safflower oil Lecithin Heavy whipping cream Lactase Multivitamin NaCl Vitamin B1 Vitamin C Vitamin E Calcium gluconate
0.23 kg 150 ml 5 ml 5 ml 200 ml 0.75 ml 1 capsule 1.25 g 250 mg 250 mg 400 IU 150 mg
Note: The whipping cream must be treated with the 0.75 ml lactase 24 hours prior to use. To prepare fish paste, use high quality fish, such as herring that is greater than 10% fat, and remove the heads, tails, and fins. Begin with 0.34 kg of fish cut into pieces and placed in a food processor with 100 ml of Pedialyte. Blend until the mixture is a smooth paste, transfer to rice strainer, and grind the larger bone fragments out of the paste. This process will yield about 0.23 kg of herring paste. Add this herring paste to a blender with the other ingredients, except the whipping cream, and blend until smooth. Pour the contents into a clean container and add the 200 ml of lactase-treated heavy whipping cream. Rock the container to blend the fish paste with the whipping cream. This yields approximately 637 ml of formula. Label the container with the date and time. The formula should be kept refrigerated until used, and warmed to approximately 30°C prior to feeding. Formula should be discarded after 24 hours.
Delivery Methods and Techniques Bottle-feeding is the preferred method for feeding sea lion pups. Some pups prefer a rubber lamb’s nipple; others accept only nursers or baby nipples for humans. It often will take 3 or 4 days of constant encouragement before a pup will accept the bottle. Pups that do not suckle readily will need to be fed via stomach tube until they accept the bottle. California and Steller sea lions imprint easily on their caregivers, and extreme measures must be taken to eliminate virtually all human contact with pups that are to be released back into the wild. This poses a problem when encouraging pups to nurse from a bottle; however, once this process has been achieved, bottles may be placed so they emerge from a large padded box or other similar arrangement that allows the pup to nurse comfortably without human contact.
0839_frame_37.fm Page 840 Tuesday, May 22, 2001 1:47 PM
840
CRC Handbook of Marine Mammal Medicine
Feeding Frequency and Daily Requirements The initial feeding should be an electrolyte formula with 5% glucose via stomach tube at 20 ml/kg body weight. The second feeding consists of 50% electrolyte solution and 50% formula giving the same volume as the initial feeding. The total volume the second day is 100 ml/kg via stomach tube divided into five or six feedings on the second day. Offer 120 to 150 ml of formula at each of the five to six feedings daily, initially. Once the pup is nursing well, the amount of formula offered each feeding may be increased gradually. By 3 to 4 weeks of age the pup should be suckling 200 to 250 ml at each of the five feedings. By 6 weeks of age pups should be nursing 250 to 300 ml/feeding four times a day. If the pup will not accept the bottle, it must be fed via stomach tube until it can be weaned. Pups fed by stomach tube will not tolerate the same amounts per feeding as pups that are nursing (Gage, 1993). The amounts given via gavage should be reduced by 15 to 20%, and the full amount should be given very slowly to help avoid regurgitation.
Weaning Procedures Although sea lions may suckle until 1 year of age in the wild, pups weighing over 12 kg (26 lb) may have fish introduced to their diet (Gage, 1990). Ground chunks of fish may be offered to the pups mixed in the bottle with their formula. Eventually, enlarge the nipple hole and feed small fish pieces through it in place of the formula. Pups may also be force-fed small pieces of fish at this point. It may take 1 to 4 weeks until they begin to eat fish voluntarily. This process may be accelerated using the force-feeding method. Gradually increase the amount of fish offered until the pups are eating 20% of their body weight per day. Vitamin supplements should be given daily. Another method that can be used to wean the pup from formula to a fish diet is the “ice cube method” (Gage, 1990). The formula feeds are reduced to twice daily and the pup is given ice cubes to play with. Small bits of fish are frozen into the ice cubes. The pups may reject the first “fish cubes” but will eventually accept them. Once the pup is eating these, a slurry of ground fish is frozen into cubes. Once the pup accepts these, offer small pieces of fish and then whole fish. The entire process usually takes about 1 week, and the pups may lose 2 kg (4.4 lb) body weight during this process (Gage, 1990).
Other Practical Information Sea lion pups are given a physical examination, weighed, and blood is drawn for complete blood counts and serum chemistry analysis. Pups are weighed daily for the first week, then twice a week, and then once a week when older than 3 months. Sea lion pups should be allowed to swim under supervision in shallow water when they are 2 to 3 days old. They may need to be supported in deeper water. Salt water is preferable, although pups may be successfully raised with access to only fresh water. These pups must be supplemented with NaCl at approximately 2 to 3 g/day. Small pups under 7 kg (15 lb) should be brought inside at night if the ambient temperature is below 20°C (68°F).
References and Suggested Further Reading Arnould, J.P.Y., and Boyd, I.L., 1995, Inter- and intra-annual variation in milk composition in Antarctic fur seals (Arctocephalus gazella), Physiol. Zool., 68: 1164–1180. Arnould, J.P.Y., and Boyd, I.L., 1997, Lactation and the cost of pup-rearing in Antarctic fur seals, Mar. Mammal Sci., 13: 516–526. Arnould, J.P., and Hindell, M.A., 1999, The composition of Australian fur seal (Arctocephalus pusillus doriferus) milk throughout lactation, Physiol. Biochem. Zool., 72: 605–612.
0839_frame_37.fm Page 841 Tuesday, May 22, 2001 1:47 PM
841
Hand-Rearing and Artificial Milk Formulas
Arnould, J.P.Y., Boyd, I.L., and Socha, D.G., 1996, Milk consumption and growth efficiency in Antarctic fur seal (Arctocephalus gazella) pups, Can. J. Zool., 74: 254–266. Dosako, S., Taneya, S., Kimura, T., Ohmori, T., Daikoku, H., Suzuki, N., Sawa, J., Kano, K., and Katayama, S., 1983, Milk of northern fur seal: Composition, especially carbohydrate and protein, J. Dairy Sci., 66: 2076–2083. Gage, L.J., 1990, Hand-rearing pinniped pups, in Handbook of Marine Mammal Medicine, Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 333–535. Gage, L.J., 1993, Hand rearing pinnipeds, in Zoo and Wild Animal Medicine, 3rd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 413–415. Gales, N.J., Costa, D.P., and Kretzmann, M., 1996, Proximate composition of Australian sea lion milk throughout the entire supra-annual lactation period, Aust. J. Zool., 44: 651–657. Kretzmann, M.B., Costa, D.P., Higgins, L.V., and Needham, D.J., 1991, Milk composition of Australian sea lions, Neophoca cinerea: Variability in lipid content, Can. J. Zool., 69: 2556–2561. Trillmich, F., and Lechner, E., 1986, Milk of the Galapagos fur seal and sea lion, with a comparison of the milk of eared seals (Otariidae), J. Zool., 209: 271–277. Trillmich, F., Kirchmeier, D., Kirchmeier, O., Krause, I., Lechner, E., Scherz, H., Eichinger, H., and Seewald, M., 1988, Characterization of proteins and fatty acid composition in Galapagos fur seal milk. Occurrence of whey and casein protein polymorphisms, Com. Biochem. Physiol. B, 90: 447–452. Werner, R., Figueroa-Carranza, A.L., and Ortiz, C.L., 1996, Composition and energy content of milk from southern sea lions (Otaria flavescens), Mar. Mammal Sci., 12: 313–317.
Walruses Formulas Beginning Formula Zoologic Milk Matrix 30/55 1 part to 2 parts boiled or bottled water Daily supplements: Pinniped mulitivitamin (Mazuri, Purina Mills, Inc.) 1 tablet Dicalcium phosphate 455 mg Taurine 250 mg Vitamin E 800 IU Maintenance formula Zoologic Milk Matrix 30/55 Water (boiled or bottled) Herring (thawed, whole, fins and tails removed) Clams (thawed, ground) Salmon oil Pinniped mulitivitamin (Mazuri, Purina Mills, Inc.) Dicalcium phosphate Taurine Vitamin E
400 g 800 ml 550 g 114 g 90 ml 1 tablet BID 500 mg BID 250 mg BID 400 IU BID
Label, refrigerate, and discard after 24 hours. Maintenance formula is strained to prevent buildup of small particles in the nipple. The ratio of Milk Matrix to water may be increased gradually to 1.5 parts Zoologic Milk Matrix (Pet-Ag, Inc., Hampshire, IL) to 2 parts water to improve the caloric content of the formula. Beware that the Milk Matrix dry product varies in consistency, and the dilution may need to be adjusted with each new lot to maintain consistency.
0839_frame_37.fm Page 842 Tuesday, May 22, 2001 1:47 PM
842
CRC Handbook of Marine Mammal Medicine
Feeding Frequency and Daily Requirements For the first 24 to 48 hours, give a mixture of electrolyte solution and water with a stomach tube (Samansky, 1995). Initially, start with a very dilute formula (10% formula to 90% water) and increase it to 1 part powdered formula to 2 parts water over the course of 2 days. Continue to increase the concentration of the formula slowly. The slow increase in concentration may prevent digestive problems. Walrus calves should be fed 100 Kcal/kg body weight/day to expect average weight gains of about 0.5 to 0.7 kg/day (1.1 to 1.5 lb) (Samansky, 1995). Weighing the calf each day prior to the first feeding will help monitor the feeding regime. Continue to weigh the calf daily for at least the first month; thereafter weekly weighing should be sufficient. At 10 to 12 weeks the calves are placed on the maintenance formula that will be fed until weaning. Feed young calves six times a day. This schedule may be reduced to five times a day starting the second or third month, while keeping the total daily formula intake constant. The number of feedings may be reduced to four times a day when the calves are 4 months old.
Delivery Methods and Techniques It is often necessary to tube-feed orphaned calves, because they often will not readily accept a bottle. A calf-type bottle with either a calf or lamb’s nipple should be tried. Some calves will accept the smaller lamb’s nipple more readily than a calf nipple. If it is necessary to use a stomach tube, extra care should be taken to avoid inadvertently placing the feeding tube into the trachea. It is recommended to use water in the bottle, especially if the calves become dehydrated. Calves appear to be very sensitive to formula temperature; suggested formula temperature is 39 to 40°C (102 to 104°F).
Weaning Procedures At 2 to 3 months of age, start adding ground fish/clams to the formula. At 4 to 6 months, offer whole fish, clams, and fish pieces in addition to the formula. At 8 to 12 months of age, begin to gradually reduce the formula. Walrus calves should be weaned completely off formula at about 18 months of age (Samansky, 1995).
Other Practical Information Walruses are given a physical examination, weighed, measured, and blood is drawn from the epidural intravertebral sinus for a complete blood count and serum chemistry analysis. Hydration status is evaluated and corrected by giving appropriate fluids. Walruses rapidly imprint on their caregivers, which is an important consideration if the animal is to be released back into the wild. Whenever possible, walrus pups should be housed together to encourage conspecific socialization. Cleanliness of the environment and the feeding equipment is of paramount importance to the success of hand rearing these animals. It is important to clean, disinfect, and dry equipment (bottles, nipples, blender, etc.) after each use. Most of the calves that have been hand-reared have received maternal milk and colostrum. If this has not happened, maternal serum should be given orally if available. Proper hygiene is important both for the caregivers and the neonates. Enteric salmonellosis occurred in walrus calves with subsequent zoonotic spread (Calle, 1995).
References and Suggested Further Reading Calle, P., 1995, Enteric salmonellosis of captive Pacific walrus (Obobenus rosmarus divergens), in Proceedings of the 26th International Association for Aquatic Animal Medicine, Mystic, CT, 92–93. Gage, L.J., Samansky, T., Rutherford, S., Allen. S., Turley, P., and Chapple, J., 1996, Hand raising orphaned walrus calves, in Proceedings of the American Association of Zoo Veterinarians, 77.
0839_frame_37.fm Page 843 Tuesday, May 22, 2001 1:47 PM
Hand-Rearing and Artificial Milk Formulas
843
Samansky, T., 1995, Hand raising orphaned walrus calves at Marine World Africa USA—The first year, in Proceedings of the 26th International Association for Aquatic Animal Medicine, Mystic, CT, 34–35.
Manatees Analysis of manatee milk has revealed that taurine is a major amino acid and that short- and medium-chained fatty acids are abundant (Pervaiz and Brew, 1986a,b; Walsh et al., 1996). With this information, it was determined that the original Esbilac (also known as Milk Matrix 33/40 milk replacer*) with 16 ml of either soybean or canola oil was comparable with manatee milk. The most common complication with this formula was usually related to constipation. The constipation was most likely due to starting a dehydrated calf on the formula before correcting and maintaining hydration during the adjustment to the bottle.
Formulas Miami Seaquarium Formula Zoologic (Milk Matrix 30/55) 400 g Goats milk 210 ml Isomil 120 ml Water 400 ml Lactinex 1 tablet/100 ml formula Children’s chewable vitamins 1 tablet/300 ml formula
Add 20 ml safflower oil or canola oil per 300 ml if calf is not gaining weight. The formula may be stored in the refrigerator up to 8 hours. Thoroughly clean and disinfect bottles between feedings. It is recommended that the bottles be boiled between feeds when raising infants <55 kg body weight. SeaWorld Formula Zoologic (Milk Matrix 33/40) Water Taurine B-complex Canola oil
500 g 1000 ml 250 mg/l formula 1 tablet/l formula 4 to 16 ml/100 g formula
Gradually increase the canola oil; at 4 ml/100 g formula, the caloric density is approximately 112 Kcal/100 g. These formulas are recommended when the ingredients are available. A number of other formulas have been utilized using goat milk alone or soy-based products. Analysis of manatee milk reveals the presence of a number of sugars, including small amounts of lactose. Manatee calves can tolerate up to 5% of the diet as a carbohydrate.
Delivery Methods and Techniques Often manatee calves will not nurse from a bottle, and it may be necessary to utilize a stomach tube to deliver the formula and prevent dehydration. A 12 to 16 French stomach tube may be used for oral or nasogastric intubation. To determine the proper depth the tube will need to * Milk Matrix 33/40 is the old spray-dried Esbilac formula with a fat source of soybean and coconut oil. The current Esbilac formula is an agglomerated product, instantized to make it easier to mix. The fat source was changed to canola oil, butter fat, and soybean oil in 1993. The new fat source combination provides a better fatty acid makeup for domestic dog puppies.
0839_frame_37.fm Page 844 Tuesday, May 22, 2001 1:47 PM
844
CRC Handbook of Marine Mammal Medicine
be inserted, place the tube alongside the calf ’s body with the tip of the tube extending to the end of the pectoral fin, and mark the tube at the calf ’s mouth with a piece of tape or indelible ink. Calves will often take a few days to accept a nipple when bottle-feeding. Formula may be fed with a Johnson and Johnson “health flow” baby bottle, using a lamb or calf nipple. Initially introduce the bottle hourly. Some calves prefer to nurse on their backs. It is important that all caregivers use similar nursing techniques to encourage the calf to accept a bottle more rapidly.
Feeding Frequency and Daily Requirements Day 1 Day 2 Day 3 to 4 Day 5 Day 6
Glucose and electrolytes 25% formula, 75% water 50% formula, 50% water 75% formula, 25% water 100% formula
Once the calf begins suckling, offer a bottle every 2 to 3 hours. The maximum formula allowed per feeding should be 300 ml. If dehydrated, the calf may need to be tube-fed between bottlefeeding attempts. Calves nursing well on a bottle may gain 0.3 to 0.5 kg (0.6 to 1.1 lb) of weight per day. A 20-kg (44-lb) calf can safely be given 125 to 150 ml of formula with a stomach tube every 3 hours. If the calf is very thin or hypoglycemic, add 1 ml of 50% dextrose/kg body weight to the formula. If glucose is added, a balanced electrolyte solution is used in place of water. The stool and blood should be monitored for excessive glucose concentrations. The amount of glucose may be reduced as other nutrients are utilized as an energy source. The utilization of fat may reduce the additional glucose requirements. If there is intestinal disease and the formula is not appropriate, the calves should be given an elemental formula comprised of: Criticare HN® (Mead Johnson, Evansville, IL) 35 ml MCT oil (medium-chain triglycerides) 28 ml Nutramigen (baby formula) 103 ml
This formula contains 200 kcal/100 g and is recommended with thin, hypoglycemic calves during the initial adjustment period or for severe colitis or pneumotosis intestinalis (Walsh, pers. comm.).
Weaning Procedures The calves may need to be fed for longer periods when they are maintained with adults, and access to solid food is limited to lettuce. When competition is a factor, the calf may need supplementation for 18 to 24 months. Access to green vegetation may be initiated as early as 1 month of age. During the weaning process, the calves should be weighed every week, more or less, depending on the progression of weight gain (or loss).
Other Practical Information Orphaned manatee calves often present with a number of medical problems, including emaciation, hypoglycemia, dehydration, hypothermia, septicemia, entercolitis, and constipation. The presence of an umbilical cord can help estimate age, but newborns typically have weight ranges from 17 to 50 kg (average 20 to 30 kg; 44 to 66 lb). Calves should have blood sampled for complete blood counts and serum chemistries. They are sampled once or twice a week to monitor their hydration status and to evaluate their overall condition. Initially, it is important to monitor blood glucose levels. With severe hypoglycemia, intravenous 5% dextrose in water is recommended, but
0839_frame_37.fm Page 845 Tuesday, May 22, 2001 1:47 PM
845
Hand-Rearing and Artificial Milk Formulas
intravenous infusion is difficult. Glucose supplementation is designed as a preventive measure until the calf is able to utilize other sources of energy. Underweight neonates that are initially hypoglycemic and respond well to glucose, but are not adequately supplemented, will reach metabolic exhaustion within a week. Therefore, supplemental nutrition must include proteins and fats within the first few days of hand rearing. Manatee colostrum contains 46 Kcal/100 g; milk during midlactation has 189 Kcal/100 g; and 130 Kcal/100 g in late lactation (Walsh, pers. comm., 2000). Some formulas make up for the lack of caloric density by increasing the daily volume. Manatee colostrum and serum immunoglobulins have been utilized with these neonates (Walsh and Bossart, 1999). Manatee IgG is given intravenously and/or orally to very young calves. Neonatal calves are often maintained in freshwater pools between 30 and 32°C (86 and 90°F) with 1 part per million chlorine.
References and Suggested Further Reading Pervaiz, S., and Brew, K., 1986a, Composition of the milks of the bottlenose dolphin (Tursiops truncatus) and the Florida manatee (Trichechus manatus latirostris), Comp. Biochem. Physiol. A, 84: 357–360. Pervaiz, S., and Brew, K., 1986b, Purification and characterization of the major whey proteins from the milks of the bottlenose dolphin (Tursiops truncatus), the Florida manatee (Trichechus manatus latirostris), and the beagle (Canis familiaris), Arch. Biochem. Biophysiol., 246: 846–854. Walsh, M., and Bossart, G., 1999, Manatee medicine, in Zoo and Wild Animal Medicine, 4th ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 513. Walsh, M.T., Oftedal, O.T., Worthy, G.A.J., Rodgers, Q.R., Innis, S.M., and Campbell, T.W., 1996, Manatee milk analysis: Changes through nursing, in Proceedings of the 27th International Association for Aquatic Animal Medicine, 67.
Sea Otters Formula and Preparation Clams, finely chopped Squid, remove beak, quill, and ink sac Dextrose 5% in water Lactated Ringer’s solution Whole milk/whipping cream 50/50 Multivitamin supplement with iron (HiVite) Dicalcium phosphate Cod liver oil
120 g 120 g 100 ml 100 ml 200 ml 2 ml 500 mg 2 ml
Chop squid and clams into fine pieces and then puree (do not whip). Add liquids and vitamins/ minerals to puree and mix well. Blend in dairy products last. Treat the dairy blend with Lactaid® (McNeil Consumer Healthcare, Fort Washington, PA) and/or lactobacillus to aid in milk digestion if diarrhea occurs. Add ALL BRAN® (Kellogg’s, Battle Creek, MI) cereal (5 g to formula batch above) for fiber, if stools are chronically soft. Mix enough formula for 24 hours, label with time and date of preparation, and refrigerate. Discard unused portion after 24 hours or immediately freeze individual portions and label with time and date, thaw/warm, and use as needed. Discard unused frozen portions after 1 week.
Delivery Methods and Techniques Very young sea otter pups seem to adapt to nursing a bottle readily. Older pups may not have a strong enough suckle urge and may need to be tube-fed until weaned onto solids. If
0839_frame_37.fm Page 846 Tuesday, May 22, 2001 1:47 PM
846
CRC Handbook of Marine Mammal Medicine
pups will nurse on a bottle, use an infant latex nurser bottle with disposable bags (4-oz size for newborns up to 4 kg, or 9 lb, body weight, and then the 8-oz size), plus a standard infant nipple with an “X”-cut opening at the nipple aperture to accommodate the thicker formula. Feed formula to the pup (face down, lying on its belly) and push the formula through the nipple by applying pressure to the nurser bag with a homemade “syringe plunger,” allowing delivery of formula more rapidly as the pup nurses and/or sucks food from the nipple. Pups like the nipple pushed up through a hole in a clean terry washcloth. If feeding via gavage is necessary, use a red rubber urethral catheter, 12 to 14 French, 16 in. long, and a 60-ml catheter-tip syringe.
Feeding Frequency and Daily Requirements Because sea otters have a high metabolic rate (see Chapter 36, Nutrition), they require a food intake of 20 to 35% of their body weight daily (see Chapter 44, Sea Otters). Initially, their daily formula intake should equal 30% of their body weight divided into eight to ten feedings for the first 1 to 3 weeks of age. At this point, gradually increase the amount of formula offered at each feeding, and decrease the feeding frequency to four to six feedings per day. Adjust total daily formula amount based on the rate of weight gain. Weigh pups at the same time each day, preferably in the morning prior to the first feeding. Expect an average weight gain of approximately 1% body weight per day. The weight may plateau every 2 to 5 days. Loss of more than 5% body weight at any time or loss of weight over two consecutive days may be because too little formula is being offered, or may indicate a medical problem.
Weaning Procedures At 3 to 4 weeks of age, offer small soft pieces of clam, squid, peeled shrimp, or other seafood before offering formula. Record total weight of solid food consumed. At 6 weeks of age or when the pup is taking soft bits readily, open shells and loosen the muscle of whole clams or mussels. At 8 to 10 weeks, start to offer unpeeled shrimp and whole crab legs, and then intact mussels by 12 to 16 weeks of age. Gradually decrease formula feedings equal to the measured amount of voluntary solid food intake. Most pups are weaned to an exclusively solid diet by 4 months of age. Continue to weigh each pup frequently and supplement with formula if weight loss occurs, or if the pup fails to gain weight.
Other Practical Information Pups should be given a physical examination upon admission. The general body condition, including estimation of age, nutritional state, weight, hydration, temperature, and evidence of disease or injury should be noted. Proper grooming is vital. Dermatitis can easily result if the coat is not kept clean and dry. Pups will chill rapidly in water if the coat is left wet. Body temperature monitoring is vital during the first few weeks of rehabilitation. Severe hypothermia or hyperthermia may occur in a 10- to 30-min period. Room temperature for the pup nursery should be 15°C (60°F), and water temperature for swimming should be about the same. Water quality is also very important. Fresh water may be used for bathing, but must be nonchlorinated. Salt water may be mixed with a home aquarium salt mix, and pups may be dipped at the end of each bath or swim period. This allows the use of fresh water for stimulation of urine and defecation and for exercise. This water can be disposed of at each cycle. Grooming and drying with salt water on the coat to condition the fur is recommended. Socialization with other otters is desirable as early in the rehabilitation process as possible.
0839_frame_37.fm Page 847 Tuesday, May 22, 2001 1:47 PM
847
Hand-Rearing and Artificial Milk Formulas
References and Suggested Further Reading Sandegren, F.E., Chu, E.W., and Vandevere, J.E., 1973, Maternal behavior in the California sea otter, J. Mammal., 54: 668–679. Tuomi, P., 1990, Sea otter pups (Enhydra lutris), Wildl. J., 13(3): 9–14. Williams, T.D., and Hymer, J., 1992, Raising orphaned sea otter pups, J. Am. Vet. Med. Assoc., 201: 688–691. Williams, T.D., Styers, D., Hymer, J., Rainville, S., and McCormick, C.R., 1995, Care of sea otter pups, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-bearing Marine Mammals, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 133–140.
Polar Bears Hand-rearing polar bear cubs is more complicated than hand-rearing most other marine mammals. This section offers general advice, but the reader is urged to read more in-depth documentation or to contact individuals with success hand-rearing these animals. Polar bear cubs should be placed in an infant incubator with a temperature range of 27 to 31°C (82 to 88°F) and 40 to 50% humidity (Michalowski, 1971; Wortman and Larve, 1974; Hess, 1976). The temperature is gradually reduced to room temperature over a period of weeks. The umbilicus should be treated with 0.5% chlorhexidine (Nolvasan 2% solution diluted with sterile water) every 6 hours for 24 hours. In most reported cases, hand-reared cubs are initially given polar bear serum both orally and parenterally (Wortman and Larve, 1974; Hess, 1976; Kenny et al., 1999). Medical problems associated with formula composition have occurred, including fatal thiamine deficiency/Chastek’s paralysis (Hess, 1976), vitamin D deficiency/rickets (Kenny, 1999), lactobezoars, constipation, dehydration, and bloating (Hess, 1976; Kenny et al., 1999).
Formulas Most reported formulas have Esbilac® (Pet-Ag, Inc., Hampshire, IL) and distilled water as the major ingredients; however, dilution varies from 1 part Esbilac to 3 parts distilled water, to a 1:1 ratio. Additional fat was added with half cream and half milk (half-and-half), whipping cream, or safflower oil. The supplements were common to most formulas. Two formulas have been successful. Esbilac Distilled water Half-and-half
1 part 1 part 2 parts
Esbilac Distilled water Whipping cream
1 part 3 parts 1 part*
or
To either of the above formulas add: Cod liver oil Karo syrup NeoCalglucon Liquid pediatric vitamins (e.g., Poly-vi-sol) Liquid iron supplement (e.g., Fer-in-sol)
5 ml/day (increase to 10 ml as cub grows) 4 ml/100 ml formula 2.5 ml/100 ml formula 1 ml/day 3 drops/100 g formula
Discard unused formula after 24 hours. * If whipping cream is used, it is recommended to pretreat with lactase for 24 hours.
0839_frame_37.fm Page 848 Tuesday, May 22, 2001 1:47 PM
848
CRC Handbook of Marine Mammal Medicine
Delivery Methods and Techniques Use a human “preemie” nipple with the bottle, eventually enlarging the nipple hole as the cub grows. In one case, a human “cleft palate” nipple worked best. If the cub will not nurse from the bottle, a nasogastric tube is utilized; a polyurethane infant feeding tube works well.
Feeding Frequency and Daily Requirements Initially, the formula should be diluted with water, 1 part formula to 3 parts water, and gradually increased to full strength by the end of the first week. Initially, with newborns, feed approximately 1 oz (30 ml) per feeding every 2 hours. Eventually reduce the feeding to six to seven times a day. A guideline for the amount of formula per day is 15 to 25% of body weight per day (Kenny, 2000). This is only a guideline; daily weight should be measured at the same time each day. Gradual change in formula type or amount is recommended.
Weaning Process At 10 to 16 weeks of age, the weaning process is initiated. Start by offering milk formula in a bowl, introduce precooked baby cereal (Gerber rice cereal®; Fremont, MI) mixed with formula. Slowly add solid food soaked in formula, such as Hill’s Prescription Diet C/D or I/D® (Hill’s Pet Nutrition, Inc., Topeka, KS), or ZuPreem omnivore diet® (Lunexa, KS). This should be a gradual process taking 2 months to complete. Dietary manipulations should be gradual with limited amounts of one item adjusted at a time.
Other Practical Information Offer water in a shallow pan to avoid a tendency for the cubs to immerse their entire heads into the bowl when learning to drink water. One case reported convulsions, resulting from inadvertent water intoxication (Michalowski, 1972). Abrasions and contusions are reported from the cubs “rooting around” in the incubators (Hess, 1976). Therefore, the material used for bedding and enclosures should be selected carefully.
References and Suggested Further Reading Baker, B.E., Hatcher, V.B., and Harington, C.R., 1967, Polar bear milk, 3. Gel-electrophoretic studies of protein fractions isolated from polar bear milk and human milk, Can. J. Zool., 45 (Suppl.): 1205–1210. Blix, A.S., and Lentfer, J.W., 1979, Modes of thermal protection in polar bear cubs—At birth and on emergence from the den, Am. J. Physiol., 236: R67–74. Cook, H.W., Lentfer, J.W., Pearson, A.M., and Baker, B.E., 1970, Polar bear milk, IV, Gross composition, fatty acid, and mineral constitution, Can. J. Zool., 48: 217–219. Derocher, A.E., Andriashek, D., and Arnould, J.P.Y., 1993, Aspects of milk composition and lactation in polar bears, Can. J. Zool., 71: 561–567. Hedburg, G., in press, Hand raising polar bear cubs, in Hand Raising Wild and Domestic Mammals, Iowa State University Press, Ames. Hess, J., 1976, Hand-reared polar bear cubs (Thalarctos maritimus) at the St. Paul Zoo, Int. Zoo Yearb., 11: 102–107. Kenny, D.E., Irlbeck, N.A., and Eller, J.L., 2000, Rickets in two hand-reared polar bear (Ursus maritimus) cubs, J. Zoo Wildl. Med., 30: 132–140. Michalowski, C., 1971, Hand-rearing polar bear cubs (Thalarctos maritimus) at Rochester Zoo, Int. Zoo Yearb., 11: 107–109.
0839_frame_37.fm Page 849 Tuesday, May 22, 2001 1:47 PM
Hand-Rearing and Artificial Milk Formulas
849
Michalowski, D., 1972, Hand-rearing polar bear cubs (Thalarctos) (second 90 days) at Rochester Zoo, Int. Zoo Yearb., 12: 175–176. Oftedal, O.T., 1993, The adaptation of milk secretion to the constraints of fasting in bears, seals, and baleen whales, J. Dairy Sci., 76: 3234–3246. Wortman, J., and Larve, M., 1974, Hand-rearing polar bear cubs (Thalarctos maritimus) at Topeka Zoo, Int. Zoo Yearb., 14: 215–240.
Acknowledgments Many individuals were responsible for the information provided in this chapter. Each of these people was very willing to share his or her experience and knowledge regarding hand-rearing marine mammals. Cetaceans—Jay Sweeney (Dolphin Quest) and Tom Reidarson (SeaWorld); Harbor seals/elephant seals—Frances Gulland and Joan Sicree (The Marine Mammal Center) and Pam Tuomi and Lynn A. Adecholt (Alaska SeaLife Center); Manatees—Maya Dougherty (Miami Seaquarium) and Mike Walsh (SeaWorld); Greg Bossart (Harbor Branch Oceanographic Institute); Sea otters—Pam Tuomi (Alaska SeaLife Center); Walruses—Terry Samansky (Six Flags Marine World); Polar Bears—Gail Hedberg (San Francisco Zoological Society), Kerri Slifka (Zoo Nutrition Services, Chicago Zoological Society Brookfield Zoo), and David E. Kenny (Denver Zoological Foundation). The authors thank Tanya Zabka for compiling the references, and give a special thanks to Kasey Cooley for her patience and help in preparing this chapter.
0839_frame_37.fm Page 850 Tuesday, May 22, 2001 1:47 PM
0839_frame_C38.fm Page 851 Tuesday, May 22, 2001 2:01 PM
38 Tagging and Tracking Michelle E. Lander, Andrew J. Westgate, Robert K. Bonde, and Michael J. Murray
Introduction Tagging and tracking of marine mammals is important to veterinarians, biologists, and managers. Monitoring the ability of rehabilitated marine mammals to forage, dive, survive, and ultimately reproduce following release is essential for determining the efficacy of the rehabilitation process. Many postrelease monitoring programs for rehabilitated marine mammals, although limited in the past, have now been implemented (Harvey, 1991; Lander, 1998; Bonde et al., 1999; Early et al., 1999). Biologists and managers have been able to assess habitat use, home range, and resource utilization of free-ranging and rehabilitated marine mammals by tracking their movements and migrations. Postrelease monitoring depends upon the ability to monitor animals in the wild using a variety of tracking techniques, which may be constrained by habitat, animal species, molting phase, and equipment design and cost. Reliability and performance of different tracking methodologies often dictate the success or failure of any tracking program, so investigators should be familiar with the use and limitations of various types of tags and equipment available (Kenward, 1987). This chapter summarizes standard techniques most commonly used to tag and track marine mammals.
Tracking Methodologies: A Brief Overview Many methods, ranging from natural markings to sophisticated instruments, are used to identify or tag marine mammals for the purpose of following or “tracking” them. Tags can be classified into two basic types: passive and active. Passive tags are those that assist solely in the identification of individual animals. There are two types of passive tags. The first is natural markings on individuals such as unique pigmentation patterns, scars, cuts, notches, or unique dorsal fin and fluke shapes. The second is devices or marks that are attached physically to animals, which may or may not be permanent. Examples include cattle tags, button tags, streamer tags, spaghetti tags, passive integrated transponder (PIT) tags, nylon flags, dye marks, tattoos, freeze brands, and heat brands. Both natural and artificial markings have been used to study many different species in the wild and have greatly advanced fields that rely on photographic identification techniques. Their ultimate utility is limited, however, because animals must be resighted and the identifying marks seen (and sometimes photographed) for these tags to be effective.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
851
0839_frame_C38.fm Page 852 Tuesday, May 22, 2001 2:01 PM
852
CRC Handbook of Marine Mammal Medicine
Resighting individuals can be difficult because most behaviors of marine mammals occur below the sea surface or at great distances from land (Wilson et al., 1986). Active tags, including telemetry devices and archival instruments, are those that transmit or collect data. In addition to helping identify individuals, they provide a means to locate animals and collect information about an animal’s behavior and its surroundings. Telemetry is the science of measuring and transmitting data from a distant source via acoustic or radio frequencies. These data can be in the form of a simple location beacon, or relayed information collected by sensors on a tag. Conventional radio telemetry allows individuals to be identified using pulsed transmitters at different frequencies for different subjects. The signals from a radio transmitter are in the form of pulses or beeps, because it is easier for the human ear to perceive a pulsed signal than a steady tone, and this increases the longevity of the battery. Pulse rates of transmitters are configured by the manufacturer and will depend on the type of application. When selecting a pulse rate, consider the typical surfacing interval of the animal being tagged and select a pulse rate that will provide at least two full beeps per surfacing. Pulse rates commonly used for deployments on smaller cetaceans are between 90 and 110 pulses per minute (ppm), whereas pulse rates of 25 to 65 ppm are typically used to study pinnipeds and manatees. Lower pulse rates may be used if one is interested only in locations of animals when they are onshore. Manipulating pulse rates can optimize the life expectancy of most transmitters. Although some radio transmitters produce low-frequency signals, most radio-tracking equipment operates in the very high frequency (VHF) range of the electromagnetic spectrum, and VHF frequency bands that are allocated to wildlife radio tags differ among countries. Most VHF transmitters operate in the 138 to 180 MHz Land Mobile Band and the frequency of the VHF transmitter used will be dictated by the reception capabilities of the receiving equipment. Allocation of legal frequency ranges are restricted and regulated in the United States by the Federal Communications Commission (FCC). Unless otherwise specified, the manufacturer will select the frequencies of the transmitters purchased within the frequency range. Should the purchaser choose to select his or her own frequencies, a frequency separation of at least 10 to 20 kHz among transmitters is recommended. The process of radio telemetry signal acquisition requires a system consisting of a receiver and an antenna. Because VHF frequencies are not within frequency ranges audible to humans, the primary function of a telemetry system is to receive, amplify, and translate signals emitted from radio transmitters down to an audible range (Beaty, 1989). The receiver is a device that converts radio waves into sound, and the antenna is a transformer, providing a mechanism for interfacing between the radio wave emitted from the radio transmitter and the receiver (Tomkiewicz, 1988). Most telemetry receivers come with 2-MHz of bandwidth (e.g., 164 to 165 MHz), although smaller and larger bandwidths are available. When selecting a specific bandwidth, other people who use telemetry in the area where a researcher intends to work should be consulted. Although there are concerns about having too many researchers working in a limited frequency space, this is far outweighed by the fact that they can assist each other with tracking needs and equipment loans. For example, most cetacean radio trackers along the eastern seaboard of the United States work in the 148 to 150 MHz range. Antennas are an integral part of any telemetry receiving system, and it is important to purchase antennas that are tuned to the appropriate bandwidth. Like receivers, antennas usually are tuned to cover a 2-MHz range. Directional antennas are used to receive signals, and determine the direction from which the signal strength is greatest. Two types of antenna systems commonly used for wildlife radio tracking include two-element H-Adcocks and three- or four-element Yagis. The latter possess superior gain and are better suited for tracking on the ocean. When tracking marine mammals on the open ocean, the elements of the antenna should be oriented perpendicular to the surface of the water. Orientation of the
0839_frame_C38.fm Page 853 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
853
antenna in the same plane as that of the transmitting antenna on the radio tag will improve the overall gain and directionality of the received signal. H-Adcocks are smaller and more commonly used when tracking from airplanes and work better in this situation. Aerial tracking is useful when working in rugged or inaccessible areas, and is often the preferred method for locating animals during periods of dispersal or migration (Kenward, 1987). Aircraft antenna mounting kits are available from most wildlife telemetry manufacturers. Additional accessories, such as data loggers and automatic directional finders, are available to enhance, support, and simplify tracking. Transmitters come in all shapes and sizes and are usually configured by the manufacturer to suit specific needs (see Table 1 for a listing of some radio tag manufacturers). If unsure about specific applications, tag manufacturers can provide insight, expertise, and support with technical options that should be considered. When ordering transmitters, it is important to consider the method of attachment, and the size, shape, and mass of tags with respect to potential alterations in the animal’s performance (Costa, 1988). For example, the weight of transmitters, including attachment mounts, should not exceed 2 to 3% of the animal’s body mass (Kenward, 1987; Advanced Telemetry Systems, Inc. (ATS) Web page, see Table 1), and they should be hydrodynamically shaped. Transmitters must also be capable of withstanding high pressure, because as animals dive, the pressure increases by 1 atm every 10 m (33 ft). When ordering transmitters, be sure to request units that have been tested beyond the pressures to which the study animal is expected to dive. The range of VHF telemetry systems is limited. As a rule of thumb, assume that the maximum reception ranges will be less than 5 km (3 miles) from boat or shore to the tagged animal. Greater reception ranges can be achieved by increasing the height of the receiving antenna. For example, when tracking from an airplane, ranges of more than 30 km (18 miles) are possible. The greatest challenge faced when radio-tracking most marine mammals results from the limited time they spend at the water’s surface, because radio signals can only be heard when the transmitter’s antenna is near or above the surface of the water. There are other disadvantages of radio telemetry techniques: they are very time intensive; the number of animals that can be studied at any one time may be limited (Kenward, 1987); there may be high costs associated with locating the animal; and adequate sampling is often constrained by weather conditions, darkness, safety considerations, and extensive animal movements (Harris et al., 1990). Although many transmitters can be equipped with mortality, temperature, and activity sensors, the amount of behavioral, ecological, and physiological data that can be collected is still limited. Archival tags are self-contained instruments that are capable of logging and archiving signals from a wide variety of sensors. Various data, such as temperature, salinity, position, and water depth, can be downloaded upon retrieval of the instrument (Bengtson, 1993). For example, time-depth recorders (TDRs), first devised for Weddell seals (Leptonychotes weddellii) during the 1960s (Kooyman, 1965), collect and store time-stamped pressure data, and thus provide a means to record the continuous diving behavior of tagged animals. Time-depth recorders are optimized to gather and record digitally large amounts of data on the behavior of diving animals. The memory capacity and resolution of these instruments have improved drastically over time. Many parameters, including depth and duration of dives, environmental temperature, vocalization recordings, swim velocity, and light level (which can ultimately be used for determining geographic location) can be sampled at preprogrammed time intervals. Some microcomputer packages can simultaneously measure and record heart rate and body temperature in conjunction with diving behaviors (e.g., see Andrews, 1998a). Communications are performed through a serial interface to a computer, and instruments are managed via user-definable options and parameters to suit data collection needs (Hill, 1997). Other references should be reviewed to obtain information for sampling designs (Boyd, 1993; Boveng et al., 1996).
0839_frame_C38.fm Page 854 Tuesday, May 22, 2001 2:01 PM
854
CRC Handbook of Marine Mammal Medicine
TABLE 1 Contact Information for Some Tag Manufacturers (listed alphabetically) and a General List of Products Commonly Used for Tagging and Tracking Marine Mammals Company and Address
Telephone and Fax Numbers, Web Site, and E-mail
Products
Advanced Telemetry Systems, Inc. 470 First Ave North P.O. Box 398 Isanti, MN 55040 U.S.A.
Tel: 763-444-9267 Fax: 763-444-9384 Web: http://www.atstrack.com E-mail:
[email protected]
Transmitters Data collection computers Global positioning system (GPS) collar systems Receivers and antennas
Holohil Systems Ltd. 112 John Cavanagh Road Carp Ontario Canada K0A 1L0
Tel: 613-839-0676 Fax: 613-839-0675 Web: http://www.holohil.com E-mail:
[email protected]
Radio transmitters
Lotek 114 Cabot Street St. John’s, Newfoundland Canada A1C 1Z8
Tel: 709-726-3899 Fax: 709-726-5324 Web: http://www.lotek.com E-mail:
[email protected]
Transmitters GPS Time-Depth Recorders (TDRs) Receivers
Sirtrack Ltd Private Bag 1403 Goddards Lane Havelock North New Zealand
Tel: +64 6 877 7736 Fax: +64 6 877 5422 Web: http://Sirtrack.landcare.cri.nz E-mail:
[email protected]
Transmitters Platform Transmitter Terminals (PTTs) Receivers and antennas
Sonotronics, Inc. 1130 E. Pennsylvania St., Suite 505 Tucson, AZ 85714 U.S.A.
Tel: 520-746-3322 Fax: 520-294-2040 Web: http://www.sonotronics.com E-mail:
[email protected]
Coded tags Pulse interval tags Ultrasonic receivers Hydrophones
Telonics 932 E. Impala Ave. Mesa, AZ 85204 U.S.A.
Tel: 480-892-4444 Fax: 480-892-9139 Web: http://www.telonics.com E-mail:
[email protected]
Telemetry subsystems Argos subsystems GPS subsystems Receivers and antennas
Trac-Pac, Inc. 251 Racetrack Road NE Fort Walton Beach, FL 32547 U.S.A.
Tel: 850-864-1857 Fax: 850-863-3980
Trac-Pacs
Wildlife Computers 16150 NE 85th Street 226 Redmond, WA 98052 U.S.A.
Tel: 425-881-3048 Fax: 425-881-3405 Web: http://www.wildlifecomputers.com E-mail:
[email protected]
Satellite-linked tags TDRs Additional archival instruments (heartrate/stomachtemperature recorders, heart-rate transmitters, and stomachtemperature pills)
Because TDRs must be recovered to retrieve data, they work best for animals that return to predictable locations, such as rookeries or haul-out sites (Stone et al., 1998). If animals cannot be recaptured, a variety of methodologies have been used to retrieve TDRs, ranging from attachment methodologies using corrodible bolts to release devices (Westgate et al., 1995; Baranov,
0839_frame_C38.fm Page 855 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
855
1996; Andrews, 1998a and 1998b; Croll, 1998; Eguchi, 1998; Hammill et al., 1999; Hooker and Baird, 2000). However, many methodologies are in the early stages of development, and are unreliable and expensive. Archival pop-up tags, which have been used primarily on pelagic fish in the past, collect and store data into memory. These data are then transmitted via satellite after the buoyant tag is released from the animal by an automatic time-delay switch (Stone et al., 1998). Archival pop-up tags are now available for marine mammals (Hill, pers. comm.). Satellite telemetry is becoming more widely used for tracking marine mammals. Satellite transmitters emit powerful ultrahigh-frequency (UHF) radio signals (401.650 MHz) to orbiting satellites and hence do not require conventional radio tracking (Harris et al., 1990). Satellite telemetry can thus be used for tracking species that are located in remote areas where conventional tracking costs would be prohibitive, or in situations where animals are pelagic or have large home ranges (Harris et al., 1990). Satellite telemetry also provides data periodically throughout the day and night, and during adverse weather conditions. Furthermore, no fieldwork is necessary after tagging to collect data, although close-range observations are necessary to document tag attachment and behavior (Mate et al., 1995). Satellite-linked transmitters that operate through the Service Argos system are termed Platform Transmitter Terminals (PTTs). Service Argos, Inc. (
[email protected] in Europe, or
[email protected] in North America) is a satellite-based location and data collection system that was established during 1978, under an agreement among the National Oceanic and Atmospheric Administration (NOAA), the National Aeronautics and Space Administration (NASA), and the Centre National d’Etudes Spatiales (CNES, the French space agency). The Argos receivers are carried on board NOAA polar-orbiting environmental satellites ∼850 km (∼510 miles) above the Earth (Figure 1). At least two satellites are simultaneously in service on sun-synchronous, polar, circular orbits, providing full global coverage (Argos, 1996).
FIGURE 1 Illustration of the Service Argos system (from Service Argos, Inc.).
0839_frame_C38.fm Page 856 Tuesday, May 22, 2001 2:01 PM
856
CRC Handbook of Marine Mammal Medicine
The Argos centers calculate transmitter locations by measuring the Doppler shift of the transmitted frequency as the satellite orbit passes over the transmitter (Kenward, 1987; Fancy et al., 1988). Two or more signals must be received during a single pass of the satellite to calculate the animal’s location. Data from the Argos archive are available via media products (i.e., floppy disk, CD-ROM), and results can be retrieved from anywhere in the world using public data networks (Argos, 1996). Researchers who are interested in using satellite telemetry are required to submit a program application form to Service Argos, which recommends submitting the application a few months before any planned deployments. This allows time for application approval, correction of the program if necessary, and transmitter testing. Some program information (i.e., PTT identification numbers, data type, and repetition rates) will need to be conveyed to the instrument manufacturers. Most vendors need at least 8 weeks notice, and transmitters that are custombuilt for specific applications may take additional time to produce. Prior to deployment, new users also should allocate some time to familiarize themselves with new instruments. The decreased size and greater capability of satellite tags has led to a significant increase in their use on marine mammals. Satellite transmitters used in the Argos system transmit messages once every 55 to 90 s, although most PTTs deployed on marine mammals transmit once every 45 to 60 s. Power output generally ranges from 0.25 to 1.0 W, depending on the model and battery configuration. Battery choice is contingent upon size constraints of the instrument and temperature of the water where the instrument will be deployed. Standard PTTs may be cast in epoxy or sealed in aluminum or titanium housings, which are designed to withstand hydrostatic pressure. Because UHF radio signals rapidly attenuate in salt water, PTTs are equipped with a saltwater switch that conserves battery power by synchronizing transmissions with surfacings and suppressing transmissions when the animal is underwater. Data collected from Service Argos vary, depending on the service agreement and PTT model. Service Argos allows 256 bits of information to be transmitted with each message. Transmitter manufacturers, therefore, have built additional sensors into their tags that allow additional data to be collected and transmitted. For example, behavioral data, or sensor data, in addition to at-sea and on-land locations, are encoded into messages emitted from satellite-linked timedepth recorders (SDRs; see Wildlife Computers in Table 1). Summary information on maximum dive depths, dive durations, and the amount of time animals spend at certain depths (“time-at-depth”) are collected and encoded into histograms, which consist of a set of bins (or accumulators) that contain counts for a given range of depth or time (Hill, 2000). The counts are accumulated for a “histogram period,” which is usually defined for 6 hours. Analysis software, which consists of a set of integrated programs and files to decode and format the data that are provided from Service Argos, is either supplied with a purchase or may be downloaded from the Internet (see Table 1). In addition to a variety of sensor data requested by the user, PTT number, date, time, latitude and longitude in decimal degrees, number of transmitter messages received, satellite identity, and location quality or class are collected. The accuracy of locations obtained through satellite telemetry is important when trying to examine an animal’s movements (Goulet et al., 1999). Service Argos provides an estimate of accuracy for each location class (LC): LC 3 = ±150 m, LC 2 = ±350 m, LC 1 = ±1 km, and LC 0, A and B = unrated (Argos, 1996). Factors that affect the quality of location estimates include movement, latitude, and altitude of the animal, antenna type and attachment, overpass elevation, oscillator stability, ionospheric propagation, transmitter power, number of uplink messages received, and the number of Doppler measurements obtained during each satellite overpass (Kenward, 1987; Stewart et al., 1989; Mate et al., 1995; Argos, 1996). Because locations determined from PTTs on marine mammals are less accurate than those predicted by Service Argos (Stewart et al., 1989; Mate et al., 1992), data generally need to be examined for erroneous
0839_frame_C38.fm Page 857 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
857
positions (e.g., position estimates that place an animal inland or well outside the study area), logical consistency, and unreasonable movements. Filtering algorithms and error indices have been developed to enable researchers to exclude locations with large errors (McConnell et al., 1992; Keating, 1994). Screening procedures are used to ensure that distances between locations are realistic, given the elapsed time between the fixes and an estimated swim speed of the animal being studied (Loughlin et al., 1987; McConnell et al., 1992; Mate et al., 1995; Merrick and Loughlin, 1997; Read and Westgate, 1997). For example, a maximum swim speed of 10 km/hour (6 mph) between locations is acceptable for many species (Loughlin et al., 1987; Merrick and Loughlin, 1997; Lowry et al., 1998; Wells et al., 1999; Melin, pers. comm.). There are a few options available to increase battery life, and thus longevity, of satellite tags. For example, the duty cycle of an instrument defines periods during which the instrument transmits vs. periods of temporary shutdown. Transmitters may be programmed to temporarily shutdown after an animal comes ashore for a user-specified period. A satellite transmitter’s duty cycle affects the probability of location determination and successful data transmission to the satellites, operational life, and data collection costs (Burger, 1991). Optimum duty cycles should be programmed according to study objectives and the timing and elevation of satellite passes over the study area (Harris et al., 1990). The number of satellite passes per day ranges from 7 at the equator to 15 at higher latitudes (Kenward, 1987). Telonics’ (see Table 1) Real-Time Satellite Display (TSD) and Satellite Predictor (TSP) program is a DOS-based application that provides a variety of satellite overpass prediction and color display features for use on IBM-compatible computers. J Pass 2.0 (http://Liftoff.msfc.nasa.gov/realtime/jpass/20), provided by NASA’s Marshall Space Flight Center, can be accessed for pass details or links to other programs. Marine mammals may also be tagged with a Global Positioning System (GPS), which receives positional data from a constellation of 24 satellites carrying transmitters (Tomkiewicz, 1996). Although still in its infancy, GPS is advantageous in that it is more accurate than Service Argos, allows for 24-hour coverage, provides extremely accurate time-stamping of position fixes, and users can establish the interval at which position fixes are acquired (Tomkiewicz, 1996). Data may be retrieved from a GPS by recovering the instrument after recapturing the animal, or by relaying the information from a transmitter on the animal to a receiver. If a GPS receiver is interfaced or built into a PTT, a processing module at the Argos centers can extract the GPS positions from Argos messages, validate them, and output them as regular Argos positions (Argos, 1996). Unfortunately, the various tracking methodologies listed above do not allow for direct observations of animals at depth (Davis et al., 1999). However, underwater video systems, or “Crittercams” (Parrish et al., 2000), have recently allowed biologists to directly observe behaviors that are otherwise typically inferred by use of transmitters or archival instruments (Williams et al., 2000). To date, hunting, foraging, swimming, and locomotor behaviors have been documented for some pinnipeds and cetaceans using these animal-borne video cameras (Davis et al., 1999; Skrovan et al., 1999; Tully, 1999; Parrish et al., 2000; Williams et al., 2000, Harvey, pers. comm.).
Pinnipeds The choice of tag and tracking system for pinnipeds depends upon the species, the objectives of the study, and the ability to restrain the animal (see Chapter 26, Ultrasonography; Chapter 41, Seals and Sea Lions). In the United States, the National Marine Fisheries Service (NMFS) requires that all rehabilitated pinnipeds be flipper-tagged when released, and resightings of these tags have been used to assess the efficacy of rehabilitation programs (Stoddart, 1978; Harvey et al., 1983;
0839_frame_C38.fm Page 858 Tuesday, May 22, 2001 2:01 PM
858
CRC Handbook of Marine Mammal Medicine
Webber and Allen, 1986; Seagars, 1988; Gavette, 1992). Cattle and sheep ear tags (e.g., jumboroto, metal monel, and temple tags) are most commonly used to flipper-tag pinnipeds (Allflex New Zealand Ltd, 931 Tremaine Ave, Private Bag, Palmerston North, New Zealand; Daltons Animal Identification Systems Ltd, Henley-on-Thames, Oxford, U.K.). These embossed numbered tags are relatively inexpensive and can be personalized and color-coded. The tags are applied using applicators provided by tag manufacturers so that the trailing edge of the tag is flush with the edge of the flipper; phocids are tagged in the webbing of the rear flippers, whereas otariids are tagged in the fore flippers. Although flipper tags are commonly used, they may be inadequate for long-term monitoring because the painted numbers may rub off and the color of the plastic may change over time with exposure to sunlight. Furthermore, the small numbers on flipper tags are difficult to read from a distance and sand, mud, or other animals may obscure visibility of tags. Flipper tags are also lost quite frequently. Retention rates and effectiveness of different types of flipper tags are summarized in Testa and Rothery (1992) and Jeffries et al. (1993). Brands provide better resighting information than flipper tags (Merrick et al., 1996). Freeze branding involves permanently marking animals with cold branding irons or liquid nitrogen sprays, resulting in unpigmented or bald brands. Freeze branding, however, has some limitations. For example, it is difficult to distinguish unpigmented hair against a light pelage, and freeze brands may disappear over time and with molting. During hot branding, heated steel irons are applied to the pelage to kill hair follicles and pigment-producing cells for a permanently bald brand (Merrick et al., 1996). Although branding is a quick and reliable method for permanently marking free-ranging California sea lions (Zalophus californianus), Steller sea lions (Eumetopias jubatus), elephant seals (Mirounga angustirostris), Weddell seals, gray seals (Halichoerus grypus), ringed seals (Phoca hispida), and harbor seals (P. vitulina), it is a controversial technique because it is difficult to assess degree of pain in wild animals. Alternative tagging methodologies, therefore, are commonly used to monitor pinnipeds. As a rule of thumb, tags should be attached where they are most exposed when the animal surfaces (Kenward, 1987). For that reason, smaller tags are commonly attached to the heads of pinnipeds. For example, “head tags” (Daltons Animal Identification Systems Ltd) (Figure 2), which were originally designed by the Sea Mammal Research Unit, Scotland, are glued directly to the pelage of the head. The tags are made of a high-impact styrene, and filled with a mixture of microspheres and a polyester casting resin, which makes them buoyant, impervious to pressure, and strong enough to withstand abrasion (Hall et al., 2000). They weigh 30 g, can be produced in several bright colors, and have a large two-digit number embossed on each
FIGURE 2 Head tag glued to the head of a harbor seal. (Photo credit: Jason Waite.)
0839_frame_C38.fm Page 859 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
859
side, enabling identification of individuals in the field (Hall et al., 2000). Head tags have been used successfully on rehabilitated gray seal pups and free-ranging harbor seals, and numbers can be read reliably at distances up to 50 m (∼165 ft) using binoculars (Hall et al., 2000; Greig, pers. comm.). Before gluing the tag to the head, the pelage is cleaned with acetone, methylated spirit, or methyl hydrate and dried with compressed air (Fedak et al., 1983; Ellis and Trites, 1992; Jeffries et al., 1993; Hall et al., 2000). Clean towels can be used to remove excess dirt and moisture from the fur. Quick-setting epoxies (e.g., Devcon® products, Devcon, Riviera Beach, FL and Titan Corporation, Lynwood, WA) or cyanoacrylic adhesives (e.g., Loctite® products, Loctite Corporation, Rocky Hill, CT; Mississauga, Ontario, Canada; Mexico City, Mexico) are commonly used to attach tags to the pelage of pinnipeds (Jeffries et al., 1993; Le Boeuf et al., 2000; RaumSuryan, pers. comm.). In contrast to epoxy, which becomes hot and may take 5 to 15 min to harden, Loctite 422 and 380 bond quickly to the fur (<1 min) and generate minimal heat (Jeffries et al., 1993). Loctite 4212 takes longer to set, but supposedly fills the interstitial spaces within the fur (Raum-Suryan, pers. comm.). However, cyanoacrylic adhesives degrade in water over time, whereas epoxies are more resistant to degradation in salt water (Bengtson, 1993). The use of telemetry techniques emerged during the 1960s as a powerful new tool for studying behavior, movements, and physiology of pinnipeds (Bengtson, 1993). Over the succeeding 40 years, several different types of tags and attachment schemes have been developed. Unfortunately, instruments that are glued to the fur are eventually lost during the molt, so permanent marks are still the only way to acquire long-term data. Some seals have been fitted with VHF radio transmitters attached to bracelets around the base of their hind flippers (Pitcher and McAllister, 1981; Bengtson, 1993; Jeffries et al., 1993), although these methodologies cannot be applied to all animals. For example, there is no constriction at the ankle of gray seals, where bracelets could be strapped on (Fedak et al., 1983). Because anklets do not allow for growth, they cannot be deployed on juveniles without the addition of a biodegradable link. Although anklet attachments are advantageous in that they are not lost during the molt (provided a biodegradable link is not used), Pitcher and McAllister (1981) found that excess rigidity of the bracelet caused some seals to develop abrasions where the bracelet encircled the ankle. If used, bracelets should be fastened such that an index finger can be slid between the bracelet and the seal’s ankle (Bengtson, 1993). Radio transmitters have also been attached to temple tags (Temple Original®, Temple Tag Co., Little River, TX; Figure 3), which are then attached to the hind flipper. Although not lost during the molt, radio flipper tags tend to migrate and tear the webbing of the flipper. Additionally, studies have indicated that some pinnipeds, especially juveniles, are quite active after release, so that attachment of the radio transmitter to the flipper or ankle may result in the animal not being detected while in the water. Placement of the radio tag on the head provides a reliable signal when the seal surfaces to breathe (Fedak et al., 1983; Harvey, 1987). Diving and respiratory patterns, as well as foraging areas, can also be documented with head-mounted radio tags. Radio transmitters are typically attached to the head, nape of the neck, or back (Figure 4) of pinnipeds using the same methods as those used for the head tags listed above (Fedak et al., 1983; Jeffries et al., 1993; Merrick and Loughlin, 1997). The radio tag can be glued directly to the fur on the animal, or the surface area of the bottom of the transmitter can be increased by attaching it to a piece of neoprene using glue or cable ties. Radio transmitters that are cast in metal housings with metal tabs can be bolted to a piece of neoprene (Jeffries et al., 1993). The antenna can be applied backward or forward, but it should be oriented in a manner to improve signal propagation (Guinn and Lee, 1996). Many researchers prefer to tag pinnipeds with the antenna facing toward the posterior end of the animal. However, it may be beneficial to orient the antenna toward the head if the animal displays behaviors that would hinder signal reception. For example,
0839_frame_C38.fm Page 860 Tuesday, May 22, 2001 2:01 PM
860
CRC Handbook of Marine Mammal Medicine
FIGURE 3 A Temple tag containing a radio transmitter is illustrated in the larger photograph. (Photo credit: Michael Weise.) Temple tags (∼5 cm) are also shown in the upper right corner. The number is prominent, but may wear off over time and may be difficult to read from a distance. (Photo credit: Michael Murray.)
FIGURE 4 California sea lions tagged with VHF transmitters on their heads and satellite transmitters on their backs. (Photo credit: The Marine Mammal Center.)
some seals tend to tip their head back while bottling (resting upright) at the surface. If the transmitter is placed on the head with the antenna facing forward, keeping in mind that power output varies with antenna dimensions (Kenward, 1987), the length of an antenna should be adjusted to avoid poking the animal in the eyes. Satellite telemetry is a preferred means for tracking remote, pelagic, and migratory species. Attachment methods and the length and positioning of the antenna on the animal are critical for maximizing the number of uplinks, and therefore data transmitted (Stewart et al., 1989). Satellite transmitters are usually attached to the fur of pinnipeds on the midline over the scapulae using marine epoxy (Fedak et al., 1983), although some animals have been tagged on the lower back (Schreer and Testa, 1996) or head (Le Boeuf et al., 1996; 2000). As with radio transmitters, the surface area of the base of the satellite transmitter can be increased. Satellite transmitters have been attached to the backs of Steller sea lions using epoxy applied to a thin mesh spread (Merrick et al., 1994) (Figure 5), and to the heads of northern elephant seals using marine epoxy spread over a nylon matrix (Le Boeuf et al., 1996). Additionally, PTTs
0839_frame_C38.fm Page 861 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
861
FIGURE 5 Satellite-linked time-depth recorder attached to the dorsum of a Steller sea lion using epoxy applied to a piece of mesh. (Photo credit: National Marine Mammal Laboratory.)
have been fastened with two cable ties and a stainless-steel hose clamp to a nylon seine net (0.48 cm mesh) glued to the head (Le Boeuf et al., 2000). Satellite transmitters also have been glued directly to the pelage of fur seals (Callorhinus ursinus and Arctocephalus sp.) and California sea lions using epoxy (Walker and Boveng, 1995; DeLong, pers. comm.) (see Figure 4). For best results, sandpaper or a metal file can be used to roughen the base of the instrument to facilitate adhesion. Use of epoxy should be limited, because it may burn the pelage and skin beneath the instrument. In addition to damaging the fur, excess glue also increases the weight of the package and may cause the hair to break, resulting in tag loss. The pelage of captive animals, which are housed in pools containing ozone, bromine, chlorine, and other bleach products, is especially prone to damage from glue. Transmitters have been attached to the tusks of walruses (Odobenus rosmarus) using stainless-steel bands (Wiig et al., 1993; Born and Knutsen, 1997), and antennas should be adjusted so that they do not interfere with the eyes or whiskers (Harris et al., 1990). Time-depth recorders have been deployed on many pinniped species (Boyd and Croxall, 1996). In earlier studies, hose clamps were used to attach TDRs to harnesses made of tubular nylon webbing (Gentry and Kooyman, 1986; Feldkamp et al., 1989), but because this increased drag, performance, and swim velocity, other attachment methods are now used. For example, Boyd and Arnbom (1991) glued a piece of nylon webbing (30 × 2 cm) to the dorsal midline, slightly posterior to the scapulae, of southern elephant seals (Mirounga leonina). Two hose clamps, distanced 10 cm (4 in.) apart, were looped around the webbing before attachment and allowed to cure in the glue. After the glue dried, a cylindrical TDR was screwed into the hose clamps, so that the long axis of the instrument was parallel with the long axis of the seal. McCafferty et al. (1998) used similar methodologies to study the foraging behavior of Antarctic fur seals (Arctocephalus gazella). Plastic cable ties were used to attach TDRs to a nylon-webbing strap that had been glued to the mid-dorsal fur. Le Boeuf et al. (2000) used a combination of stainless-steel hose clamps and cable ties to attach dive recorders to elephant seals on the dorsal midline above the shoulders using the same methods as they did for the PTTs described above. Boness et al. (1994) embedded hose clamps (which later secured a TDR), a radio transmitter, and a piece of nylon mesh into an epoxy mount. The nylon mesh, which had been embedded in the base of the mount, was epoxied to the fur of harbor seals. The radio tag enabled researchers to locate, recapture, and retrieve TDRs. Inclusion of a radio transmitter in the epoxy mount also allows for recovery of
0839_frame_C38.fm Page 862 Tuesday, May 22, 2001 2:01 PM
862
CRC Handbook of Marine Mammal Medicine
the TDR if the mount detaches from the animal while on shore. Similar epoxy mounts have been used successfully on California sea lions (Melin, pers. comm.) and elephant seals (DeLong and Stewart, 1991; Stewart and DeLong, 1995). For species that are not as easily recaptured, TDRs and radio transmitters can be encased in buoyant packs made of micro-balloon-glass bubbles and epoxy (Ellis and Trites, 1992) or syntactic foam (Flotation Technologies, Biddeford, ME; Eguchi, 1998). Release devices can then be used to retrieve the packs. Animal welfare and well-being should always be considered before conducting a tagging study, and instrument effects must be considered during data analysis and interpretation. Unfortunately, few studies have been conducted to examine the adverse effects of tagging pinnipeds. It is often difficult to design an appropriate experiment to study the impacts of tracking devices, because they are used to collect data that are difficult to obtain in any other way (White and Garrott, 1990), and that makes it difficult to have controls. A mark–recapture study of northern fur seals tagged with metal monel tags indicated the return rates of nontagged animals were greater than for tagged animals (York, 1989). Walker and Boveng (1995) found that the average duration of foraging trips and nursing visits for Antarctic fur seals was greater for seals that were tagged with TDRs and radio transmitters than for seals tagged with radio transmitters alone. In contrast, Baker and Johanos (1999) found that telemetry devices had no deleterious effects on individual free-ranging Hawaiian monk seals (Monachus schauinslandi).
Cetaceans Cetaceans lack hair, are very streamlined, and do not exhibit haul-out behavior, which makes them very difficult to tag. Despite these hurdles, biologists have succeeded in deploying a wide variety of tags on both mysticetes and odontocetes. A wide variety of passive tags have been used on cetaceans with varying degrees of success, including cattle ear-tags, spaghetti tags, buttons tags, and freeze brands (see Scott et al., 1990, for review). Cattle ear or roto-tags (Jumbo Roto Tag®, Dalton Supplies, Nettlebed, U.K.) have been used extensively in field studies of bottlenose dolphins (Tursiops truncatus) (Scott et al., 1990). They have proved effective for short-term identification of individuals, although photographs often are necessary to actually read roto-tag numbers. Roto-tags are attached close to the trailing edge of the dorsal fin and often migrate out of the tissue, leaving a notch. By attaching roto-tags in a selective manner, unique notches can be made on individual dorsal fins. Roto-tags are attached using special pliers supplied with the tags by the manufacturer. Freeze branding is considered the best way to mark individual cetaceans permanently. These marks have remained legible on individual bottlenose dolphins for up to 11 years (Wells, pers. comm.). Effective freeze brands are achieved by applying the supercooled branding irons to the epidermis for a set period of time. Marks are usually made on the dorsal fin as well as on the lateral flank. The time interval for the brand to be applied to the skin is very species specific. There are three types of telemetry tags used on cetaceans: acoustic, VHF, and satellite. The type of telemetry used will be dictated by the animal, the research goals, and the budget. Acoustic telemetry has not been used widely in the field because acoustic transmitters that work efficiently in seawater do so at frequencies (10 to 100 kHz) that can be heard by some cetaceans (Richardson, 1995), and this may influence the behavior of the tagged animal. Because signals from higher-frequency transmitters attenuate very quickly in seawater, it would not be feasible to use an acoustic transmitter that transmitted above the hearing level of most odontocetes (>200 kHz). Acoustic transmitters do have a major advantage over other types of telemetry devices, because they can transmit through water. This means that it is possible to track and record data from animals while they are swimming underwater. However, wildlife acoustic
0839_frame_C38.fm Page 863 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
863
transmitters have a limited transmission range of a few kilometers or less, which makes tracking cetaceans challenging. Since the 1960s, VHF radio telemetry has been used to study the movements of wild cetaceans, resulting in many valuable insights (Evans, 1971; Gaskin et al., 1975; Irvine et al., 1982; Mate et al., 1983; Harvey and Mate, 1984; Mate and Harvey, 1984; Read and Gaskin, 1985). Attachment techniques have varied, but usually involved attaching tags directly to the dorsal fin. Early VHF packages were large and cumbersome (Evans, 1971) and covered most of the dorsal fin, but modern VHF tags can be as small as a thimble. One of the problems encountered during VHF studies was short deployment times. The reasons for this were not always clear, but were assumed to be caused by either transmitter failure or animals moving out of the limited tracking range of the VHF receivers. VHF telemetry is thus most suited for species whose general home range is known or when continuous tracking is planned. Satellite tags are especially suited for application to cetaceans because they transmit remotely to orbiting satellites and, as such, do not require field-based monitoring, operate around the clock, and during all weather conditions. Satellite transmitters have decreased in size considerably since their introduction in the mid-1980s (Mate, 1989) and these tags have been successfully deployed on cetaceans as small as harbor porpoises (Phocoena phocoena) (Read and Westgate, 1997) and as large as blue whales (Balaenoptera musculus) (Mate et al., 1999). In addition to telemetry devices, data loggers have been deployed on cetaceans. To date, TDRs have been the most common type of data logger used and there have been published accounts from a wide variety of species (Hooker and Baird, 2000). Time-depth recorders are attached to cetaceans directly (Westgate et al., 1995) or by way of suction cups (Hooker and Baird, 1999). One of the major challenges faced when deploying data loggers is recovering the units once the deployment is over. This is usually accomplished by incorporating the data logger with a VHF transmitter into a buoyant package. This enables the tagged animals to be tracked and the loggers recovered after they have detached. The attachment methods that are used for any tag will be dependent on the species being tagged, the objectives of the study, and the duration of the deployment. Many cetacean species have never been tagged and techniques have not been developed. Thus, more effort will be required before a safe and effective tag can be deployed. If the goal is data collection for a few days, the best system to deploy is one that attaches the tag to the animal using suction-cups. There are two types of suction-cup-based attachments available. The first relies on a single large suction cup to which the tag is attached either directly or by a tether. This kind of tag is suited for remote deployment from either crossbows or poles. Obvious reactions to these tags have been documented in bottlenose dolphins (Schneider et al., 1998), so they may not be applicable to all species. The second type of suction-cup-based tagging system, called the Tracpac® (Trac-Pac, Inc., see Table 1), was developed for use on bottlenose dolphins (Figure 6). This unique package relies on a series of smaller suction cups that are attached to a hinged thermoplastic saddle. The suction cups hold the saddle to the dolphin’s dorsal fin. The saddle is conformed to the shape of the dorsal fin and contains special pockets to hold the transmitters or data loggers. One limitation of this system is that free-ranging animals must be captured before these devices can be attached. Both methods are best suited for short-term deployments (usually less than 1 day) where animals will be tracked continuously using VHF telemetry. Another short-term tagging method that is commonly used is the roto-radio, which combines a small, epoxy-encapsulated VHF transmitter with a cattle-ear roto-tag. The roto-tag can be attached to the trailing edge of the dorsal fin to provide a quick, reliable, and less invasive way to attach VHF radios to cetaceans. The system is safe because it only requires a single small hole that is located near the trailing edge of the dorsal fin, but like Trac-pacs, requires that the
0839_frame_C38.fm Page 864 Tuesday, May 22, 2001 2:01 PM
864
CRC Handbook of Marine Mammal Medicine
FIGURE 6 Trac-pac on a bottlenose dolphin. The tag is held in place by a series of small suction cups that line the inside of the package. In this view the VHF radio and TDR can be seen. (Photo credit: Chicago Zoological Society, Sarasota Dolphin Research Program.)
animal be captured. Roto-radios tend to have shorter life expectancies than traditionally mounted systems, with typical deployments lasting approximately 25 to 30 days. When longer deployments are required, it is necessary to attach the tag using more invasive methods. This usually requires attachment by way of pins through the dorsal fin, or for larger cetaceans, by implanting the tag into the blubber layer. Because these methods could potentially harm animals, they should be approached with caution and by someone who has previous experience. Generally speaking, tags are not usually available from manufacturers in a configuration that is ready to deploy. Although Wildlife Computers manufactures a fin mount that has been used on various cetaceans (Figure 7), packages typically have to be customized by the purchaser. Most tags attached to odontocetes use pins that go through the tissue of the dorsal fin (Scott et al., 1990; Westgate et al., 1995; Read and Westgate, 1997) or dorsal ridge (Martin and Smith, 1992) (Figure 8). These pins are used either to attach the tag directly or to attach a saddle containing the tag. When securing a tag to a dorsal fin using pins, the quantity, size, and placement of the pins should be considered. A greater number of pins provide increased stability, but there is also an increased risk of damaging the vasculature of the dorsal fin. Too few pins may be safer for the animal, but may not provide enough support for the tag. As a general rule, use as few pins as possible to provide a secure attachment. Pin diameter is also important. Small pins have an increased tendency to break or migrate out of the tissue, whereas large pins are more likely to impact a major vessel. Generally, use 6.5-mm-diameter pins for smaller odontocetes such as harbor porpoises and 8.0-mm-diameter pins for larger animals such as bottlenose dolphins. Delrin or nylon has been commonly used as pin materials because each is strong and has good biocompatibility (Scott et al., 1990). Extreme caution should be exercised if stainless-steel bolts are used when tagging cetaceans; such bolts can result in serious fin damage by migrating out of tissue (Irvine et al., 1982). Pins should be secured with some bimetallic combination of nuts and washers because these will eventually corrode. The in situ dynamics of bimetallic corrosion is not well understood. All nuts should be backed with oversized delrin or nylon washers and open cell foam to reduce the effects of localized pressure necrosis. The tightness of the package on the dorsal fin is difficult to gauge. Overtightening can lead to tissue necrosis and undertightening can allow the tag to move around, which may lead to unnecessary tissue abrasion. Holes for pins usually are cut using specialized
0839_frame_C38.fm Page 865 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
865
FIGURE 7 A fin-mount containing a SDR-T16 satellite-linked tag (using the Telonics ST-16 transmitter), designed and manufactured by Wildlife Computers. (Photo credit: Wildlife Computers.)
FIGURE 8 A Telonics ST-10 satellite transmitter attached to a harbor porpoise. (Photo credit: Grand Manan Whale and Seabird Research Station.)
0839_frame_C38.fm Page 866 Tuesday, May 22, 2001 2:01 PM
866
CRC Handbook of Marine Mammal Medicine
hole cutters. It is better to cut a hole that is slightly smaller than the pin that will be used because this will help reduce bleeding and reduce loosening of the tag should the hole size increase over time. Prior to any surgery, the site should be cleaned properly, a local anesthetic applied, and antibiotic cover considered. Implantable tags have been deployed remotely by crossbow or modified shotguns, usually on large whales (Goodyear, 1993; Watkins et al., 1993; Mate et al., 1999). The heads of these tags usually have cutting points that assist in penetrating the blubber. Implantable tags are sometimes designed so that just a small anchor penetrates the blubber layer to which the externally mounted tag is attached. To date, most implantable tags have contained satellite transmitters (Mate et al., 1997; 1999), but acoustic transmitters have also been deployed (Goodyear, 1993; Watkins et al., 1993). When designing implantable tags, it is important to have a good understanding of the blubber depth in the area that the tag will be attached, so the correct penetration will be achieved. Unlike tags that are attached using other techniques, implantable tags undergo extreme ballistic forces when deployed and need to be designed to withstand high impact. Tags and penetrating heads should be soaked in a disinfectant solution prior to implantation. To date, little attention has focused on the physical or physiological effects of tagging cetaceans. When placing tags on dorsal fins, it is important to remember that these appendages function as both hydrodynamic and thermoregulatory surfaces, and increased drag means increased energy expenditure for locomotion (Pabst et al., 1999). It is important, therefore, to design tags that minimize any disruption of normal water flow, especially for long-term deployments. Recent results indicated that saddle-based packages had more drag than packages without saddles, and increased drag may have contributed to decreased deployment time (Hanson et al., 1998). It is imperative to understand the vasculature of the dorsal fin, so a site reasonably devoid of vessels can be selected (see Chapter 9, Anatomy). This will minimize impact to important heat-exchanger vessels, as well as to the general blood supplies of the fin. This can be accomplished by examining dorsal fins of dead animals or by probing selected pin sites with a hypodermic needle prior to the actual surgery. Little is understood about the long-term fate of implantable tags. Whether short- or long-term deployments are planned, the actual deployment length is often far less than expected. Incorporating small changes to a tag design in an iterative manner is usually more successful than trying to effect many major changes quickly, because it is possible to learn from sequential trials.
Manatees Research on marking, tagging, and tracking manatees (Trichechus spp.) has been implemented since the 1960s and various methods have been adopted. Passive integrated transponder (PIT) tags (e.g., InfoPet Identification Systems®, Burnsville, MN), which typically function as a complement to other means of identifying individuals, have been employed in manatee research for the past 10 years (Wright et al., 1998). These tags are glass-encapsulated microchips (11.0 × 2.2 mm and 54 mg in weight) that are programmed with a unique identification code. The tags are surgically placed subdermally into the cutaneous space of the fat layer over both the right and left shoulder regions using a cut-down procedure and a modified injection syringe (Figure 9). There have been no observed ill effects of these tags in manatees and this system of simple mark–recapture methodology allows researchers to positively identify a captured, stranded, or dead manatee at a later time. The electronics have an extensive life span because they are passive until activated by a handheld scanner (Thomas et al., 1987). Additionally, PIT tags are relatively inexpensive, small, accurate, cause minimal chance of infection, and can be retrieved and reused (Thomas et al., 1987).
0839_frame_C38.fm Page 867 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
867
FIGURE 9 PIT tag and applicator. Note the modification of the syringe plunger to facilitate placement of the transponder in the subcutis. (Photo credit: Michael Murray.)
Freeze branding also has been used to help facilitate reidentification of manatees. Standardized 5-cm (2-in.) numeric brands are cooled in liquid nitrogen and held firmly over a clean, dry surface of the skin for approximately 30 s. Only manatees that do not have unique scar patterns are branded on each shoulder and dorsal tail stock. Numbers are assigned from a centralized computer database prior to branding. Extensive studies using radio telemetry have been conducted on manatees over the last few decades. Throughout the years many attachment systems have been used, but currently, two principal types of tag configurations are used. Early projects employed a belt-mounted VHF transmitter for use in freshwater systems and were first introduced during the late 1970s (Bengtson, 1981; Irvine and Scott, 1984). Since that time, extensive modifications to the VHF transmitter housing have been developed that allow for use of a floating transmitter package. Conventional VHF radio-transmitters (Telonics, model MOD-550, 15-mW output into 50-Ω load), which emitted pulsed signals (25 to 65 ppm) at unique frequencies in the 164-MHz band, were installed in floating housings that were 33 cm long and 6 cm in diameter (Rathbun et al., 1987; Deutsch et al., 1998). These instruments were able to broadcast with a maximum effective range of approximately 15 km (9 mi) when tracking from land or sea and up to 50 km (30 mi) from air. These smaller floating units primarily were used during early years to locate animals visually or through triangulation in the field. Since that time, several modifications to the tracking assembly have been incorporated. Advances in technology and larger floating housings have allowed researchers to employ more powerful units. Satellite-monitored PTTs have become the predominantly used tag. Floating PTT units (Telonics, models ST-3 and ST-4, 1 W into 50-Ω load) are 39 cm long, 9 cm in diameter, and include a VHF transmitter (Telonics, model MOD-200 and MOD-225; 24 to
0839_frame_C38.fm Page 868 Tuesday, May 22, 2001 2:01 PM
868
CRC Handbook of Marine Mammal Medicine
FIGURE 10 Illustration of a GPS remote tracking system (Lotek, see Table 1) used on manatees. (Photo credit: United States Geological Survey.)
32 ppm, 164-MHz band) and an ultrasonic beacon (Sonotronics, Inc., see Table 1, model CHP87-L, 72-79 kHz) to aid researchers in locating manatees or detached units. Additional tracking methodologies utilizing GPS technology and cellular telephone communication equipment may be integrated into existing tracking packages (Figure 10). A noninvasive technique is used to attach radio tags to manatees. The current tag design includes an adjustable belt that attaches with a buckle around the caudal peduncle just cranial to the insertion of the tail paddle (Figure 11). The belts are constructed in various sizes and breaking strengths (Rathbun et al., 1987). Corrodible nuts and bolts allow the belt to release after as long as 4 years. Radio transmitters (Telonics, model MOD-550, 15 mW output into 50-Ω load) and ultrasonic beacons (Sonotronics, Inc., model CHP-87-L) have been incorporated into the belt assemblies, making it possible to locate manatees that have broken free of the floating transmitter or to recover detached belts. The floating transmitter is attached to the belt by a 1.5- to 2.0-m-long and 9.5-mm-diameter flexible nylon tether. Each tether has a weak link and is designed to break free if the transmitter becomes entangled. More-detailed technical descriptions of the manatee radio tag and belt assemblies are available in reports by Reid and O’Shea (1989), Rathbun et al. (1990), and Reid et al. (1995). Because this system has worked so well for manatees (T. manatus latirostris) in Florida, it has been modified and adapted for use on dugongs (Dugong dugon) in Australia and Saudi Arabia and manatees in Belize (T. m. manatus), Brazil (T. inunguis and T. m. manatus), Mexico (T. m. manatus), Puerto Rico (T. m. manatus), and West Africa (T. senegalensis). Prototypes of floating manatee radio tags have also been used successfully to temporarily track entangled large whales. The advantages for tagging a herbivorous, coastal marine mammal are obvious. Manatees commonly are found in shallow-water environments where floating transmitters usually are exposed at the surface. This affords excellent exposure of the antenna to air when the manatee is feeding or resting near the surface in salt or brackish water. When traveling, however, the transmitter often is pulled beneath the surface and radio signals are attenuated in salt water. Unfortunately, floating radio tags also are vulnerable at the surface and many are hit by boats, grabbed by alligators, or become snagged and detach. Tag life is limited and can be directly
0839_frame_C38.fm Page 869 Tuesday, May 22, 2001 2:01 PM
869
Tagging and Tracking
Antenna Antenna
Transmitter Transmitterhousing housing Nose Nose cone cone Chain Chain link linkconnector connector Peduncle Peduncle belt belt Tether Tether Buckle swivel Buckle and and swivel Tightening Tightening strap strap
Eye Eye bolt bolt Joiner Joiner
Chain link connector Chain link connector
FIGURE 11 Details of the belt assembly used on manatees. A satellite-monitored PTT tag is shown. VHF tags are similar in appearance, but the housing is smaller and has a longer antenna. (From Reid et al., 1995; photo credit: United States Geological Survey.)
0839_frame_C38.fm Page 870 Tuesday, May 22, 2001 2:01 PM
870
CRC Handbook of Marine Mammal Medicine
affected by battery life, duration of selected duty cycle, and sinking due to fouling with barnacles and other epiphytes. There have been few or no adverse effects of radio tagging observed in previously tagged manatees. However, care should be taken to assure that belts are not too tight, fit properly to allow for future growth of the animal, and are rated with the appropriate breaking strengths based on the animal’s age and size class. Minor scarring of tissue due to belt chafing along the anterior edges of the tail margin has been observed in some cases. Behavior does not appear affected and reproductive cycles do not seem to be interrupted. This noninvasive technique of monitoring a nearshore, coastal marine species is ideal and has been used successfully by many manatee biologists. For example, over 50,000 tracking days have been amassed from the 250 manatees tagged by the U.S. Geological Survey (USGS; Sirenia Project) during the last 20 years. One manatee was tagged intermittently for more than 6 years.
Sea Otters Tagging and tracking of sea otters (Enhydra lutris) may be problematic, despite their tendency to spend the majority of their lifetime in nearshore areas. The combination of the small external ear and the dexterity exhibited during normal grooming processes precludes use of ear tags. Branding interferes with the otter’s pelage and subsequent thermoregulatory abilities. Tattoos would be difficult to find and read in the densely furred otter. Despite these limitations, several techniques have been employed to facilitate identification and tracking of individual otters. Passive integrated transponder tags are small, 1- × 4-mm chips (Loomis, 1993) that were originally implanted in the subcutaneous tissue between the scapulae (Thomas et al., 1987). Because application was in close proximity to the otter’s mouth, PIT tags are now placed in the subcutaneous space in the caudal femoral region, to minimize risk to the handler. The fur over the right inguinal area may be parted with a 1:1 combination of sterile lubricating jelly and povidone-iodine solution. The implanting tool is a syringe/needle combination with a plunger extension that forces the PIT tag from its location within the bore of the needle. The tag should be tested by passing the scanning wand over the tag prior to and after insertion. Chip failure may occur and it is believed that rough handling of the chips may cause damage to the glass casing. There have recently been reports of soft-tissue sarcomas associated with transponder chip implantation in two small mammals in zoos (Pessier et al., 1999), although there have been no such reports in the sea otter. The most significant limitation to transponder chip use is its relatively short reading range of 5 to 38 cm (2 to 15 in.), depending upon the manufacturer, which necessitates having the animal in hand. The chip cannot be utilized to identify animals at a distance. Other marking techniques, therefore, are typically used in conjunction with PIT tags to facilitate the monitoring process. A variety of methods for attachment of flipper tags to sea otters have been attempted. The use of metallic tags was abandoned, as otters would frequently bend the tags, causing pressure necrosis of the underlying webbing and subsequent loss of tags. Plastic tags of a variety of colors and designs also have been employed. Techniques for anchoring tags with either doubleor single-puncture anchoring have been used (Ames et al., 1983). Currently, plastic tags (Temple brand) are attached to the hind flipper by a single-puncture technique. A 5-mm puncture is made through the webbing with a sterilized leather punch or Baker’s biopsy punch. The skin puncture must be placed such that tension is not created when the tag is inserted (puncture is too proximal), or such that there is too much space between the margin of the flipper and the tag (puncture is too distal). In young animals, the tag should be fitted to allow for growth. The plastic tag is inserted by its open end through the skin hole and then rotated 180° so that the open end is at the margin of the flipper webbing. The hole in the
0839_frame_C38.fm Page 871 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
871
open end of the tag is then filled with glue and a screw placed to secure the tag. By utilizing a variety of colors and color combinations, a number of individuals may be identified. Furthermore, tags may be placed in one of two locations: either the interdigital webbing between digits 4 and 5 or between digits 1 and 2 (Figure 12). Limitations to the use of the flipper tags include a limitation in the number of colors that can be distinguished at a distance; tags may be lost, either by damage to the interdigital webbing or by actual failure of the tag itself; and the otter and its tag must be within sight. The attachment of a radio transmitter to the tag within a smooth, waterproof nonreactive resin has been used as an adjunct to flipper tags (see Figure 12). These transmitters have enclosed batteries and antennae. They are attached in the same way as flipper tags; however, they are typically placed in the interdigital webbing between digits 2 and 3, or 3 and 4, to allow for greater stability and support, as well as placement of the traditional plastic tags. Flipper radio transmitters have the advantage of permitting tracking without having to resight the otter. Unfortunately, there are significant limitations to their use. Transmitters of excessive weight tear
FIGURE 12 Flipper tag and flipper transmitter on an adult male sea otter. The flipper tag is placed in the interdigital webbing between digits 1 and 2 and the transmitter between digits 3 and 4. This placement provides additional support for the heavier transmitter/tag combination. (Photo credit: Michelle Staedler.)
the interdigital webbing. Currently, transmitters that have a battery life span of 90 days weigh approximately 18 g. When the battery life was extended to 150 days, the weight increased to approximately 27 g and resulted in tearing of the flipper (Staedler, pers. comm.). Flipper radio transmitters typically are used for animals requiring short-term, relatively intensive monitoring. The most effective means of tracking sea otters involves the use of intra-abdominal radio transmitters. Surgical implantation of radio transmitters into the peritoneal cavity (Figure 13) has been used in several mustelids (Eagle et al., 1984; Hoover, 1984; Reid et al., 1986; Spelman et al., 1997; Stoskopf et al., 1997; Schwantje et al., 1998; Johnson and Berkley, 1999) and intraperitoneal implantation of radio transmitters has been used in sea otters by a number of investigators (Loughlin, 1977; Ribic, 1982; Garshelis and Siniff, 1983; Williams and Siniff, 1983; Ralls et al., 1989). Intra-abdominal radio transmitters are self-contained with batteries, antenna, and transmitter encased in a smooth, waterproof, nonreactive epoxy resin (Figure 14). Although there is some variation among transmitters, most are approximately 10 × 6 × 2.5 cm, and weigh between 140 and 180 g. In most cases the battery life span of the transmitter is 2 years, although duty cycling may prolong it. A mortality switch in the transmitter resulting in a change in pulse rate is activated when the body temperature
0839_frame_C38.fm Page 872 Tuesday, May 22, 2001 2:01 PM
872
CRC Handbook of Marine Mammal Medicine
of the otter decreases. The ranges of implanted transmitters are variable, with a maximum range of 0.8 to 1.2 km (0.5 to 1.0 miles) when the receiver antenna is at sea level during calm seas. By moving the receiver to a cliff, the range is increased to approximately 3.2 km (2 miles). Air-based tracking generally will have a range of 8 km (5 miles). Range also is dependent upon otter posture. When the abdomen is underwater, either when diving or posturally held underwater, the transmitter signal is attenuated, decreasing both range and signal strength. The primary advantages to this more invasive transmitter application are increased life span of the transmitter, and increased transmitter range. Obviously, the need for enhancements in these areas must be weighed against the invasive nature of the procedure. The technique for surgical implantation of the transmitter within the abdominal cavity is straightforward, except that the patient must be returned to the water following recovery from the anesthetic event. For that reason, surgical technique, particularly as it applies to tissue handling and suturing, is critically important. The technique currently used is similar to the original description (Williams and Siniff, 1983), but has been updated by frequent users (Murray, pers. obs.). The surgery tends to be well tolerated by most animals, with foraging
FIGURE 13 Radiographic appearance of an abdominal transmitter in an adult, male sea otter. Note that the transmitter has rotated 180°. Such positioning may attenuate signal strength. (Photo credit: Michael Murray.)
and diving typically observed within hours of recovery. The most significant postoperative concerns are the formation of surgical site seromas or dehiscence of the incision. Guynn et al. (1987) found that transmitters implanted into beavers (Castor canadensis) became coated with fibrinous connective tissue within 6 weeks of surgery, but there is no report of such coating occurring on transmitters implanted in sea otters. Several sea otter carcasses containing transmitters have been examined (Murray, pers. obs.). The celiotomy incisions appeared to heal well, although there was some localized cutaneous and subcutaneous inflammation and occasionally dehiscence, probably a result of grooming. The linea alba had not been affected in otters that were examined, but there was some degree of adhesion of omentum to the incision site. The transmitters appeared to be relatively innocuous, and the omentum had not adhered to them. The exterior coating of the transmitter package was consistently unaltered and the serosal response to the presence of this equipment was minor to nonexistent. Some transmitters did not necessarily stay where they were implanted, as some of them turned in a variety of directions and were located in any of all four abdominal quadrants. Ralls et al.
0839_frame_C38.fm Page 873 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
873
FIGURE 14 Intra-abdominal transmitter. The wider end of the transmitter contains the batteries, the transmitter is located in the midsection, and the antenna is coiled in the narrow end. (Photo credit: Michael Murray.)
(1989) also found that few, if any, of the free-floating transmitters implanted in sea otters became immobilized by adhesions. Similar results were found in yellow-bellied marmots (Marmota flaviventris), but three of ten transmitters implanted into beavers adhered to intestines, causing the death of one animal from intestinal obstruction (Guynn et al., 1987; Van Vuren, 1989). Ralls et al. (1989) recovered a transmitter from a sea otter killed accidentally 1195 days after implantation, with no pathology observed on necropsy. The effects of intraperitoneal implants on otters are thus believed to be minor or absent, but confirmatory data are lacking. However, a male sea otter on exhibit at the Monterey Bay Aquarium, Monterey, California is 15 years old and has had a transmitter since 5 months of age. Furthermore, numerous transmitters have been implanted in female otters that have subsequently given birth and reared pups without incident. Ralls and Siniff (1989) implanted transmitters in several pregnant sea otters, one of which died, and concluded that reproduction in implanted sea otters was normal; however, they conducted no detailed studies. Despite that, intentional implantation in pregnant animals is probably inappropriate. There also is some concern that implantation in very young animals weighing less than 8 to 9 kg (17 to 20 lb) may be problematic. Subcutaneous transmitters have not received much attention in sea otters. Garshelis and Siniff (1983) and Williams and Siniff (1983) implanted transmitters subcutaneously into ten sea otters. The sea otters manipulated the transmitters through the skin, but only one pup removed the sutures from the incision. The death of a subcutaneously implanted female from previously existing pneumonia, and her dependent pup (the one that had removed its sutures), discouraged further investigation of the subcutaneous space for transmitter implantation.
0839_frame_C38.fm Page 874 Tuesday, May 22, 2001 2:01 PM
874
CRC Handbook of Marine Mammal Medicine
Polar Bears Some tracking studies of polar bears (Ursus maritimus) have been conducted. Several different designs have been created to mount transmitting devices to the ears of polar bears, but problems associated with irritation of the ear, antenna dimensions, and casting and size of the transmitters prevented the widespread use of conventional telemetry ear-tag transmitters (Hansen, 1995). Furthermore, transmitters attached to the fur of polar bears using epoxy have only provided short-term data (Andriashek, pers. comm.). Radio and satellite transmitters, therefore, have been incorporated into neck collars for polar bears (for a description of some expandable/breakaway collars, including collar materials, release features and devices, hardware, accessories, and performance issues, see Web pages for ATS, Lotek, and Telonics; see Table 1). Because polar bears occupy one of the most remote habitats in the world, satellite telemetry is predominantly used to discern their movements and activities (Harris et al., 1990). The use of satellite transmitters incorporated into neck collars has been limited to females, however, because male polar bears have necks that are larger in diameter than their heads, which enables them to shed a collar quickly. Implantation of satellite transmitters with a percutaneous antenna into the subcutaneous space of the dorsal cervical region appears to be a promising approach for tracking male polar bears (for a detailed description of the surgical procedures, see Mulcahy and Garner, 1999). As technology advances, it is possible that the subcutaneous methods used for polar bears may be useful for attaching other types of transmitters and instruments to other marine mammal species (Mulcahy and Garner, 1999). Techniques, subclinical effects on individuals, and equipment should be evaluated under controlled conditions for other species of marine mammals.
Conclusion The methodologies used for any tracking study will depend upon the objective of the study, the type of data to be collected, funding, field conditions, equipment limitations, and the species being studied (White and Garrot, 1990). When choosing a tag and means of attachment, factors such as mass and expected behavior or activity of the species, tag size, weight, operational life, power consumption, and power output must be considered. Animal welfare, tag effects on the animal, experimental design, and data analysis and interpretation must all be taken into account when planning a tracking study (Heezen and Tester, 1967).
Acknowledgments The authors thank James Harvey, Tom Loughlin, and Michael Scott for reviewing this chapter, and Jack Ames, Michelle Staedler, and Pam Tuomi for their assistance in preparing the manuscript.
References Ames, J.A., Hardy, R.A., and Wendell, F.E., 1983, Tagging materials and methods for sea otters, Enhydra lutris, Calif. Fish Game, 69: 243–252. Andrews, R.D., 1998a, Instrumentation for the remote monitoring of physiological and behavioral variables, J. Appl. Physiol., 85: 1974–1981. Andrews, R.D., 1998b, Remotely releasable instruments for monitoring the foraging behaviour of pinnipeds, Mar. Ecol. Prog. Ser., 175: 289–294.
0839_frame_C38.fm Page 875 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
875
Argos, 1996, User’s Manual, Satellite Based Data Collection and Location System, Service Argos, Inc., Landover, MD, 176. Baker, J.D., and Johanos, T.C., 1999, Effects of research handling and instrumentation on Hawaiian monk seals, Monachus schauinslandi, in Abstracts of the 13th Biennial Conference on the Biology of Marine Mammals, Wailea, HI, 28 November–3 December, 226. Baranov, E.A., 1996, A device for data retrieval and recapture of diving animals in open water, Mar. Mammal Sci., 12: 465–468. Beaty, D., 1989, Technical notes on receivers, Telonics Q., 2(3). Bengtson, J.L., 1981, Ecology of Manatees (Trichechus manatus) in the St. Johns River, Florida, Ph.D. dissertation, University of Minnesota, St. Paul, 126. Bengtson, J.L., 1993, Telemetry and electronic technology, in Antarctic Seals, Research Methods and Techniques, Laws, R.M. (Ed.), Cambridge University Press, Cambridge, U.K., 119–139. Bonde, R.K., Lefebvre, L.W., Ward, L.I., Wright, I.E., and Valade, J.A., 1999, Post-release monitoring: An important aspect of the captive manatee release program, in Abstracts of the 13th Biennial Conference on the Biology of Marine Mammals, Wailea, HI, 28 November–3 December, 20. Boness, D.J., Bowen, W.D., and Oftedal, O.T., 1994, Evidence of a maternal foraging cycle resembling that of otariid seals in a small phocid, the harbor seal, Behav. Ecol. Sociobiol., 34: 95–104. Born, E.W., and Knutsen, L.O., 1997, Haul-out and diving activity of male Atlantic walruses (Odobenus rosmarus rosmarus) in NE Greenland, J. Zool. London, 243: 381–396. Boveng, P.L., Walker, B.G., and Bengtson, J.L., 1996, Variability in Antarctic fur seal dive data: Implications for TDR studies, Mar. Mammal Sci., 12: 543–554. Boyd, I.L., 1993, Selecting sampling frequency for measuring diving behavior, Mar. Mammal Sci., 9: 424–430. Boyd, I.L., and Arnborn, T., 1991, Diving behaviour in relation to water temperature in the southern elephant seal: Foraging implications, Polar Biol., 11: 259–266. Boyd, I.L., and Croxall, J.P., 1996, Dive durations in pinnipeds and seabirds, Can. J. Zool., 74: 1696–1705. Burger, B., 1991, Duty cycling of satellite transmitters, Telonics Q., 4(1). Costa, D.P., 1988, Methods for studying the energetics of freely diving animals, Can. J. Zool., 66: 45–52. Croll, D.A., Tershy, B.R., Hewitt, R., Demer, D., Hayes, S., Fielder, P., Popp, J., and Lopez, V.L., 1998, An integrated approach to the foraging ecology of marine birds and mammals, Deep-Sea Res., 11, 45: 1353–1371. Davis, R.W., Fuiman, L.A., Williams, T.M., Collier, S.O., Hagey, W.P., Kanatous, S.B., Kohin, S., and Horning, M., 1999, Hunting behavior of a marine mammal beneath the Antarctic fast ice, Science, 283: 993–996. DeLong, R.L., and Stewart, B.S., 1991, Diving patterns of northern elephant seal bulls, Mar. Mammal Sci., 7: 369–384. Deutsch, C.J., Bonde, R.K., and Reid, J.P., 1998, Radio-tracking manatees from land and space: Tag design, implementation, and lessons learned from long-term study, Mar. Tech. Soc. J., 32: 18–29. Eagle, T.C., Choromanski-Norris, J., and Kuechle, V.B., 1984, Implanting radio transmitters in mink and Franklin’s ground squirrels, Wildl. Soc. Bull., 12: 180–184. Early, G., Cooper, R., Kraus, S., and Williamson, M., 1999, The movements and behavior or released rehabilitated seals, in Abstracts of the 13th Biennial Conference on the Biology of Marine Mammals, Wailea, HI, 28 November–3 December, 52. Eguchi, T., 1998, Morphology of the Pacific Harbor Seal (Phoca vitulina richardsi) Using Elkhorn Slough, California, and Their Movements and Diving Behavior in the Monterey Bay Area, M.Sc. thesis, Moss Landing Marine Laboratories, California State University, Fresno, 105. Ellis, G.M., and Trites, A.W., 1992, The RAM-packs came back: A method for attaching and recovering pinniped dive recorders, Aquat. Mammals, 18: 61–64. Evans, W.E., 1971, Orientation behavior of delphinids: Radio telemetric studies, Ann. N.Y. Acad. Sci., 188: 142–160.
0839_frame_C38.fm Page 876 Tuesday, May 22, 2001 2:01 PM
876
CRC Handbook of Marine Mammal Medicine
Fancy, S.G., Pank, L.F., Douglas, D.C., Curby, C.H., Garner, G.W., Amstrup, S.C., and Regelin, W.L., 1988, Satellite telemetry: A new tool for wildlife research and management, U.S. Fisheries Wildlife Resources Publ. 172. Fedak, M.A., Pullen, M.R., and Kanwisher, J., 1983, Attachment of a radio tag to the fur of seals, J. Zool. London, 200: 298–300. Feldkamp, S.D., DeLong, R.L., and Antonelis, G.A., 1989, Diving patterns of California sea lions, Zalophus californianus, Can. J. Zool., 67: 872–883. Garshelis, D.L., and Siniff, D.B., 1983, Evaluation of radio-transmitter attachment for sea otters, Wildl. Soc. Bull., 11: 378–383. Gaskin, D.E., Smith, G.J.D., and Watson, A.P., 1975, Preliminary study of movements of harbour porpoises (Phocoena phocoena) in the Bay of Fundy using radio telemetry, Can. J. Zool., 53: 1466–1471. Gavette, C.A., 1992, The Dispersal, Distribution, and Disposition of Released Rehabilitated Pinnipeds, M.A. thesis, San Francisco State University, San Francisco, CA, 118. Gentry, R.L., and Kooyman, G.L., 1986, Methods of dive analysis, in Fur Seals: Maternal Strategies on Land and at Sea, Gentry, R.L., and Kooyman, G.L. (Eds.), Princeton University Press, Princeton, NJ, 28–40. Goodyear, J.D., 1993, A sonic/radio tag for monitoring dive depths and underwater movements, J. Wildl. Manage., 57: 503–513. Goulet, A.-M., Hammill, M.O., and Barrette, C., 1999, Quality of satellite telemetry locations of gray seals (Halichoerus grypus), Mar. Mammal Sci., 15: 589–594. Guinn, M., and Lee, J., 1996, Testing the possibilities, Telonics Q., 9(2). Guynn, D.C.J., Davis, J.R., and Von Recum, A.F., 1987, Pathological potential of intraperitoneal transmitter implants in beavers, J. Wildl. Manage., 51: 605–606. Hall, A., Moss, S., and McConnell, B., 2000, A new tag for identifying seals, Mar. Mammal Sci., 16: 254–257. Hammill, M.O., Lesage, V., Lobb, G., and Carter, P., 1999, A remote release mechanism to recover timedepth recorders from marine mammals, Mar. Mammal Sci., 15: 584–588. Hansen, B., 1995, The latest word in Ursine accessories, Telonics Q., 8(2). Hanson, M.B., Westgate, A.J., and Read, A.J., 1998, Evaluation of small cetacean tags by measuring drag in wind tunnels, Marine Mammal Protection Act and Endangered Species Act implementation program 1997, Hill, P.S., and DeMaster, D.P. (Eds.), AFSC Processed Rep. 98-10: 51–62. Harris, R.B., Fancy, S.G., Douglas, D.C., Garner, G.W., Amstrup, S.C., McCabe, T.R., and Pank, L.F., 1990, Tracking wildlife by satellite: Current systems and performance, U.S. Department of the Interior Fish and Wildlife Service, Washington, D.C., Fish and Wildlife Technical Report 30: 52. Harvey, J.T., 1987, Population Dynamics, Annual Food Consumption, Movements, and Dive Behaviors of Harbor Seals, Phoca vitulina, in Oregon, Ph.D. dissertation, Oregon State University, Corvalis, 177. Harvey, J.T., 1991, Survival and behavior of previously captive harbor seals after release into the wild, in Marine Mammal Strandings in the United States, Proceedings of the Second Marine Mammal Stranding Workshop, Reynolds, J.E., and Odell, D.K. (Eds.), NOAA Technical Report NMFS 98: 117–122. Harvey, J.T., and Mate, B.R., 1984, Dive characteristics and movements of radio-tagged gray whales in San Ignacio Lagoon, Baja California Sur, Mexico, in The Gray Whale (Eschrichtius robustus), Jones, M.L., Swartz, S.L., and Leatherwood, S. (Eds.), Academic Press, Orlando, FL, 561–575. Harvey, J.T., Brown, R.B., and Mate, B.R., 1983, Two sightings following release of rehabilitated harbor seals, Murrelet, 64: 18. Heezen, K.L., and Tester, J.R., 1967, Radio-tracking by triangulation, J. Wildl. Manage., 31: 124–141. Hill, R., 1997, MK5/MK6 Time-Depth Recorder Instruction Manual, Wildlife Computers, Redmond, WA, 68. Hill, R., 2000, Satellite-Linked Time-Depth Recorder Instruction Manual, Wildlife Computers, Redmond, WA, 57.
0839_frame_C38.fm Page 877 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
877
Hooker, S.K., and Baird, R.W., 1999, Deep-diving behaviour of the northern bottlenose whale, Hyperoodon ampullatus (Cetacea: Ziphiidae), Proc. R. Soc. London Biol. Sci., 266: 671–676. Hooker, S.K., and Baird, R.W., 2001, Diving and ranging behavior of odontocetes: A methodological review and critique, Mammal Rev., 31: 81–105. Hoover, J.P., 1984, Surgical implantation of radio telemetry devices in American river otters, J. Am. Vet. Med. Assoc., 185: 1317–1320. Irvine, A.B., and Scott, M.D., 1984, Development and use of marking techniques to study manatees in Florida, Fla. Sci., 47: 12–26. Irvine, A.B., Wells, R.S., and Scott, M.D., 1982, An evaluation of techniques for tagging small odontocete cetaceans, Fish. Bull., 80: 135–143. Jeffries, S.J., Brown, R.F., and Harvey, J.T., 1993, Techniques for capturing, handling and marking harbour seals, Aquat. Mammals, 19: 21–25. Johnson, S.A., and Berkley, K.A., 1999, Restoring river otters in Indiana, Wildl. Soc. Bull., 27: 419–428. Keating, K.A., 1994, An alternative index of satellite telemetry location error, J. Wildl. Manage., 58: 414–421. Kenward, R., 1987, Wildlife Radio Tagging: Equipment, Field Techniques and Data Analysis, Academic Press, San Diego, CA, 222. Kooyman, G.L., 1965, Techniques used in measuring diving capacities of Weddell seals, Polar Rec., 12: 391–394. Lander, M.E., 1998, Success of Free-Ranging and Rehabilitated Harbor Seal (Phoca vitulina richardsi) Pups in the Wild, M.S. thesis, Moss Landing Marine Laboratories, San Francisco State University, San Francisco, CA, 108. Le Boeuf, B.J., Morris, P.A., Blackwell, S.B., Crocker, D.E., and Costa, D.P., 1996, Diving behavior of juvenile northern elephant seals, Can. J. Zool., 74: 1632–1644. Le Boeuf, B.J., Crocker, D.E., Costa, D.P., Blackwell, S.B., Webb, P.M., and Houser, D.S., 2000, Foraging ecology of northern elephant seals, Ecol. Monogr., 70: 353–382. Loomis, M.R., 1993, Identification of animals in zoos, in Zoo and Wildlife Medicine: Current Therapy 3, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 21–23. Loughlin, T.R., 1977, Activity Patterns, Habitat Partitioning, and Grooming Behavior of the Sea Otter, Enhydra lutris, in California, Ph.D. dissertation, University of California, Los Angeles, 110. Loughlin, T.R., Bengtson, J.L., and Merrick, R.L., 1987, Characteristics of feeding trips of female northern fur seals, Can. J. Zool., 65: 2079–2084. Lowry, L.F., Frost, K.J., Davis, R., DeMaster, D.P., and Suydam, R.S., 1998, Movements and behavior of satellite-tagged spotted seals (Phoca largha) in the Bering and Chukchi Seas, Polar Biol., 19: 221–230. Martin, A.R., and Smith, T.G., 1992, Deep diving in wild, free-ranging beluga whales, Delphinapterus leucas, Can. J. Fish. Aquat. Sci., 49: 462–466. Mate, B.R., 1989, Watching habits and habitats from Earth satellites, Oceanus, 32: 14–18. Mate, B.R., and Harvey, J.T., 1984, Ocean movements of radio-tagged gray whales, in The Gray Whale (Eschrichtius robustus), Jones, M.L., Swartz, S.L., and Leatherwood, S. (Eds.), Academic Press, Orlando, FL, 577–589. Mate, B.R., Harvey, J.T., Hobbs, L., and Maiefski, R., 1983, A new attachment device for radio-tagging large whales, J. Wildl. Manage., 47: 869–872. Mate, B.R., Nieukirk, S., Mesecar, R., and Martin, T., 1992, Application of remote sensing methods for tracking large cetaceans: North Atlantic right whales (Eubalaena glacialis), Final Report for U.S. Department of the Interior, Minerals Management Service, Washington, D.C., 167. Mate, B.R., Rossbach, K.A., Nieukirk, S.L., Wells, R.S., Irvine, A.B., Scott, M.D., and Read, A.J., 1995, Satellite-monitored movements and dive behavior of a bottlenose dolphin (Tursiops truncatus) in Tampa Bay, Florida, Mar. Mammal Sci., 11: 452–463. Mate, B.R., Nieukirk, S.L., and Kraus, S.D., 1997, Satellite monitored movements of the north right whale, J. Wildl. Manage., 61: 1393–1405.
0839_frame_C38.fm Page 878 Tuesday, May 22, 2001 2:01 PM
878
CRC Handbook of Marine Mammal Medicine
Mate, B.R., Lagerquist, B.A., and Calambokidis, J., 1999, Movements of north Pacific blue whales during the feeding season off southern California and their southern fall migration, Mar. Mammal Sci., 15: 1246–1257. McCafferty, D.J., Boyd, I.L., Walker, T.R., and Taylor, R.I., 1998, Foraging responses of Antarctic fur seals to changes in the marine environment, Mar. Ecol. Prog. Ser., 166: 285–299. McConnell, B.J., Chambers, C., and Fedak, M.A., 1992, Foraging ecology of southern elephant seals in relation to the bathymetry and productivity of the Southern Ocean, Antarc. Sci., 4: 393–398. Merrick, R.L., and Loughlin, T.R., 1997, Foraging behavior of adult female and young-of-the-year Steller sea lions in Alaskan waters, Can. J. Zool., 75: 776–786. Merrick, R.L., Loughlin, T.R., Antonelis, G.A., and Hill, R., 1994, Use of satellite-linked telemetry to study Steller sea lion and northern fur seal foraging, Polar Res., 13: 105–114. Merrick, R.L., Loughlin, T.R., and Calkins, D.G., 1996, Hot branding: A technique for long-term marking of pinnipeds, NOAA Technical Memorandum, NMFS-AFSC-68, 21. Mulcahy, D.M., and Garner, G., 1999, Subcutaneous implantation of satellite transmitters with percutaneous antennae into male polar bears (Ursus maritimus), J. Zoo Wildl. Med., 30: 510–515. Pabst, D.A., Rommel, S.A., and McLellan, W.A., 1999, Functional morphology of marine mammals, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 15–72. Parrish, F.A., Craig, M.P., Ragen, T.J., Marshall, G.J., and Buhleier, B.M., 2000, Identifying diurnal foraging habitat of endangered Hawaiian monk seals using a seal-mounted video camera, Mar. Mammal Sci., 16: 392–412. Pessier, A.P., Stalis, I.H., Sutherland-Smith, M., Spelman, L.H., and Montali, R.J., 1999, Soft tissue sarcomas associated with identification microchip implants in two small zoo mammals, in Proceedings, American Association of Zoo Veterinarians, Columbus, OH, 9–14 October, 139–140. Pitcher, K.W., and McAllister, D.C., 1981, Movements and haulout behavior of radio-tagged harbor seals, Phoca vitulina, Can. Field Nat., 95: 292–297. Ralls, K., Siniff, D.B., Williams, T.D., and Kuechle, V.B., 1989, An intraperitoneal radio transmitter for sea otters, Mar. Mammal Sci., 5: 376–381. Rathbun, G.B., Reid, J.P., and Bourassa, J.B., 1987, Design and construction of a tethered, floating radiotag assembly for manatees, National Technical Information Service, PB87-161345/AS, Springfield, VA, 49. Rathbun, G.B., Reid, J.P., and Carowan, G., 1990, Distribution and movement patterns of manatees (Trichechus manatus) in northwestern peninsular Florida, Florida Marine Research Publication, 48: 1–33. Read, A.J., and Gaskin, D.E., 1985, Radio-tracking the movements and behavior of harbor porpoises Phocoena phocoena (L.), in the Bay of Fundy Canada, Fish. Bull., 83: 543–552. Read, A.J., and Westgate, A.J., 1997, Monitoring the movements of harbour porpoises (Phocoena phocoena) with satellite telemetry, Mar. Biol., 130: 315–322. Reid, D.G., Melquist, W.E., Woolington, J.D., and Noll, J.M., 1986, Reproductive effects of intraperitoneal transmitter implants in river otters, J. Wildl. Manage., 50: 92–94. Reid, J.P., and O’Shea, T.J., 1989, Three years operational use of satellite telemetry on Florida manatees: Tag improvements based on challenges from the field, in Proceedings of the 1989 North American Argos Users Conference and Exhibit, San Diego, CA, 15–17 May, 361. Reid, J.P., Bonde, R.K., and O’Shea, T.J., 1995, Reproduction and mortality of radio-tagged and recognizable manatees on the Atlantic Coast of Florida, in Population Biology of the Florida Manatee (Trichechus manatus latirostris), O’Shea, T.J., Ackerman, B.B., and Percival, H.F. (Eds.), National Biological Service Information and Technical Report 1: 171–191. Ribic, C.A., 1982, Autumn movement and home range of sea otters in California, J. Wildl. Manage., 46: 795–801. Richardson, W.J., 1995, Marine mammal hearing, in Marine Mammals and Noise, Richardson, W.J., Greene, C.R., Malme, C.I., and Thomson, D.H. (Eds.), Academic Press, San Diego, CA, 205–240.
0839_frame_C38.fm Page 879 Tuesday, May 22, 2001 2:01 PM
Tagging and Tracking
879
Schneider, K., Baird, R.W., Dawson, S., Visser, I., and Childerhouse, S., 1998, Reactions of bottlenose dolphins to tagging attempts using remotely-deployed suction-cup tag, Mar. Mammal Sci., 14: 316–324. Schreer, J.F., and Testa, J.W., 1996, Classification of Weddell seal diving behavior, Mar. Mammal Sci., 12: 227–250. Schwantje, H.T.M., Weir, R., and McAdie, M., 1998, Capture and immobilization of mustelids in British Columbia, in Joint Conference Proceedings of the American Association of Zoo Veterinarians and American Association of Wildlife Veterinarians, 450. Scott, M.D., Wells, R.S., Irvine, A.B., and Mate, B.R., 1990, Tagging and marking studies on small cetaceans, in The Bottlenose Dolphin, Leatherwood, S., and Reeves, R.R. (Eds.), Academic Press, San Diego, CA, 489–514. Seagars, D.J., 1988, The fate of released rehabilitated pinnipeds based on tag-resight information: A preliminary assessment, NMFS, SWR Admin. Rep. SWR-88-1, 31. Skrovan, R.C., Williams, T.M., Berry, P.S., Moore, P.W., and Davis, R.W., 1999, The diving physiology of bottlenose dolphins (Tursiops truncatus) II. Biomechanics and changes in buoyancy at depth, J. Exp. Biol., 202: 2749–2761. Spelman, L.H., Jochem, W.J., Sumner, P.W., Redmond, D.P., and Stoskopf, M.K., 1997, Postanesthetic monitoring of core body temperature using telemetry in North American river otters (Lutra canadensis), J. Zoo Wildl. Med., 28: 413–417. Stewart, B.S., and DeLong, R.L., 1995, Double migrations of the northern elephant seal Mirounga angustirostris, J. Mammal., 76: 196–205. Stewart, B.S., Leatherwood, S., Yochem, P.K., and Heide-Jorgensen, M.D., 1989, Harbor seal tracking and telemetry by satellite, Mar. Mammal Sci., 5: 361–375. Stoddart, D.M., 1978, The growth and behaviour of a hand-reared common seal (Phoca vitulina vitulina) after release to the sea, J. Zool. London, 186: 535–574. Stone, G.S., Tausig, H.C., and Schubel, J.R., (Eds.), 1998, Marine animal telemetry tags, New England Aquarium Aquatic Forum Series Rep. 98-3, 14–15 May, Boston, MA, 62. Stoskopf, M.K., Spelman, L.H., Sumner, P.W., Redmond, D.P., Jochem, W.J., and Levine, J.F., 1997, The impact of water temperature on core body temperature of North American river otters (Lutra canadensis) during simulated oil spill recovery washing protocols, J. Zoo Wildl. Med., 28: 407–412. Testa, J.W., and Rothery, P., 1992, Effectiveness of various cattle ear tags as markers for Weddell seals, Mar. Mammal Sci., 8: 344–353. Thomas, J.A., Cornell, L.H., Joseph, B.E., Williams, T.D., and Dreischman, S., 1987, An implanted transponder chip used as a tag for sea otters (Enhydra lutris), Mar. Mammal Sci., 3: 271–274. Tomkiewicz, S., 1988, Why it works: Almost everything you ever wanted to know about antennas, Telonics Q., 1(1). Tomkiewicz, S., 1996, GPS applications for wildlife—a review, Telonics Q., 9(1). Tully, D., 1999, Analysis of Foraging Behaviour of Adult Male Harbour Seals Using Animal-Borne Video Data: Effects of Prey Type on Tactics and Profitability, M.Sc. thesis, Dalhousie University, Halifax, Nova Scotia, Canada. Van Vuren, D., 1989, Effects of intraperitoneal transmitter implants on yellow-bellied marmots, J. Wildl. Manage., 53: 320–323. Walker, B.G., and Boveng, P.L., 1995, Effects of time-depth recorders on maternal foraging and attendance behavior of Antarctic fur seals (Arctocephalus gazella), Can. J. Zool., 73: 1538–1544. Watkins, W.A., Daher, M.A., Fristrup, K.M., Howland, T.J., and Di Sciara, G.N., 1993, Sperm whales tagged with transponders and tracked underwater by sonar, Mar. Mammal Sci., 9: 55–67. Webber, M.A., and Allen, S.G., 1986, Resightings of two rehabilitated and released harbor seals in California, Calif. Fish Game, 45: 60–61. Wells, R.S., Rhinehart, H.L., Cunningham, P., Whaley, J., Baran, M., Koberna, C., and Costa, D.P., 1999, Long distance offshore movements of bottlenose dolphins, Mar. Mammal Sci., 15: 1098–1114.
0839_frame_C38.fm Page 880 Tuesday, May 22, 2001 2:01 PM
880
CRC Handbook of Marine Mammal Medicine
Westgate, A.J., Read, A.J., Berggren, P., Koopman, H.N., and Gaskin, D.E., 1995, Diving behaviour of harbour porpoises, Phocoena phocoena, Can. J. Fish. Aquat. Sci., 52: 1064–1073. White, G.C., and Garrott, R.A., 1990, Analysis of Wildlife Radio-Tracking Data, Academic Press, San Diego, CA, 383. Wiig, O., Gjertz, I., Griffiths, D., and Lydersen, C., 1993, Diving patterns of an Atlantic walrus (Odobenus rosmarus rosmarus) near Svalbard, Polar Biol., 13: 71–72. Williams, T.D., and Siniff, D.B., 1983, Surgical implantation of radio telemetry devices in the sea otter, J. Am. Vet. Med. Assoc., 183: 1290–1291. Williams, T.M., Davis, R.W., Fuiman, L.A., Francis, J., Le Boeuf, B.J., Horning, M., Calambokidis, J., and Croll, D.A., 2000, Sink or swim: Strategies for cost-efficient diving by marine mammals, Science, 288: 133–136. Wilson, R.P., Grant, W.S., and Duffy, D.C., 1986, Recording devices on free-ranging marine animals: Does measurement affect foraging performance? Ecology, 67: 1091–1093. Wright, I.E., Wright, S.D., and Sweat, J.M., 1998, Use of passive integrated transponder (PIT) tags to identify manatees (Trichechus manatus latirostris), Mar. Mammal Sci., 14: 641–645. York, A.E., 1989, Lack of controls in tagging experiments: How reliable are findings from mark–recapture studies? in Abstracts of the 8th Biennial Conference on the Biology of Marine Mammals, Pacific Grove, CA, 7–11 December.
0839_frame_C39.fm Page 881 Tuesday, May 22, 2001 2:02 PM
39 Marine Mammal Transport Jim Antrim and James F. McBain
Introduction During the last 30 years, specialized marine mammal transport techniques and equipment have been developed to cope with the unique physiology of marine mammals. Therefore, safe, successful transportation is possible with the proper attention to detail and the use of technology appropriate to the type of marine mammal being transported. In moving marine mammals, one must always be aware of, and comply with, local, state/ provincial, federal, and international laws and regulations or mandates regarding transportation. Since constant monitoring is necessary for the early detection and correction of respiratory, thermoregulatory, postural, and behavioral abnormalities, U.S. law mandates that a qualified attendant be present when transporting cetaceans, pinnipeds, sirenians, and sea otters (Department of Agriculture, 1999).
Regulations Two published regulatory documents deal with the transport of marine mammals and should be consulted as appropriate. The first of these is applicable only to individuals transporting marine mammals under the jurisdiction of the United States. This document is commonly known as “The Animal Welfare Act—Specifications for the Humane Handling, Care, Treatment, and Transportation of Marine Mammals” (Department of Agriculture, 1999) (see Chapter 33, Legislation). The portion of this document covering marine mammal transportation standards includes specifications for consignments to carriers and intermediate handlers, primary enclosures used to transport marine mammals, primary conveyances, food and water requirements, care in transit, terminal facilities, and methods of handling. Enforcement of this regulation is the responsibility of the U.S. Department of Agriculture (USDA), Animal and Plant Health Inspection Service (APHIS). The second document is “Live Animals Regulations,” published by the International Air Transport Association (IATA), and accepted by the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) (Convention Resolution 4.20) and the Office International des Epizooties (OIE) as guidelines for transporting animals by air (International Air Transport Association, 1999). The European Economic Community, as well as many other countries, governmental entities, and airlines, have adopted these regulations as laws, policies, or minimum standards for transporting live animals in containers, pens, and stalls.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
881
0839_frame_C39.fm Page 882 Tuesday, May 22, 2001 2:02 PM
882
CRC Handbook of Marine Mammal Medicine
History of Marine Mammal Transport The overland transport of cetaceans has been documented at least as far back as 1554, when Rondelet, governing physician of the University of Medicine at Montpellier, France, recorded the regular transportation of live dolphins 130 miles inland by fishermen. These animals were shipped for the purpose of human consumption and were kept alive during transit to arrive at the market unspoiled (Rondelet, 1554). In about 1861, P.T. Barnum reported the transport of six belugas or white whales (Delphinapterus leucas) from the St. Lawrence River to New York in a box on a bed of seaweed. Only one of these animals survived the transport and was exhibited for 2 years with a bottlenose dolphin (Tursiops truncatus) at Barnum’s Museum (Wyman, 1863). In 1877, a beluga, collected in Labrador, was shipped by sailing vessel to Montreal, then by train to New York, with a total of 14 days travel time. The animal was subsequently placed on a bed of seaweed for the 10-day steamship voyage to England, where it survived only 4 days after arrival. In the following year, the same route was used to transport four belugas. Three of these animals survived the trip (Lee, 1878). Another early documented cetacean transport occurred in the 1870s when two belugas were sent from Labrador to the Brighton Aquarium in England in the flooded hold of a ship. Unfortunately, both animals died during the transport (Slijper, 1979). More closely approximating current transport methods, the New York Aquarium transported a group of bottlenose dolphins within water-filled boxes in 1907. About 5 years later, Taylor first employed a soft stretcher for marine mammal transportation (Townsend, 1914). Early transport of killer whales (Orcinus orca) occurred much later and somewhat differently, due to the size of the animals involved. In June 1965, a floating pen pulled by a boat, measuring 18 × 12 × 5 m (59 × 40 × 16 ft), was used to transport an adult killer whale from Namu, British Columbia, to Seattle, Washington. The journey took 2 weeks and the animal traveled a distance of 675 km (422 miles) (Newman and McGreer, 1966). In December 1965, a female killer whale was transported by air in a box from Seattle, Washington, to SeaWorld, Inc. in San Diego, California. The animal was transported easily, and quickly acclimated to the zoological environment.
Cetaceans Cetaceans spend their entire lives in water, which provides uniform support by equal distribution of pressure over the entire body (Ridgway, 1972). The result is near-weightlessness, allowing normal respiration. Additionally, water rapidly dissipates cetacean metabolic heat at 25 times the rate of air (Ridgway, 1972). As a result, when removed from their free-swimming state, the primary criteria that must be met to transport cetaceans successfully are adequate body support for their comfort and normal respiration, and a temperature control mechanism to assist with thermoregulation (see Figures 1 and 2). Body support techniques have been developed to allow normal breathing during transport. Cetaceans are now moved in fabric stretchers and suspended in water-filled transport units, closely approximating the near-weightlessness of water. Also, the widespread use of aircraft for long-distance travel has shortened the amount of time cetaceans are removed from a free-swimming water environment (Cornell, 1978). Temperature control is of paramount importance. Since most of a cetacean’s body is enclosed within a layer of thick, insulating blubber, thermoregulation in the water is controlled through constriction or dilation of the peripheral vessels in the heavily vascularized pectoral flippers, tail flukes, and dorsal fin (Slijper, 1979). Consequently, cetaceans are less able to dissipate excess
0839_frame_C39.fm Page 883 Tuesday, May 22, 2001 2:02 PM
Marine Mammal Transport
883
FIGURE 1 Beluga (Delphinapterus leucas) transport unit, side cutaway view. (Image courtesy of SeaWorld.)
FIGURE 2 Bottlenose dolphin transport (Tursiops truncatus) unit, side cutaway view. (Image courtesy of SeaWorld.)
0839_frame_C39.fm Page 884 Tuesday, May 22, 2001 2:02 PM
884
CRC Handbook of Marine Mammal Medicine
heat when removed from the water, due to the lower thermal conductivity of air. Thermoregulatory assistance can be provided in several ways. Suspending the cetacean within a waterfilled container will allow water temperature adjustments according to the animal’s needs. In an aircraft or enclosed-truck transport, the interior temperature may also be adjusted. It is best to conduct overland cetacean transports in open trucks at night or on overcast days to prevent sunburn. If this is not possible, light, moistened towels may be placed over the cetacean’s skin to prevent sunburn and desiccation. In early years, much of the difficulty in transporting cetaceans was due to equipment failures. Most failures can be avoided through proper planning and thorough preparation. Therefore, based on the possibility of problems, there must always be contingency plans to ensure the welfare of the animals being transported. Such plans require adequate personnel and equipment, either to return cetaceans to holding facilities if the transport is aborted, or to care for the animals if a delay occurs at an unexpected location. Certainly, the transport should be canceled if there are no contingency plans prepared to manage significant delays. Animal health problems resulting from early, unsophisticated cetacean transports included muscular stiffness upon return to water, depression of appetite, anemia as a result of abrasions, pressure necrosis, and respiratory infections. Because of the improved transport techniques used today, which provide appropriate water temperatures and allow for lateral and vertical flexion, muscular stiffness can be minimized, and usually disappears within a few hours to a few days, or is avoided entirely. Now, properly transported cetaceans resume feeding immediately upon removal from their transport units, especially animals that have experienced transport previously. Abrasions and pressure necrosis are avoided through the use of fitted stretchers, nonabrasive materials, and proper positioning within the stretcher and transport unit. The utilization of modern rapid transport has decreased or eliminated the previously common use of antibiotics and corticosteroids for the prevention and treatment of transport-related respiratory infections. Proper support equipment, well-trained, experienced personnel, and strict attention to logistical details are important factors influencing the success of any cetacean transport. A thorough assessment of the health status of the cetacean must be made prior to transport. Such an assessment should include behavioral observations, a physical examination, hematological examination, and, if possible, urinalysis. Cetaceans are generally not fed for 24 hours prior to shipment to minimize the volume of body wastes discharged into the transport container. Fasting may also reduce the incidence of regurgitation during transit. The initial step in cetacean transport is lifting the whale or dolphin from its pool or holding facility in a custom-fitted stretcher made of soft material, generally nylon or canvas. These stretchers may be lined with wool or chamois, if desired, to minimize the possibility of abrasion. Stretchers must be constructed according to accurate measurements of the body of the individual cetacean to ensure proper fit and provide equal distribution of body weight along the stretcher. Paired openings in the stretcher allow the pectoral flippers to extend in an unrestricted, natural position (Ridgway, 1972; Cornell, 1978). The animal is positioned in the stretcher so that pressure is not increased on specific points along the body or flippers, which might lead to pressure necrosis or abrasion. During loading, every effort must be made to avoid abrading the highly vascularized skin of the cetacean. The danger of pressure necrosis is greatest in the axillary region (region just posterior to the insertion of the pectoral flippers). Metal poles supporting the stretcher must be sturdy enough to support the weight of the cetacean, even if the weight suddenly shifts. The pole ends should be rounded or covered with caps to eliminate sharp edges. Rope, nylon webbing, chain, or vinyl-covered steel cable may be used to connect the stretcher poles to the lifting crane. Eyebolts on the stretcher poles where the lifting tackle is attached should be padded to prevent lacerations and abrasions to both cetaceans and personnel. When engaged through the eyebolts, lifting hooks must face away from the animal. Body surfaces likely
0839_frame_C39.fm Page 885 Tuesday, May 22, 2001 2:02 PM
Marine Mammal Transport
885
to be exposed to the drying effects of air may be covered with a protective ointment or kept wet by spraying or dousing with water to prevent desiccation (Cornell, 1978). A crane is required to lift most cetaceans safely because of their weight and the possibility that they may move while being lifted. Therefore, the crane’s lifting capacity must be equal to the animal’s weight plus an acceptable safety factor. At the location of the cetacean lift, the supporting surface should be inspected and all details of the lift must be discussed with the crane operator. Hydraulic cranes are preferred over mechanical cranes because they operate more smoothly. For safety, a cetacean must be kept as low to the ground as practical during all phases of the lift. Tags and/or guidelines must be attached to the stretcher, not to the eyebolts, where they might bind with lifting tackle. When lifting small cetaceans (<400 kg; <880 lb) two tags or guidelines should be used. These should be attached to the stretcher poles on opposite ends. For large animals (>400 kg), four lines should be used, two at each end attached directly to the stretcher poles. Guidelines should be attended during all phases of the lift to control unwanted motion. Once fitted into its custom stretcher, the cetacean is lowered into a watertight transport unit of appropriate size, allowing the animal clearance from the ends, sides, and bottom of the transport unit. Transporters currently in use at Sea World are constructed of fiberglass-covered foam, or polyvinyl chloride sheet stock (Divinycell®, DIAB, Inc., DeSoto, TX, and DIAB International, Laholm, Sweden), or marine plywood. Some of these transport units are reinforced and protected by a steel frame. A removable lining of vinyl is installed on the inside walls of the transport unit to prevent abrasions. Lids or baffles are secured over both ends of the unit to reduce the possibility of water spillage. A stretcher support system may be incorporated into transport units to enable attendants to adjust the position of the animal easily by raising or lowering the stretcher poles during loading or transport (Figures 3 and 4). Once positioned within the transport unit, sufficient fresh water is added to submerge the lower 2/3 to 3/4 of the animal’s body. It is recommended that salt water not be used, because of potential damage to aircraft components if spillage occurs. Suspension in water provides cooling and buoyancy, permitting exertion-free respiration throughout transport. If transported in an adequately supported posture, there should be little or no detectable variation in respiratory rhythm during transport (Lilly, 1961; Cornell, 1978). The temperature of the water in the transport unit should be the same or close to the temperature to which the cetacean is habituated. If the cetacean is being moved from water of one temperature to a different temperature, it is best to have the water in the transport unit at a temperature between the two. If the ambient air temperature is significantly warmer than the temperature of the water in the transport unit, ice may be added to maintain the appropriate water temperature. For ground transport, the planned route of travel must be examined ahead of time with attention to road conditions, such as overhead clearances (i.e., trees, electrical wires, low bridges), as well as any special limitations on weight or size. If the load is excessively heavy or overly wide, special permits may be required and must be obtained in advance of transport. A whale transport unit is usually carried on the back of a flatbed or lowboy trailer pulled by a diesel tractor. It is best if the tractor is equipped in a manner that the exhaust is directed toward the road surface to minimize exposure of the animals and human attendants. A spare tractor should accompany the transport caravan as assurance against delays due to tractor malfunction. Attendants accompanying the cetacean transport unit must be briefed prior to transport concerning all aspects of safety. A special permit allowing attendants to ride on the back of trucks may be required in some areas. Escort vehicles may precede and follow the caravan, and communications between all vehicles in the convoy can be maintained using radios or cellular phones. In many areas, notifying the appropriate authorities (i.e., police, highway patrol) of the
0839_frame_C39.fm Page 886 Tuesday, May 22, 2001 2:02 PM
886
CRC Handbook of Marine Mammal Medicine
FIGURE 3 Beluga transport unit. (Image courtesy of SeaWorld.)
planned move and route will aid ground transport and may eliminate traffic delays. An on-theground logistical team formed prior to transport can help coordinate communications among involved staff (i.e., husbandry, administration, media interest) at the time of transport itself. Most recent long-distance cetacean transports have involved the use of aircraft. Large cargo aircraft are preferred because of the size and weight of the cetaceans and their transport units. However, a major problem encountered over the years has been the relatively small size of many aircraft cargo doors. It is essential prior to transport to confirm that the transport unit and associated equipment will fit easily through the cargo door of the transport aircraft. Accurate weights on the animal transport units and other support equipment must also be obtained, so the aircraft can be balanced properly. Additionally, the aircraft needs to have provisions to secure the load safely to prevent shifting during flight. If, in transporting marine mammals, one uses a local airport regularly, it may be of help to brief airport and air carrier service personnel periodically, either in person or by written correspondence. Depending upon the size, weight, and configuration of the transport unit, various types of loading and unloading equipment may be used, including platform loaders, forklift trucks, and cranes. Platform loaders and forklift trucks decrease loading time, but are not always available with adequate lifting capacity and often must be scheduled in advance. Mechanical problems or inclement weather might necessitate landing at an alternative airport rather than the originally scheduled destination. For these reasons, large cetacean transport units should be designed and constructed for lifting by a crane, since one is usually available on short notice. However, crane companies frequently will not have the proper equipment to lift large cetacean transport units safely, making it essential that tackle such as steel cables, spreader bars, and
0839_frame_C39.fm Page 887 Tuesday, May 22, 2001 2:02 PM
Marine Mammal Transport
887
FIGURE 4 Bottlenose dolphin transport unit. (Image courtesy of SeaWorld.)
shackles accompany transport units at all times. Animal care ground personnel should remain at the airport until the transport aircraft takes flight, since occasional mechanical problems have required deplaning or have caused takeoff delays. Attendants accompanying the cetacean air transport may need to brief the flight crew before takeoff regarding the appropriate angle of takeoff and descent (both of which should be as gradual as possible to avoid spillage from the transport unit), as well as cabin pressure and air temperature. Since cetaceans appear to be susceptible to the effects of high-altitude sickness (acute mountain sickness), it is recommended that the altitude or pressure inside the aircraft be maintained to the equivalent of 1067 m altitude (3520 ft) or less. Keeping the cabin air temperature the same as the temperature of the water in the transport unit will avoid any unwanted temperature loss or gain. U.S. federal regulations specify most marine mammals be attended during transport, yet do not specify how many attendants are required. It is recommended there be at least one experienced attendant for each transported cetacean. This person should not only be experienced in animal husbandry, but also have emergency medical equipment on-site, and know how to use it, should it become necessary. The experienced attendant will also ensure close attention to each animal’s behavior, which can lead to early detection of abnormalities in respiration, posture, and activity level, and thereby provide attendants who can rapidly correct problems. Attendants should have ready access to personal protective equipment (i.e., rain gear, wet suit, waders) to aid in animal repositioning should it be necessary during transport. Attendants can also prevent desiccation of the cetacean’s skin and the subsequent development of “hot spots,” which later blister and slough, by regularly wetting exposed skin with water. Appropriate equipment for unloading and ground transport is necessary at the destination airport. Such equipment must be in position at the airport prior to the arrival of the aircraft to avoid delays.
0839_frame_C39.fm Page 888 Tuesday, May 22, 2001 2:02 PM
888
CRC Handbook of Marine Mammal Medicine
Upon arrival at the final destination, the cetacean is removed from the transport unit while still suspended in its fabric stretcher, lowered into a pool of water, and released. Such pools need not be shallow or small, but plans and equipment should be available in case the transported animal needs assistance, immediately following release or in subsequent days. Animals must remain under close observation for at least the first 24 hours following transport. Respiratory rates must be monitored and food offered frequently. Follow-up physical examinations and hematological examinations are prudent within the week following transport, or sooner if problems arise.
Pinnipeds Pinniped transport is less complex than that of cetaceans because pinnipeds live a semiaquatic existence, and are able to tolerate long periods out of water if kept cool and/or moist. Cool air temperatures ranging from 5 to 13°C (41 to 55°F) are suggested if transported by air or enclosed truck, but not mandatory unless the pinnipeds are exhibiting evidence of discomfort or hyperthermia. If ground transport is utilized, provisions for cooling, such as ice and water sprays, are necessary during transport. Ice may be placed within the transport unit, providing a freshwater source and cooling for the animal. As an alternative, ice may be placed on top of transport containers to furnish a cooling drip during transport. As is the case with cetaceans, pinnipeds should be fasted for 24 hours prior to the move, and should not be fed during transport. Transport containers need to be well ventilated, strong enough to contain the animal(s) being moved, and possess watertight bottoms extending at least 25 cm (10 in.) up the cage sides to prevent spillage of wastes and water. Any wood or metal used for cage construction must be free of sharp edges, splinters, or burrs on the interior surface. The openings in a wire or net mesh must not exceed 5 cm (2 in.) to prevent possible trauma during investigation or attempts to escape (Ridgway, 1972). Pinnipeds may be transported singly, especially in the case of large or aggressive individuals, or as groups. It is best to avoid overcrowding, as it can lead to overheating. Cage dimensions must be large enough to allow the animal to turn around and exhibit normal posturing during transport. As with cetaceans, the presence of qualified attendants is required by U.S. law and is necessary to note and correct difficulties arising during transport. Depending on the size of the animal, net and gloves should be available if it is necessary to do something to the animal during transport.
Sea Otters Sea otters, unlike cetaceans and pinnipeds, have a high metabolic rate and need to be fed wellrefrigerated food items such as clams and shrimp before and during transport. Due to this high metabolic rate, sea otters are as temperature sensitive during transport as cetaceans, maintaining primary thermoregulation through flipper contact with water. Therefore, a layer of ice must be placed on an elevated, draining cage floor, to serve as a source of fresh water and cooling. Sea otter transport cages may be constructed of wood, fiberglass, or smooth metal with mesh side panels, allowing good ventilation and unobstructed observation. The use of nylon netting instead of metal mesh for the side panels has minimized dental damage. A perforated floor can be placed above the waste-containing watertight cage bottom allowing uneaten food, feces, and urine to fall through. Cage bottoms should be removable, allowing emptying of waste during transport (Fitz-Gibbon and Hewlett, 1984). Plastic sky kennels are not recommended, because they provide inadequate ventilation, and otters may break teeth on the steel mesh door or
0839_frame_C39.fm Page 889 Tuesday, May 22, 2001 2:02 PM
Marine Mammal Transport
889
side vents. Furthermore, sky kennels promote fur soiling by forcing the otter to lie in its own waste products. Cage size must be large enough for the otter to turn around and groom freely. The presence of an attendant is required by U.S. law and is as imperative for sea otters as it is for pinnipeds and cetaceans. Thermoregulatory status must be constantly monitored as hyperthermia is the greatest obstacle to successful sea otter transport. Lethargy, panting, and flippers warm to the touch are all signs of overheating in sea otters. If the otters become too warm, spraying with cool water is helpful. It is important to keep a sea otter’s fur as clean as possible during transport. Soiled fur can be gently rinsed with a freshwater sprayer.
Sirenians Transport of sirenians is less complex than that of cetaceans, pinnipeds, or sea otters, because they actually require and are tolerant of warmer temperatures. Transport appears to be best accomplished using a temperature-controlled (18 to 23°C; 64 to 73°F) truck or airplane. Current IATA regulations stipulate both manatees and dugongs be transported in the same manner and using the same containers as cetaceans (International Air Transport Association, 1999). Manatees are easily transported out of water on soft, closed cell foam in a watertight container constructed of fiber-glassed Divinycell® or marine plywood. These containers should be slightly larger in length and width than the manatee, to enable the animal to make normal postural adjustments. Initially, animals may move around considerably. However, they soon settle down and begin resting comfortably in a ventral, lateral, or dorsal position. During air transport, standard nylon aircraft tie-down straps are used over the open tops of the containers to avoid any vertical movement of the animals that may be caused by air turbulence. Freshwater spray can be applied throughout the transport process to aid in cooling. Manatees do not require feeding during transport because of their slow metabolic rates. Fasting prior to transport produces a negligible change in waste output. Any solid stool passed during transport must be removed in a sanitary manner.
Polar Bears Transport of polar bears requires metal caging sturdy enough to contain and support the bear safely with a watertight cage floor to catch waste. Polar bears are best transported during cool weather. Aircraft cabin or enclosed trailer temperatures should be maintained below 13°C (55°F) during flight or ground transport. If a segment of the route involves transport outside a refrigerated vehicle, ice must be provided either within or on top of the cage. This ice can also provide a freshwater drinking source in extended transports. Food is not necessary during transport. Upon arrival, it may be a few days before the animal becomes acclimated enough to begin feeding. Although tooth breakage is a possibility, it is unwise to tranquilize polar bears during transport, because thermoregulatory mechanisms may be compromised. To promote calmness, consider conducting the transport in darkness. Despite the lack of requirement by U.S. law, an attendant can assure the polar bear is maintained at a comfortable temperature and can protect bystanders from unwanted contact.
Additional Medical Considerations Animals that have never been transported before should be given the opportunity to become familiar with the transport container. This means allowing the animal to enter the container
0839_frame_C39.fm Page 890 Tuesday, May 22, 2001 2:02 PM
890
CRC Handbook of Marine Mammal Medicine
or a similar unit in a way that allows it to become accustomed to the container prior to transport. In the case of cetaceans, this means arranging to put the animal into a stretcher and loading it into the water-filled transport unit. Allowing the animal to become desensitized to the transport device before the actual event is well worth the investment in time and effort. This makes the process less stressful for the animal, and also allows the transport staff to become aware of the type of behaviors to be expected. Prior to transport, a decision must be made regarding sedation. If the animal is expected to travel without difficulty, pre-transport sedation is unnecessary. If there is uncertainty, one may administer an oral dose of diazepam (Valium®, Roche Pharmaceuticals, Exton, PA) 30 to 60 min before any transport-related activities are apparent to the animal (see Chapter 29, Anesthesia, and Chapter 31, Pharmaceuticals). In the case of cetaceans, this means before the pool water is lowered or before the animals become aware of any other unusual activity or atypical accumulation of people. In spite of the fasting recommendations, the oral administration of diazepam does not seem to produce any problems. In manatees, pre-transport administration of sedation would most likely be given by intramuscular injection. If during the transport, the animal’s activity is deemed to be potentially threatening to its health, appropriate sedation should be administered. Diazepam or midazolam (Versed®‚ Roche Pharmaceuticals, Exton, PA) alone, or in combination with meperidine/pethidine (Demerol®‚ Sanofi Winthrop Pharmaceuticals, Overland Park, KS), has been the most widely used for transport sedation. Dosages are discussed in Chapters 29 (Anesthesia) and 31 (Pharmaceuticals). During a transport, the availability of refrigerated food fish may be helpful for the administration of oral medication. Offering fish can also provide a useful diversion for an animal upset by transport activities. As mentioned previously, the use of vinyl liners in cetacean transport units will minimize the possibility of skin abrasions. This is very important because most cetaceans demonstrate prolonged bleeding times from external wounds when in fresh water. In the event that an abrasion is hemorrhaging, ferric subsulfate (14% iron) (Medical Chemical Corporation, Torrance, CA; or Kwik-Stop, Arc Laboratories, Atlanta, GA) applied topically is usually effective. Yunnan Paiyao® (Yunnan Paiyao Factory), a Chinese herbal medication, applied topically or given orally, also appears to be useful for decreasing bleeding time. Occasionally during air transport of cetaceans, dramatic elevation of respiratory rate will occur. The causes for this can range from life-threatening to insignificant. A dolphin may act as if it is experiencing mild colic. In most cases these signs are the result of flatus, which is relieved when the gas passes. Cetaceans experiencing apparent high-altitude sickness (acute mountain sickness) can exhibit similar symptoms. However, this condition does not go away untreated and is life-threatening. Maintaining control of cabin pressure appears the best method of dealing with this disease. Since previous attempts to treat altitude sickness in cetaceans have met with limited success, it is best to opt for prevention. A well-planned transport with appropriate equipment, executed in a professional manner, results in minimal requirement for medical intervention.
Conclusion Safe, successful transportation of marine mammals may be routinely achieved with constant attention to detail and proper planning, preparation, communication, and execution. For cetaceans, care must be taken to ensure correct support of the animal while in transit. In the case of all marine mammals, and especially with sea otters, attention must be given to the thermoregulatory needs to prevent overheating. Competent attendants must be constantly aware of each animal’s condition to ensure the animal is traveling comfortably. The logistics
0839_frame_C39.fm Page 891 Tuesday, May 22, 2001 2:02 PM
Marine Mammal Transport
891
of proper marine mammal transport must include, but not be limited to, adequate staffing, correct equipment, and appropriate scheduling and routing.
Acknowledgments The authors thank Steve Clark and Marion and Mick Heller for their editorial assistance and Laurie Gage and Jeff Boehm for their peer reviews of the chapter. The authors also want to thank Dee Dee Dilworth and Judy Swift for assistance in the preparation of this manuscript. This manuscript was assigned SeaWorld publication no. 200-04-C.
References Cornell, L.H., 1978, Capture, transportation, restraint and marking, in Zoo and Wild Animal Medicine, Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 764–770. Department of Agriculture, 1999, Specifications for the Humane Handling, Care, Treatment, and Transportation of Marine Mammals, Animal and Plant Health Inspection Service, 9 Code of Federal Regulations (9 CFR), Chapter 1 (January 1, 1999 Edition) Subpart E: 85–107. Fitz-Gibbon, J., and Hewlett, K.G., 1984, Transport cage developed at the Vancouver public aquarium for sea otters, Int. Zoo Yearb., 23: 223–224. International Air Transport Association, 1999, Live Animals Regulations, 26th ed., 1 October, Montreal, 368 pp. Lee, H., 1878, The White Whale, R.K. Burt & Co., London, 16 pp. Lilly, J., 1961, Man and Dolphin, Doubleday, New York, 312 pp. Newman, M.A., and McGeer, P.L., 1966, The capture and care of a killer whale, Orcinus orca, in British Columbia, Zoologica (N.Y.), 51: 59–70. Ridgway, S.H., 1972, Homeostasis in an aquatic environment, in Mammals of the Sea, Ridgway, S.H. (Ed.), Charles C Thomas, Springfield, IL, 590–747. Rondelet, G., 1554, Libri de Piscibus Marinis, Lugduni, 2 vols., in 1 vol. Slijper, E.J., 1979, Whales, Cornell University Press, Ithaca, NY, 511 pp. Townsend, C.H., 1914, The porpoise in captivity, Zoologica (N.Y.), 1: 289–299. Wyman, J., 1863, Boston Nat. Hist. Soc. J., 7: 603–612.
0839_frame_C39.fm Page 892 Tuesday, May 22, 2001 2:02 PM
0839_frame_C40 Page 893 Tuesday, May 22, 2001 2:06 PM
VIII Specific Medicine and Husbandry of Marine Mammals
0839_frame_C40 Page 894 Tuesday, May 22, 2001 2:06 PM
0839_frame_C40 Page 895 Tuesday, May 22, 2001 2:06 PM
40 Cetacean Medicine James F. McBain
Introduction It is the author’s intention in this chapter to provide enough nuggets of information and philosophy that veterinarians will feel more comfortable with the challenges and pitfalls presented by the practice of cetacean medicine. It is assumed that the reader is a competent veterinary practitioner, since the practice of cetacean medicine largely depends on the same knowledge required for other species. There are differences in working with cetaceans that may seem intimidating. As with other exotic or unusual animals, once the differences are accommodated, the veterinarian can feel confident in applying his or her full range of knowledge and experience to the problems of working with a new species.
Philosophy No one likes to think behavioral change in a dolphin is due to illness. If one thinks there is a problem, one should deal with it. If one knows there is a problem, it may be too late. The best day to begin the search for answers is today. If there is clinical evidence that a dolphin has an illness, no one wants to believe it is serious. If the condition could be serious, assume that it is serious. If it is not known what disease the cetacean has, it is pneumonia until proved otherwise. When a dolphin or whale is sick, it is best to call a meeting to decide what will be done (not if something is to be done). Do not be caught off-guard by traps that present themselves in the practice of cetacean medicine. Humans, including veterinarians, are characterized by optimism and avoidance of the unknown. People always want things to turn out well, because that makes them happy, and they like being happy. A veterinarian closing the abdomen on an overweight canine ovariohysterectomy is not hoping that the small amount of uncontrolled seepage will turn into a fatal gusher. In addition to being optimists, most people would prefer not to try to fix something they do not fully understand. This is especially true if they are not even sure it is broken. For example, if a computer will no longer respond to commands, people shut it off, let it cool down, then reboot it. When the computer again responds, they are happy and will likely repeat the same behavior in the future. These human tendencies of optimism and avoidance of the uncomfortable unknown, which have generally served humans well, have resulted in the deaths of many cetaceans (pers. obs.). There are responsibilities that accompany the practice of veterinary medicine. Practitioners must consider themselves the animal’s advocate. Always temper what is easiest for people with what is best for the animal. It is easier to give a dolphin an oral medication in its food, but the
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
895
0839_frame_C40 Page 896 Tuesday, May 22, 2001 2:06 PM
896
CRC Handbook of Marine Mammal Medicine
best medication for the condition may require injection. To inject the animal daily may require its removal from its normal social group. To remove it from the social group may impact its interest in eating and its overall sense of well-being. These decisions as well as the day-to-day decisions affecting the life of the animal should be made in its interest. Most veterinarians are aware that many species of animals will mask signs of illness. This is also the case in dolphins and whales. In spite of this knowledge, many veterinarians will gravitate toward the most innocuous explanations for conditions like anorexia. It is true that there are reasons for anorexia that are not life-threatening, but the cetacean veterinarian will witness many surprise deaths if he or she assumes the cause of anorexia to be harmless. Anorexia is one of the most serious signs of disease presented to a cetacean veterinarian. Yes, it can accompany something as minor as a failure of understanding between the animal and the trainer, but it can also be as serious as a fulminating septicemia. More than once a veterinarian has said something like, “The dolphin is off its food today, so we will wait to see if the animal is still off its food tomorrow.” This may sound reasonable, but how does it sound if restated, “The dolphin is anorexic and alive today, let’s see if it is still alive tomorrow.” Trying to detect signs of illness in a species that naturally masks signs is problematic at best and sheer folly for a clinician with limited experience working with dolphins and whales. If a clinician thinks the animal may be sick, he or she should do something about it. If a clinician knows it is sick, it may be too late.
Clinical Examination The clinical examination of a cetacean includes history and examination of the animal and its environment. These are the same components as with any other animal undergoing a clinical evaluation. When a veterinary clinician hears that the dolphin or whale is not acting normally, the steps necessary to understand the problem are similar to those used for most other animal species. Clinicians need to adjust their thinking to accommodate social animals that spend their entire lives in water. A weak terrestrial animal will lie down; a cetacean will just float. Diarrhea leaves an easily recognized sign on land; in water no sign is left. The same is true with emesis if no solids are found on the pool bottom. Hematuria in a land animal leaves a tell-tale stain; in water the animal must be seen in the act of urinating. A sick animal may try to isolate itself, but in a social species it is not unusual for conspecifics to stay near an ill companion. A terrestrial herd animal butting a companion may be seen as aggression; in a dolphin, this may be an attempt to aid a sick member of the pod. Clinicians need to spend enough time observing the dolphins and whales under their care to understand the often subtle implications of what is observed.
History Collection of history is the first step in the diagnostic process. This reminder is important, because there is a tendency for clinicians to go straight to clinical laboratory findings when working with cetaceans. As is the case with other species, history is the most important part of a cetacean clinical examination. When collecting history on dolphins and small whales, do not overlook the social aspect of their lives. The quality of their interactions with trainers and pool mates can be just as important as their food intake, when looking for clues to the beginning of a period of illness. Observations of recent vomiting or diarrhea are obviously important. However, the fact that no one has seen these events does not mean they have not occurred. The appearance of a fish skeleton on the pool bottom can only have occurred as a result of emesis (unless the animal is filleting its own fish). The presence of vomited fish skeletons in the pool of a cetacean that is being treated with H2 blockers, or any other drugs that increase
0839_frame_C40 Page 897 Tuesday, May 22, 2001 2:06 PM
Cetacean Medicine
897
gastric pH, may suggest the animal’s inability to demineralize bones. In dolphins, the normal postprandial gastric pH of 1.5 will rapidly demineralize fish bones (unpubl. data). If the pH is too high, the fish bones will not demineralize and will begin to accumulate in the stomach until they produce gastric upset and vomiting (pers. obs.).
Visual Examination Visual examination of the patient both up close and at a distance is the next step in the clinical examination. When viewing the animal, examine its exterior and, at the same time, try to gain a sense of how the animal feels, and why its behavior has changed. To do this requires critical observation of the animal’s skin, eyes, body condition, its general appearance, and its interaction with its environment and companions. On at least one occasion while observing a whale that was anorexic, it became apparent that the animal, which had ceased eating, might not be the sick one. The anorexic animal was being very attentive to a companion, and this behavior is not typical of a sick individual. Blood samples collected from both animals confirmed that the companion was the sick individual. It is common for visual examination to reveal as much as, or more, than hands-on examination. Logical, commonsense knowledge of normal behavior goes hand-in-hand with medical competence, and together have a great deal to do with what a clinician can learn by looking at the animal.
How Does the Animal Feel? Are the animal’s eyes wide open or are they partially closed? This may indicate that it does not feel very badly; or it may indicate that the animal feels badly enough that it wants to be left alone and has very little interest in its surroundings. Of course, it may just indicate that its eyes hurt or light bothers its eyes. It is wise to note whether both eyes are affected. A simple method of determining if corneal injury is causing a closed eye is to squirt ophthalmic local anesthetic between the lids. This is accomplished by squirting the liquid, with a syringe, at the orbital fissure from a few centimeters distance. If some of the local anesthetic finds its way to the cornea, it will temporarily relieve the pain and allow the animal to open its eye for examination. This procedure is helpful, since it is difficult, if not impossible, to visualize a closed cetacean eye by forcing its lids open. Is the animal subdued or active? Most animals that feel sick prefer to rest and are not inherently active. Although they may swim along with pool mates, the instinctive efforts of a weak animal to keep up are usually easy to distinguish from normal swimming behavior. Is the animal alert or unaware? If the dolphin is alert and responsive to its surroundings, it is most likely not feeling ill. This fact does not mean that it is not ill, it just does not feel that way. Diseases like nocardiosis (see Chapter 16, Bacterial Diseases) or neoplasia (see Chapter 23, Noninfectious Diseases) can produce clinical laboratory indications of disease, but the animal may look as if it feels fine. This is especially true in the early stages of disease.
Buoyancy Buoyancy is a feature of physiology that is not examined in terrestrial animals. In cetaceans, buoyancy is too important to ignore in a visual examination. In simple terms, is the animal floating higher or lower than normal, and/or is it listing in the water? It is best to evaluate buoyancy in an animal at rest during the normal inspiratory breath-hold. Pneumonia in cetaceans may lead to an increase or a decrease in buoyancy. Observed changes in buoyancy do not automatically lead to a diagnosis, but observing these changes can help when developing differential diagnoses.
0839_frame_C40 Page 898 Tuesday, May 22, 2001 2:06 PM
898
CRC Handbook of Marine Mammal Medicine
Decreased Buoyancy
A decreased buoyancy will cause the animal to float lower in the water than normal. The clinician needs to be aware that a normal cetacean can decrease its buoyancy without expelling air by compressing its chest and abdomen, and this results in decreased displacement of the animal’s body. So the fact that the animal takes a deep breath and sinks to the bottom is not necessarily the result of a decrease in resting buoyancy. A true decrease in resting buoyancy is the result of diminished lung capacity. A space-occupying mass or fluid accumulation in or around the lungs is the probable cause of this alteration. Increased Buoyancy
Increased buoyancy usually results from abnormal gas accumulation in the gastrointestinal tract, abdomen, or thorax. A dolphin with a pneumothorax will appear much like a cork as it either bobs on the surface or displays difficulty diving or remaining submerged. Increased buoyancy can also be observed as a result of pain, which is relieved by inhibiting the abdominal and thoracic press that occurs normally during the inspiratory breath-hold. Listing
If listing is observed at rest, it is usually caused by a unilateral alteration in buoyancy, a postural aberration, or by choice. It is best to begin evaluating a listing animal by looking for any postural contributions. If the animal curves to one side, it may either be inducing listing or compensating for it, depending on whether the curve is toward or away from the least-buoyant side. Watching a dolphin swim that is listing at rest should reveal whether it has a righting defect. Even a dramatic unilateral buoyancy aberration can disappear when the animal is swimming, only to reappear when forward motion ceases. Normal newborn or early postpartum calves will often list at the surface if temporarily abandoned by the mother. Is the dolphin always listing to the same side? Listing is common when dolphins are looking up, as occurs when the water in a pool is lowered, because rolling to one side makes viewing the activities above much easier. Cetaceans with unilateral impairment of vision may roll to one side to make viewing activities around the pool easier. The causes of listing are usually differentiated by attention to history and visual examination, with confirmation provided by ultrasonography, centesis, and/or radiography (see Chapters 24 through 27).
Social Behavior To assess social behavior, it is important for the veterinarian to be familiar with the normal behavior of the group in question or at least to know who among the animal caretakers is a consistent and reliable observer. A cetacean’s social behavior or interaction with conspecifics provides a useful insight into its sense of well-being. A social animal that is isolating itself is most likely not feeling well. Occasionally, other dolphins will join a pod member that is not feeling well. If a dolphin or whale is being harassed by others, it deserves a second look from the veterinarian, since it may be ill. A weak female may be chased by males, as if she were sexually receptive; however, this probably occurs because the ill female is only able minimally to resist advances by the males. Humans are also a part of the social environment of captive dolphins. A change in the quality of interaction between trainer and dolphin is a potential early indicator of deteriorating animal health. Occasionally, one will observe a dolphin that actively avoids its trainer by staying as far away as it can. This animal may not be sick, but just telling its trainer that something is wrong with the environment or their relationship. Such changes need examination for clues to causes. It is the task of the clinician to sort out the subtleties of cetacean social behavior. A dolphin that does not have a positive social environment is likely to become a sick dolphin. There is
0839_frame_C40 Page 899 Tuesday, May 22, 2001 2:06 PM
Cetacean Medicine
899
evidence in humans and other social species that an individual’s social environment is a primary determinate of long-term health (House et al., 1988). The author chooses to express that thought as “Every dolphin needs a friend.”
Hands-On Examination After taking a history and completing a visual examination, feelings of uncertainty are not uncommon for the cetacean clinician. Unless a clinician has compelling evidence to the contrary, he or she should not be an optimist. Assume the worst. The usual next steps, hands-on physical examination and blood sampling (see Chapter 19, Clinical Pathology, for venipuncture sites), are generally preformed at the same time, because they require direct contact with the animal. The value of hands-on examination is limited by cetacean anatomy (see Chapter 9, Anatomy), but it is still a valuable source of information. Even auscultation has significant limitations. Auscultation is problematic because of the animal’s thick blubber layer, the rapid expiratory–inspiratory cycle, and loud transmitted sounds that can obliterate subtle rales. In spite of limitations, auscultation is an occasionally useful tool for evaluation of the thorax and abdomen. The rest of a complete examination should include a closer look at the eyes and examination of each of the body openings.
Urine Collection Male and female dolphins can be catheterized for urine collection, although many cetaceans have been trained to provide these samples on request. In bottlenose dolphins, a 5-French (1.67-mm) catheter is suitable for males, while an 8-French (2.7-mm) works well for females (pers. obs.) (see Chapter 19, Clinical Pathology).
Stool Samples If a stool sample is desired, a 16 French Levin-type stomach tube (Professional Medical Products, Inc., Greenwood, SC) works very well (pers. obs.). Levin tubes are open ended with side ports. Insert the tube into the rectum; then advance it into the descending colon. Then allow the sample to flow passively into the tube. Next, clamp or plug the tube to prevent the sample from running out and withdraw it. The tube should contain enough stool for cytology (see Chapter 20, Cytology) and culture (see Chapter 16, Bacterial Diseases). If the procedure is unsuccessful, try again using a small volume of saline flush. Do not apply suction, since there is a high likelihood it will cause some bleeding. Samples collected with swabs are also likely to contain blood contamination. If fecal occult blood is of interest, it is important to be aware that this test will always be positive if done on stool from cetaceans eating whole fish diets. If the patient is fed a diet consisting entirely of washed fish fillets for 2 to 3 days, normal stool will convert to occult blood negative (unpubl. data). This is a useful technique when there is an interest in determining if the patient is experiencing gastrointestinal bleeding that is not visible by endoscopy (see Chapter 27, Endoscopy). Always use a “normal” animal as a control for this procedure.
Milk Samples Milk samples are readily aspirated from the mammary glands of lactating females, with the assistance of external massage. A 60-ml catheter tip syringe, with a short length of tubing and an appropriately sized funnel, provides the pieces necessary to make a dolphin milk sample-collecting device. If a calf has recently nursed, it can be very difficult to obtain a useful sample.
0839_frame_C40 Page 900 Tuesday, May 22, 2001 2:06 PM
900
CRC Handbook of Marine Mammal Medicine
Blowhole The blowhole is another anatomical opening that deserves scrutiny. With adequate light, it is possible to visualize the nasal passages. The view is usually fleeting because of the rapid respirations common to cetaceans. Video or still photography through the open blowhole can provide a better look than is achievable with the naked eye. It will also yield documentation, which is useful for examining change in a known condition. Specimens can be collected for culture and cytology directly by swabbing affected tissue or indirectly by exposing an agar plate to exhaled breath (see Chapter 20, Cytology). These samples may be useful for evaluating a condition affecting the upper respiratory passages, but cannot be expected to provide accurate information about lower respiratory tract disease. Bronchoscopy and bronchoalveolar lavage are the most appropriate procedures for determining etiological agents in pneumonia cases (see Chapter 27, Endoscopy). In cetaceans, both processes are performed with minimal difficulty (Harrell et al., 1996; Hawkins et al., 1997; Reidarson et al., 1996).
Additional Diagnostic Aids Body Weight One of the most important, but often overlooked, features of a cetacean physical examination, is body weight. It is difficult, even for experienced individuals, to detect weight loss in cetaceans. When weight loss is noticed, it is often excessive. Whatever the difficulty associated with its determination, body weight is essential for medical and husbandry information. The advent of slide-out scales has made the weighing of dolphins and whales into a trained husbandry procedure.
Ultrasonography The real-time imagery of ultrasonography and the availability of waterproof transducers have resulted in the increased use of this techique in cetaceans (see Chapter 26, Ultrasonography). It is especially helpful for identifying and monitoring pulmonary disease (pers. obs.). Ultrasonography has also become a fundamental addition to the armamentarium of the cetacean reproductive physiologist (see Chapter 11, Reproduction).
Radiography Radiography has limitations in cetaceans because of their size, thick outer blubber layer, and poor abdominal contrast. There is also the logistical problem of taking the animal to the X-ray machine. Regardless of the barriers, a stationary grid makes diagnostic radiography possible on cetaceans of a size up to and including that of bottlenose dolphins (Tursiops truncatus) (Dalton et al., 1990) (see Chapter 25, Radiography).
Clinical Laboratory Tests Clinical laboratory test results are frequently the most informative part of a clinical examination. Historical data accumulated during routine examinations are very useful for comparison, when trying to interpret laboratory test results from an animal with a suspected illness. The decision to proceed with clinical laboratory evaluation should be an easy one, but often is not, because of a lack of access to an uncooperative patient or other unnecessary complications. The ability to collect a blood sample from a captive cetacean, whenever needed, is essential. A dolphin that is trained to present its flukes voluntarily to allow blood sampling may be unwilling
0839_frame_C40 Page 901 Tuesday, May 22, 2001 2:06 PM
Cetacean Medicine
901
to do so when it is feeling ill. Make contingency plans for this type of event long before it occurs. Without preplanning, the veterinarian and patient are both compromised. The clinician may pay with feelings of guilt, while death may be the toll for the animal. Obviously, the best plan is to have a medical pool that is usable any time, including during guest hours. After blood sample collection, its timely analysis must follow (see Chapter 19, Clinical Pathology). Preparation for a worst-case scenario is the responsibility of the cetacean facility director and the veterinarian. Disease is not confined to a 40-hour week and is never convenient. The price for failure to be prepared can be costly.
Clinical Pathology Case Example: Pulmonary Disease A case of pneumonia is a good way to highlight some important qualities of an effective cetacean medical program. Pneumonia is probably the most common serious disease of dolphins. Subtle signs and less than dramatic hematology and serum chemistry results are characteristic of this malady. Often anorexia or declining appetite is the only apparent sign. The veterinarian may not be made aware of change in the animal’s behavior until a day or more after the first signs appear. Frequent communication with husbandry staff is an important first line of defense. History and visual examination, in most cases, will yield few clues to indicate that the animal has pneumonia. Physical examination may be equally unrevealing. As mentioned earlier, auscultation is problematic in cetaceans. Regardless of the frequent lack of overt or objective clues to pneumonia’s existence, this disease in cetaceans can have rapidly fatal consequences. Clinicians must always remember how well cetaceans can mask signs of disease. One needs to be willing to take immediate action. Remember, “If you think there is a problem, deal with it. If you know there is a problem, it may be too late.” Clinicians must not convince themselves that, since the dolphin does not look too badly, it will be okay to wait until tomorrow before collecting a blood sample. The medical management of pneumonia is most successful with early initiation of treatment. This reinforces the need for rapid access to the animal, even if the animal chooses to be uncooperative, and the need for timely availability of clinical laboratory test results. This means that basic laboratory tests must be able to be run whenever there is a need, 24 hours a day, 7 days a week. With so little information to go on, how is the clinician to determine that the problem afflicting the animal is pneumonia? Assuming equivocal history and examination results, look to the hematology and serum chemistry tests to uncover the presence of inflammatory disease. One would reasonably assume that the total white cell and differential counts would give clear evidence of inflammatory disease. However, in cetaceans, these two parameters can be misleading, as a result of unpredictable variability. This adds to the difficulty of interpreting subtle white blood cell and differential count changes that accompany many pneumonias, and can lead the clinician to conclude that there is no significant inflammatory disease unless other parameters are examined (McBain, 1996). If inflammatory disease is present, consider the case to be pneumonia until there is compelling evidence to the contrary. The basis for this recommendation is the high probability that a sick cetacean with vague presenting signs has pneumonia. A pneumonia diagnosis can often be corroborated with ultrasonography and/or radiography. Indicators of Inflammatory Disease
The tests that have proved to be most helpful as corroborative indicators of inflammatory disease are reticulocyte count, white blood cell count, differential count, erythrocyte sedimentation rate, plasma fibrinogen, serum albumin, alkaline phosphatase, and serum iron (McBain, 1996)
0839_frame_C40 Page 902 Tuesday, May 22, 2001 2:06 PM
902
CRC Handbook of Marine Mammal Medicine
(see Chapter 19, Clinical Pathology). These tests are nearly all affected in cetacean pneumonia and other inflammatory diseases. Plasma fibrinogen is currently the most reliable indicator of inflammatory disease in cetaceans. This statement is true with the proviso that it is analyzed in the laboratory using the photo-optical test. The heat precipitation test for fibrinogen is prone to inaccuracy, and has limited utility. Plasma fibrinogen is not only useful for the early detection of inflammation, but also for determining when inflammation is under control. Elevations of as little as 20% above the animal’s high normal levels are important, but they will usually be elevated 50% or more with significant inflammation (unpubl. data). Erythrocyte sedimentation rate (ESR) is a traditional tool for detecting the presence and severity of inflammation. It is easily run without expensive equipment, which is the primary reason for its continued use. It is also a useful tool for corroborating an elevated fibrinogen level. The test is prone to fluctuations, which accounts for the fact that this test has lost favor in diagnostic testing of most other species. Many in cetacean medicine continue to use the ESR because of its historical familiarity. Serum iron decreases acutely in animals experiencing bacterial infection. This response is dramatic in cetaceans. It can plummet to levels 20% of normal or less, in a matter of 24 hours (unpubl. data). As a result, iron can be an excellent indicator of infection, but the test will not necessarily be a good indicator of the severity of the problem. Iron will fluctuate during the course of treatment, so it appears to be less reliable than fibrinogen for evaluating therapy. It is important to understand that this decrease in iron is protective for the host. The animal’s body is sequestering iron in the liver in a form that is not available to pathogenic bacteria that can readily utilize transferrin-bound iron in the serum (Lowenstine and Munson, 1999). Do not automatically begin supplementing iron just because serum levels are low. Hepatocellular damage is often associated with higher-than-normal serum iron levels (unpubl. data) (see Chapter 19, Clinical Pathology, Markers of Inflammation). Reticulocyte counts are not routinely requested by most clinicians. Few would think of it as an indicator of inflammatory disease. In cetaceans, however, the reticulocyte count is often low when the animal is experiencing a chronic low-grade infection. Many cetaceans have been treated for gastric ulcers because serum hemoglobin was low and no other parameters were significantly out of normal ranges. If a chronic, low hemoglobin exists, it is most likely because of either a slow, chronic bleed, or decreased red cell regeneration. Admittedly, there are other possible scenarios, but those two will cover the bulk of the cetacean cases (unpubl. data). The reticulocyte count will usually provide the information needed to differentiate blood loss from decreased regeneration. Values for normal cetaceans under a veterinarian’s care will be the best source of comparative data. Chronic low-grade pneumonia is not rare in cetaceans. It is often only clinically apparent because of slight elevation of inflammatory parameters and the presence of a low-grade nonregenerative anemia (anemia of chronic disease) (see Chapter 19, Clinical Pathology). The myth that most captive dolphins have gastric ulcers could cause a clinician to believe that a low serum hemoglobin is the result of bleeding ulcers; the reticulocyte count can save the cetacean veterinarian the embarrassment of treating a chronic pneumonia with iron and H2 blockers. Serum albumin levels regularly decrease below normal in the presence of infection (unpubl. data). The decrease can occur rapidly over a period of a few days. Serum albumin is not the first test to consult for evidence of infection, but it is good for confirming the results of other tests. There are obviously numerous causes of a drop in serum albumin (see Chapter 19, Clinical Pathology), so it is important to recall that low albumin is common in the presence of bacterial infection. Alkaline phosphatase has, for many years, been considered by cetacean veterinarians to be a very reliable prognostic indicator. Alkaline phosphatase levels in cetaceans are usually much
0839_frame_C40 Page 903 Tuesday, May 22, 2001 2:06 PM
Cetacean Medicine
903
higher than would be expected based on terrestrial species. Clinicians historically considered an alkaline phophatase of less than 50 IU/l in a killer whale (Orcinus orca) to be indicative of very serious disease, and values less than 25 IU/l to be equivalent to a death sentence (pers. obs.). Alkaline phosphatase normally declines with age, and also with a decrease in food intake. Even though alkaline phosphatase drops dramatically during illness, it is important to remember that it is affected by many other things (Fothergill et al., 1991; Dover et al., 1993) (see Chapter 19, Clinical Pathology). Total white blood cell count is understood by most veterinarians to be a reliable indicator of inflammatory response. There is no need to expand on this beyond repeating what was stated previously, that life-threatening pneumonia in cetaceans is frequently associated with an unremarkable total white blood cell count. Differential count is the means by which most veterinarians evaluate the nature and significance of a change in the white blood cell count. Differential counts in cetaceans are interpreted in much the same way they are in terrestrial animals. There is a potential pitfall, however. Both total white cell and differential counts may fluctuate unpredictably over a very short interval of time in cetaceans. Two blood samples taken from the same fluke vein less than 30 s apart may have different total white cell and differential counts. These variations are large enough to affect interpretation. The clinician must look to other clinical data to support diagnostic conclusions (McBain, 1996).
Therapeutics Surgery The therapeutic options in cetaceans are much the same as in terrestrial species, with the exception that abdominal surgery is rarely considered. The recent introduction of laparoscopy, as an alternative, may change this deficiency (see Chapter 27, Endoscopy). The bulk of surgery in cetaceans has been limited to dentistry, wound management, abscess treatment, superficial biopsy, endoscopic procedures, and mandibular and maxillary fracture repair. There are a couple of features regarding abscesses in cetaceans that are worthy of attention. Purulent infections deep to the blubber layer will tend to dissect along the muscle/blubber interface rather than rupturing through to the surface. These infections are often difficult to identify visually. In the author’s experience, abscesses dissect or migrate dorsally, rather than ventrally as is expected in terrestrial species. This finding is based on a small number of cases (pers. obs.). The tendency to migrate in a direction contrary to that in a land animal may be related to the water pressure gradient on the animal’s body. In a normal swimming posture, the water pressure on the ventrum of a cetacean will be greater than on its dorsal aspect. In spite of its limitations, surgery should not be ruled out as an option for cetaceans.
Medical Therapy Medical treatment options for cetaceans are virtually the same as for their terrestrial counterparts, with one notable exception (see Chapter 31, Pharmaceuticals). In cetaceans, the administration of long-term intravenous therapy is an option that is rarely used, or considered. Oral Route
The oral route is preferred for the administration of fluids and pharmaceuticals to cetaceans. Feeding medication hidden in fish is the simplest and most common approach used for per os (PO) administration. If anorexic, a stomach tube becomes the best means for administering oral medication, including fluids. Cetaceans can accommodate a small amount of saltwater
0839_frame_C40 Page 904 Tuesday, May 22, 2001 2:06 PM
904
CRC Handbook of Marine Mammal Medicine
consumption each day, but may need additional fluid support if they are anorexic or are consuming excessive amounts of seawater (pers. obs.). Nauseous cetaceans will sometimes run elevated serum sodium and chloride levels that will decrease in response to administration of oral tap water. Anorexic cetaceans normally obtain water as a metabolic by-product of fat metabolism and may also drink small amounts of seawater (Telfer et al., 1970) (see Chapter 36, Nutrition). If the animal is sick, its fluid needs may outpace the available innate water supplies (see Chapter 30, Intensive Care). These animals usually respond very well to oral fluids. A stomach tube is relatively well accepted by most cetaceans, and many have been conditioned to voluntarily accept a gastric tube. Subcutaneous Route
If vomiting of oral fluids is occurring, the “subcutaneous” route can be used effectively. For the purpose of this discussion “subcutaneous” in cetaceans means the interface between the blubber layer and the skeletal muscle layer. This space can receive fluid amounts equivalent to the subcutaneous space in many terrestrial animals (pers. obs.). To administer subcutaneous fluids to a cetacean, use fluid bags with a standard intravenous infusion setup. Advance the needle through the blubber with the fluids under gravity pressure and the tubing clamp released (open). The fluids will begin to flow freely when the needle enters the subcutaneous space. Most cetacean practitioners use pressure cuffs (Pressure Administration Cuff, McGaw, Inc., Irvine, CA) to speed the flow once the needle is in the proper location. Intramuscular Route
In cetaceans, intramuscular injection requires a longer needle than might normally be considered necessary, because of the thickness of the animal’s skin and blubber. The injections are made off the midline, slightly anterior to, parallel to, or just posterior to the dorsal fin. It is the author’s rule to limit intramuscular injection volume to a maximum of 20 ml per site. This arises from the concern that larger volumes have led to apparent ischemic necrosis at the site of the injection (pers. obs.). Exercise care to avoid the thoracic cavity, if injecting in the dorsal musculature anterior to the dorsal fin. If the animal is wiggling or thrashing during an intramuscular injection, there is also a real potential for the needle to be sheared off by the heavy fascial planes and muscle sheaths. Intravenous Route
Intravenous injection of medication can be accomplished in cetaceans via a fluke vessel if the volume is low, and in most cases the medication is not harmful if delivered perivascularly. If slow infusion or repeated administration of intravenous pharmaceuticals is necessary, an indwelling catheter may be required. The maintenance of indwelling catheters is at best difficult in an animal that wants to be swimming. To accomplish long-term infusion, the animal must be confined to a very small pool or box, such as a transport container. Neither of these options is well received by most cetaceans. A better option, if intravenous therapy is required, is to try to accomplish the needed infusions with short periods of confinement, allowing the animal to swim between treatments. This requires either leaving the catheter in place, in the hope that it will remain, or replacing it for each treatment. Both of these methods have been used with limited success (McBain and Reidarson, 1994; Van Bonn et al., 1996; Stetter et al., 1997; Robeck and Dalton, 2000). Topical Route
Topical treatment is rarely used in cetaceans for the obvious reason that the medication usually washes away very rapidly. There have been attempts to treat corneal lesions with topical medications. It is possible that this strategy may succeed if the animal is keeping its eye(s)
0839_frame_C40 Page 905 Tuesday, May 22, 2001 2:06 PM
Cetacean Medicine
905
tightly closed. The clinical response to this type of therapy is difficult to evaluate because corneal lesions usually heal without treatment. So, what is the best treatment for corneal ulcers in a cetacean? To recommend doing nothing has worked in the past. The best treatment probably has not been tried as yet and possibly not even contemplated.
Final Thoughts When embarking on a medical treatment regimen, it is important to remember that many medications have had limited use in cetaceans. If a veterinarian is going to use a pharmaceutical for the first time, it is wise to seek the advice of a colleague who has had experience with that drug. The bulk of published information is based on a small number of trials. Someone always has to be first. A veterinarian in this position must realize that adverse reaction or unexpected results can occur. The author has had this experience many times. It is obligation of individuals that have negative experiences to share their failures with others, so the failure will not occur again. There are no experts in cetacean medicine. There are not enough cetaceans in captivity to provide any one clinician with adequate experience to qualify as an expert. The opportunities for gaining experience are limited by a relatively small number of cases. There has also been a historical tendency to delay intervention or deny its necessity until it is too late to achieve a successful outcome. Little is learned from those cases beyond the recognition that to advance knowledge of therapy it must be attempted. Veterinarians have accommodated the paucity of cetacean cases by sharing their problems with others in the cetacean medicine field. This process has been expedited by the formation of professional groups such as the International Association for Aquatic Animal Medicine (IAAAM) and the European Association for Aquatic Mammals (EAAM). The only thing worse than observing the death of a patient is to find that it was preventable. Sharing problems and solutions is essential to the growth of collective medical knowledge.
Acknowledgments The author acknowledges the assistance of Linda Dunn, Judy St. Leger, Tom Reidarson, Marty Haulena, and Andy Draper. Linda was the final grammar and punctuation checker, as well as guardian of the office door when the author needed quiet time. Judy and Tom were the reality checks. Without these three, this chapter would still be a work in progress. Marty and Andy are thanked for taking on the task of review. The author knows they “volunteered” for that duty, but they deserve thanks anyway. Last, I want to thank my wife, Judy, for giving my life purpose and helping me add clarity to some of my germinal thoughts.
References Dalton, L.M., Mathey, S.W., and Hines, R.S., 1990, Radiology as a diagnostic aid in marine animal medicine, in Proceedings of the 21st Annual International Association for Aquatic Animal Medicine, 21: 15–18. Dover, S., McBain, J., and Little, K., 1993, Serum alkaline phosphatase as an indicator of nutritional status in cetaceans, in Proceedings of the 24th Annual International Association for Aquatic Animal Medicine, 24: 44. Fothergill, M.B., Schwegman, C.A., Garratt, P.A., Govender, A., and Robertson, W.D., 1991, Serum alkaline phosphatase-changes in relation to state of health and age of dolphins, Aquat. Mammals, 17(2): 71–75.
0839_frame_C40 Page 906 Tuesday, May 22, 2001 2:06 PM
906
CRC Handbook of Marine Mammal Medicine
Harrell, J., Reidarson, T., McBain, J., and Sheets, H., 1996, Bronchoscopy of the bottlenose dolphin (Tursiops truncatus), in Proceedings of the 27th Annual International Association for Aquatic Animal Medicine, 27: 33. Hawkins, E.C., Townsend, F.I., Lewbart, G.A., Stamper, M.A., Thayer, V.G., and Rhinehart, H.L., 1997, Bronchoalveolar lavage in a dolphin, J. Am. Vet. Med. Assoc., 211: 901–904. House, J.S., Landis, K.R., and Umberson, D., 1988, Social relationships and health, Science, 241: 540–545. Lowenstine, L.J., and Munson, L., 1999, Iron overload in the animal kingdom, in Zoo and Wild Animal Medicine 4, Fowler, M., and Miller, E.R. (Eds.), W.B. Saunders, Philadelphia, 260–268. McBain, J., 1996, Clinical pathology interpretation in Delphinidae with emphasis on inflammation, in Proceedings of Annual Meeting of the American Association of Zoo Veterinarians, 308. McBain, J., and Reidarson, T., 1994, A case of renal failure in a Pacific pilot whale (Globicephala macrorhynchus) and attempted therapy, in Proceedings of the 25th Annual International Association for Aquatic Animal Medicine, 25: 23. Reidarson, T., McBain, J., and Harrell, J., 1996, The use of bronchoscopy and fungal serology to diagnose Aspergillus fumigatus lung infection in a bottlenose dolphin (Tursiops truncatus), in Proceedings of the 27th Annual International Association for Aquatic Animal Medicine, 27: 34–35. Robeck, T., and Dalton, L., 2000, Treatment of cutaneous, subcutaneous Apophysomyces elegans, a mucormycotic fungi infection in a bottlenose dolphin (Tursiops truncatus) with the new antifungal agent, Nyotran®, in Proceedings of the 31st Annual International Association for Aquatic Animal Medicine, 31: 374–375. Stetter, M.D., Calle, P.P., McClave, C., and Cook, R.A., 1997, Marine mammal intravenous catheterization techniques, in Proceedings of the American Association of Zoo Veterinarians Annual Meeting, 194–196. Telfer, N., Cornell, L.H., and Prescott, J.H., 1970, Do dolphins drink water? J. Am. Vet. Med. Assoc., 157(5): 555–558 Van Bonn, W., Jensen, E., Miller, W.G., and Ridgway, S., 1996, Contemporary diagnostics and treatment of bottlenose dolphins: A case study, in Proceedings of the 27th Annual International Association for Aquatic Animal Medicine, 27: 36–37.
0839_frame_C41 Page 907 Tuesday, May 22, 2001 2:07 PM
41
Seals and Sea Lions Frances M. D. Gulland, Martin Haulena, and Leslie A. Dierauf
Introduction Seals and sea lions are among the marine mammals most commonly maintained in display facilities and most commonly rehabilitated after stranding. The biology, natural history, physiology, and ecology of pinnipeds are reviewed by Riedman (1990) and Reynolds and Rommel (1999). As veterinarians take an increasing interest in the management of marine mammals, information on their special care and medicine is developing rapidly. Most of the medicine presented here has been learned from care of the commonly stranded and displayed species of pinnipeds, particularly the California sea lion (Zalophus californianus) and the harbor seal (Phoca vitulina). Disease problems may be very different in some of the less accessible species, and in wild animals on rookeries, from those in these two species.
Husbandry Pools, Haul-Out Areas, and Enclosures Pinnipeds require both water and haul-out space on land. Although seals and sea lions can survive in pens without access to water for weeks at a time, they appear more content when given free access to water. Most pinniped species will eat more readily if offered food in water rather than on land. Fur seals will readily defecate and groom when given access to a pool, while if left in a dry haul-out area they may appear clinically depressed. Pools are also important for exercise, physical therapy, mental stimulation, and reducing soiling. In the United States, housing requirements for pinnipeds on display are governed by the Animal and Plant Health Inspection Service, U.S. Department of Agriculture (APHIS, USDA), which sets minimum sizes for both pools and haul-out areas for pinnipeds. These standards change, so the reader is advised to check current standards directly with the USDA. Currently, pool size and haul-out area sizes are based on the body length of the species housed. Pool design should be aimed at accommodating the behavioral needs of the animals housed, as well as maintaining water quality (see Chapter 35, Water Quality). For otariids, the pool may be sunken below ground or raised, with access by ramps. As phocids do not have the climbing abilities of sea lions, they are better housed with sunken pools where the water level is close to the edge, so that exiting the pool is easy. Gently sloping sides or edges below the surface of the water allow seals to rest in shallow water. Cages around pools and haul-out areas should be constructed of nontoxic, corrosion-resistant, nonabrasive material, and should be
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
907
0839_frame_C41 Page 908 Tuesday, May 22, 2001 2:07 PM
908
CRC Handbook of Marine Mammal Medicine
of sufficient height to contain the species housed (otariids climb, while phocids do not). The choice of fresh or salt water for housing is often based on economics or logistics, rather than medicine. Although pinnipeds are often housed in freshwater systems, ophthalmic problems may be more common in fresh water than in salt water (Dunn et al., 1996). When housed in fresh water, care must be taken to provide oral salt supplementation (see below). Lighting should mimic the appropriate natural photoperiod for the species as closely as possible. Harbor seals maintained in continuous light conditions have been shown to have disrupted molt cycles that revert to normal when a natural photoperiod is reinstated (Mo et al., 2000). Extremes of both heat and cold should be prevented, although, generally, most species are better able to tolerate cold than heat. Geraci (1986) states that healthy, robust harbor, gray (Halichoerus grypus), harp (Pagophilus groenlandicus), and ringed seals (Phoca hispida) can tolerate water at freezing temperatures, and air temperature below −20°C (–4°F), although a northern elephant seal (Mirounga angustirostris) died after being exposed suddenly to an outdoor temperature of −15°C (5°F) for 30 min. Hyperthermia can be avoided by providing access to shade, pools, or sprinklers when ambient temperatures rise above 26°C (79°F). Hypothermia is rare, but can be a problem in undernourished animals. Provision of waterproof heating pads or kennel areas with heat lamps can prevent hypothermia.
Feeding Although free-ranging pinnipeds feed on a variety of prey, captive animals are usually maintained on a diet of herring (Clupeidae), smelt (Osmeridae), mackerel (Scombridae), capelin (Mallotus villosus), and squid (Loligo spp.). As herring is a relatively fatty fish, it is commonly fed to produce rapid weight gain. Details of the nutritional content of different diets and the methods to calculate calorific requirements of marine mammals are provided in Chapter 36 (Nutrition) while hand-rearing techniques are given in Chapter 37 (Hand-Rearing). As a rough guideline, young pinnipeds are fed 8 to 15% of their body weight of food per day, older animals 4 to 8% per day. Daily supplementation with sodium chloride at 3 g/kg fish, thiamine at 25 to 35 mg/kg fish, and vitamin E at 100 IU/kg fish is recommended to prevent nutritional disorders associated with captivity (see Chapter 36, Nutrition). When supplementing an animal’s diet, it is advisable to feed a fish containing the supplements prior to the main feed, to ensure the animal receives all of its medications.
Restraint The methods commonly used to restrain pinnipeds may be classified into physical, mechanical, and chemical. The factors that dictate the choice of an appropriate restraint method include human safety, safety to the animal, and the ability to accomplish the desired objective. For example, if the desired objective is to perform an abdominal ultrasound, the use of a mechanical squeeze may be safe for the human operator and fairly safe for the animal, but may position the animal in such a way that exposure of the abdomen is inadequate to carry out the ultrasound examination. Different types of restraint are often used in combination. For example, a chemical sedative agent such as a benzodiazepine may be given to an animal to augment physical restraint.
Physical Restraint Physical restraint is limited by the size and species of the animal, the animal’s level of aggression, and the experience and physical ability of the restrainers. It is usually very safe for the animal, but human safety may be a concern with larger animals, and access to the animal, once restrained,
0839_frame_C41 Page 909 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
909
may be limited. Physical restraint requires a thorough knowledge of the behavior and anatomy of the species being restrained. For example, larger otariids have tremendously strong forelimbs in comparison with phocids. The foreflippers may have to be secured by additional personnel to prevent the animal from gaining leverage and rising up (Gentry and Holt, 1982). Creative use of towels, blankets, bags, and nets will aid physical restraint and increase the safety of personnel.
Mechanical Restraint Mechanical restraint is limited by the availability of adequate equipment, the cost of which varies considerably. There are many varieties of mechanical restraint devices that have been used with pinnipeds, including chutes, herding boards, restraint boards, stretchers with straps, restraint boxes, squeeze cages, and slings (Cornell, 1986; Gentry and Casanas, 1997). As some mechanical restraint equipment can be very large and heavy, it may be difficult to use in field situations. In general, mechanical restraint devices are designed to maximize safety to human operators, but may pose some risk to the animal. Some restraint boards require a padded, hinged guillotine to secure the neck, and may partially obstruct the airway. Mechanical squeeze cages should be used with caution, since it is possible to use excessive pressure and cause trauma or interfere with adequate ventilation. Full access to the animal may be limited by some mechanical restraint devices.
Chemical Restraint The ability to use chemical restraint relies on the expertise of the operators, and often requires the presence of a specially trained veterinarian. The availability of some immobilizing agents may be limited because of expense, or because they are controlled substances (see Chapter 29, Anesthesia). Some agents used at The Marine Mammal Center (TMMC), Sausalito, CA, for sedation and immobilization of phocids include diazepam (0.1 to 0.2 mg/kg IM/IV), butorphanol (0.05 mg/kg IM), tiletamine/zolazepam (0.8 mg/kg IV), propofol (3 to 5 mg/kg IV), and isoflurane. Some agents used at TMMC in otariids include midazolam (0.1 to 0.2 mg/kg IM), medetomidine (140 µg/kg IM) + ketamine (2.5 mg/kg IM) with atipamezole (200 µg/kg IM) reversal, medetomidine (70 µg/kg IM) + tiletamine/zolazepam (1.0 mg/kg IM) with atipamezole (200 µg/kg IM) reversal, and isoflurane.
Physical Examination A complete physical examination of a pinniped is often difficult to accomplish. Physical examination may have to be quick and cursory for an untrained animal that requires restraint in order to maximize safety to the animal and operators. The minimal data collected should include an age estimate, sex, standard length, and a body condition score. In contrast, a trained animal may be directed to lie out in different positions for enough time to allow a more complete examination. The physical examination begins by observing an animal from a distance, noting behavior, attitude, locomotion, paresis, and any obvious discharge, masses, swellings, or asymmetry. A thorough knowledge of the normal appearance and behavior of the species under investigation is required. Furthermore, adequate examination of the musculoskeletal system requires a thorough knowledge of the species’ normal patterns of movement. Palpation of limbs and body contours may reveal swelling, masses, or areas of increased temperature, indicative of inflammation (see Chapter 28, Thermography). Flexion of limbs may show increased or decreased mobility, or crepitus suggestive of fractures or arthritis. Thick blubber layers may obscure subcutaneous masses or abscesses, may prevent thorough abdominal palpation, and may interfere with thoracic auscultation. Thus, young or thin animals can often be examined more
0839_frame_C41 Page 910 Tuesday, May 22, 2001 2:07 PM
910
CRC Handbook of Marine Mammal Medicine
easily. Abdominal masses, such as pelvic tumors, have been diagnosed in physically restrained adult California sea lions by abdominal palpation. Thick blubber layers may also prevent diagnosis of ascites. However, a pendulous and firm abdomen in young animals is suggestive of fluid in the abdomen. Fluid waves may be detected in the abdomen of thin animals.
Diagnostic Techniques Blood Collection Phocids may be sampled relatively easily from the extradural intravertebral vein or the plantar interdigital veins of the pelvic flippers. Choice of site is governed by size of the animal, ease of restraint, volume of blood to be collected, and age of the animal. Use of the epidural intravertebral vein in northern elephant seal pups has resulted in inadvertent bone marrow contamination of samples (Goldstein et al., 1998), so caution should be used when using this site in young phocids. Illustrations showing venipuncture sites are presented in Chapter 19 (Clinical Pathology). To collect blood from the epidural intravertebral vein, restrain the seal in ventral recumbency, and locate the spines of lumbar vertebrae 3 and 4 by palpating the iliac crest and moving cranially. In fat seals, the tail is a useful guide to locate the midline. Insert the needle perpendicularly between the two vertebral bodies until blood is observed in the needle hub. The size of the needle depends upon the size and condition of the seal. A 20-gauge, 1-in. needle is preferred for harbor seal pups, while an 18-gauge, 3-in. spinal needle is needed for fat adult harbor seals. The plantar interdigital veins of the hind flippers are located by inserting the needle at 10 to 20° to the skin directly over the second digit, or medial to the fourth digit, at the origin of the interdigital webbing. The sample from this site is often an arterial/venous mix, so postsampling bleeding must be avoided by applying firm pressure. In otariids, the caudal gluteal vein is commonly used as it can be readily accessed while the animal is manually restrained. It is located lateral to the sacral vertebrae, one third of the distance between the femoral trochanters and the base of the tail. A 21-gauge, 1-in. needle is suitable for thin fur seal and sea lion pups, a 1.5-in. needle for animals up to about 150 kg, and a 3-in. needle for larger sea lions. The interdigital vessels of the hind flippers of otariids are small, but can be dilated for visualization by placing the hind flipper on a bag of warm fluids, or pouring warm water over the flipper. Tourniquets placed over the tarsus may aid in dilating interdigital vessels. Because of the small size of the vessels and the slow rate of blood collection, a heparinized butterfly needle and catheter can be useful. The jugular vein can be used as a collection site in otariids, but it can be frustrating to locate. It runs from the angle of the jaw to the thoracic inlet, and may be sampled from the angle of the jaw, the midcervical region, or at the base of the neck. The subclavian vein may be used in emergency situations in otariids. It is located by palpating the sternum and the first rib, and inserting the needle perpendicularly into the angle between the two. Because of the size of this vessel and its proximity to the heart, this site is only recommended for emergency use.
Urine Urine is most commonly collected during urination, although catheterization of pinnipeds is possible using the technique similar to that for domestic dogs. Abdominal compression of anesthetized California sea lions has been used successfully to collect urine (Acevedo, pers. comm.). Cystocentesis is also used to collect urine from anesthetized animals, by inserting a
0839_frame_C41 Page 911 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
911
sterile 2-gauge, 3-in. needle cranial to the pelvis and ventrally. Little is known about clinical changes in urine composition with dehydration and disease. However, urine samples can be useful in the detection of leptospiruria, and exposure to water-soluble toxins excreted in urine, such as domoic acid.
Cerebrospinal Fluid Cerebrospinal fluid may be collected from the epidural space at the level of the atlanto-occipital joint from both phocids and otariids in lateral recumbency by a technique similar to that used in the domestic dog.
Biopsies Skin and blubber biopsies are often performed on marine mammals for research purposes. Samples may be collected with disposable biopsy punches, or by using a scalpel blade and forceps. Local anesthetic infiltrated around the site provides analgesia, but sedation may be needed if the animal cannot be adequately restrained to prevent movement. After removal of skin or blubber samples (typically less than 2 cm in diameter), biopsy sites are usually left unsutured and allowed to heal by secondary intention, to reduce likelihood of retaining infected tissue. Abdominal organ biopsies may be performed under general anesthesia using laparoscopic techniques similar to those used in small animal surgery (see Chapter 27, Endoscopy).
Therapeutic Techniques Therapeutic agents may be delivered to pinnipeds in a variety of ways. Choosing a route of administration must take into account the required frequency, efficacy, and feasibility of administration.
Topical The use of topical agents may be limited in pinnipeds, especially if the agents are readily removed once the animals enter the water. However, most pinnipeds can remain out of water for prolonged periods of time (Geraci, 1986), although thermoregulation, ingestion, excretion, and coat quality may be affected. Topical administration may also require restraint, although wounds can be sprayed lightly with chlorhexidine or povidone iodine solutions from a short distance away. The frequency that some topical agents need to be delivered may limit their use in some instances. Many ophthalmic products, for example, require application at least four times per day. The application of topical ophthalmic agents may also be limited by the constant lacrimation typical of pinnipeds, by animal aggression, and by blepharospasm. The use of subpalpebral lavage systems has been effective (Borkowski et al., 1999).
Oral Oral medications are easily administered if an animal is eating. Pills, capsules, and small amounts of liquids may be placed into the coelomic cavity of fish. Many animals can detect pills in overstuffed fish and may refuse to eat them. Oral medications can be crushed and mixed into formula or fluids that are either bottle-fed or stomach-tubed to animals. Moderately dehydrated animals may be given fluids via a stomach tube.
0839_frame_C41 Page 912 Tuesday, May 22, 2001 2:07 PM
912
CRC Handbook of Marine Mammal Medicine
Aerosol Nebulization therapy has been used in pinnipeds that have been placed into sealed cages or pens. Care must be taken to ensure the animal does not develop hyperthermia during treatment in enclosed spaces. Placing ice bags underneath a cage grate can be a useful addition when treating small pinnipeds.
Subcutaneous Most pinnipeds may be given subcutaneous fluids, as long as the animal can be maintained in a certain position for a prolonged period of time. This may require some restraint, and may be more easily accomplished in debilitated animals. The use of smaller pens or fencing material to limit movement may facilitate subcutaneous fluid administration. The subcutaneous route may not be useful in severely debilitated animals with some degree of circulatory collapse that are no longer able to absorb fluids from the subcutaneous space. Other medications such as ivermectin and certain antibiotics have also been delivered subcutaneously. The most common site for subcutaneous administration of medications is the craniodorsal thorax between the scapulae.
Intramuscular Intramuscular injections require the use of relatively long needles to place medications under the blubber. Injections into the blubber may cause sterile abscess formation and/or result in poor absorption. Intramuscular injection usually requires some degree of restraint, although handling may be minimized with the use of alternative delivery methods such as pole-syringes or darts. However, accurate placement of medications may be compromised with their use. The muscles surrounding the pelvis, femur, and tibia are suitable for intramuscular injection in phocids, while in otariids the large muscles overlying the scapulae as well as those around the pelvis, femur, and tibia are appropriate sites.
Intravenous Compared with terrestrial mammals of similar size, there are a limited number of sites for intravenous access in marine mammals (see Blood Collection above; Chapter 19, Clinical Pathology). Catheters have been placed and maintained for prolonged periods of time in the extradural intravertebral sinus of phocids and in jugular, subclavian, and interdigital veins of anesthetized otariids. However, catheters are difficult to maintain in most otariids for any length of time. The ability of pinnipeds to shunt and pool blood away from peripheral tissues makes some peripheral venipuncture sites a poor choice for emergency drug administration.
Intraosseous Intraosseous catheters have been placed into the medullary cavity of the proximal tibia in otariids to provide fluid therapy and for administration of other therapeutic agents, including blood transfusions. Unfortunately, the maximum flow rate has not been sufficient to maintain hydration in larger animals. The catheters are difficult to maintain unless the animals are sufficiently debilitated and unable to remove the catheter.
Intraperitoneal The intraperitoneal route has been used to deliver fluids to severely compromised pinnipeds, especially in otariids with poorly accessible peripheral blood vessels. Care must be taken to avoid damaging vital organs, or introducing bacteria to the abdominal cavity either via a contaminated needle or by puncturing the gastrointestinal tract.
0839_frame_C41 Page 913 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
913
Diseases Details of viral, bacterial, fungal, parasitic, and noninfectious diseases of pinnipeds are provided in Chapters 15, 16, 17, 18, and 23, respectively. To avoid repetition, this chapter focuses on the clinical signs of these diseases, described by affected organ system for ease of differential diagnosis, and discusses treatment. Drug dosages for recommended therapeutic agents are given in Chapter 31 (Pharmaceuticals).
Integumentary System Common viruses infecting the skin of seals and sea lions are poxviruses and caliciviruses (see Chapter 15, Viral Diseases). Seal pox usually occurs in animals that have been recently weaned or been brought into captivity and is contagious to susceptible animals (Hastings et al., 1989). It causes skin nodules 0.5 to 1 cm in diameter that gradually increase in size over the first week, and may ulcerate (Figure 1A). Satellite lesions appear in the second week, and may spread rapidly. The lesions are usually self-limiting and regress after 4 weeks, although some have persisted for months. Animals usually remain active when affected, although lesions around the lips and eyes may cause sufficient discomfort to reduce appetite. Marked neutrophilia may occur in association with nodule development. Diagnosis is based on the presence of eosinophilic intranuclear inclusions in epidermal cells in skin biopsies. Treatment is usually unnecessary, although broadspectrum antibiotics may be needed to control secondary bacterial infections. San Miguel sea lion virus causes vesicles on both dorsal and ventral surfaces of the flippers, occasionally around the lips, on the dorsum of the tongue, and on the hard palate of California sea lions (Gage et al., 1990; Smith and Boyt, 1990; Van Bonn et al., 2000). The vesicles usually erode, leaving rapidly healing ulcers, but may become secondarily infected by bacteria, especially in malnourished animals. Diagnosis is based on isolation of the virus from aspirated vesicular fluid. Treatment is supportive, aimed at preventing secondary infection and enhancing nutritional status of the sea lion. Occasionally, stranded sea lions are observed with severe gangrenous necrosis of the phalanges (Figure 1B). Although progression of these lesions has not been observed, it is suspected that they may result from vesicles that became infected with bacteria. These lesions are treated by debridement and systemic antibiotic therapy based on culture and antibiotic sensitivity of bacteria from the lesions. Herpesviruses have been isolated from harbor and gray seals with small erosive skin lesions, and observed in epithelial plaques in harbor seals and California sea lions (see Chapter 15, Viral Diseases). Although infrequent, herpesviruses should be considered in the differential diagnosis of skin lesions, and skin biopsies should be examined for inclusions. Multifocal circular ulcers 1 to 2 cm in diameter have been observed in California sea lions and elephant seals. Histologically these appear to be the consequence of vasculitis and thrombosis. Microabscesses are also common on the ventral abdomen of sea lions following septicemia (Figure 1C). Diagnosis is based on the histological appearance of biopsies, and treatment with systemic antibiotics is recommended. Subcutaneous abscesses due to infection with Mycobacterium chelonii in a captive gray seal (Stoskopf et al., 1987) and M. smegmatis in a captive California sea lion (Gutter et al., 1987) were diagnosed after culturing the organisms from aspirated fluid. The gray seal was treated successfully with minocycline, while the sea lion died with pulmonary abscesses. Acanthosis and alopecia associated with Candida albicans and Fusarium spp. infections are typically observed at mucocutaneous junctions, around nail beds, and in the axillae (see Chapter 17, Mycotic Diseases) (Figures 1D and E). Lesions are most often observed in captive animals maintained in fresh water. Diagnosis is based on culture and histological examination of biopsies.
0839_frame_C41 Page 914 Wednesday, May 23, 2001 11:06 AM
914
CRC Handbook of Marine Mammal Medicine
D
A
E
B
F
C
G FIGURE 1 Common skin conditions of pinnipeds. (A) Pox in a California sea lion. (B) San Miguel sea lion virus complicated by gangrenous necrosis in a debilitated California sea lion. (C) Vascular thrombosis resulting in ulceration in a California sea lion. (D) Periocular Fusarium spp. in a northern elephant seal. (E) Fusarium spp. in the axilla of a northern elephant seal. (F) Lice in California sea lions. (G) Northern elephant seal skin disease.
0839_frame_C41 Page 915 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
915
Topical treatment is difficult without limiting access to water, but systemic treatment of an elephant seal with fluconazole at 0.5 mg/kg was effective in clearing clinical signs (Gulland, unpubl. obs.). Alopecia, broken hair shafts, and pruritus are common in debilitated seals and sea lions associated with lice infestation (Figure 1F). Most infections are species specific: the California sea lion louse is Antarctophthirius microchi, and the harbor seal louse is Echinophthirius horridis. The lice may be observed with the naked eye, and are readily treated with ivermectin, dichlorvos, or disophenol systemically, or topical rotenone louse powder. Demodicosis, also characterized by alopecia and pruritus, has been observed in California sea lions and northern fur seals (Callorhinus ursinus) (Spraker, pers. comm.). Diagnosis is based on histological detection in biopsies, and treatment with amitraz has been effective (Sweeney, 1986b). Skin wounds are common consequences of trauma in stranded animals (see Chapter 23, Noninfectious Diseases). Net entanglements, fishhooks, and gunshot injuries are especially common in California sea lions (Goldstein et al., 1999). Diagnosis of gunshot injuries is dependent upon radiographic detection of lead fragments or pellets, or recovery of the projectile, although many wounds suggestive of exit wounds are observed in pinnipeds from which no evidence of gunshot can be detected. Differential diagnoses include bite wounds (usually paired holes of similar size) and bird damage. The characteristics of shark bite wounds vary with species of shark (see Chapter 23, Noninfectious Diseases). Management of traumatic skin wounds is based on removal of foreign bodies and devitalized tissue, control of infection, and promotion of healing, as in domestic animals. Placing an animal in salt water rather than fresh water may enhance wound cleansing. Tetracycline and penicillin have been used to treat shark wounds, as Vibrio spp. and Clostridium spp. are frequently isolated from these wounds (Pavia et al., 1989; Klontz et al., 1993). Neoplastic skin lesions are differentiated from granulating wounds by biopsy and histological examination. Alopecia and acanthosis have occurred in captive harbor seals that failed to molt when maintained in constant photoperiod (Mo et al., 2000). Diagnosis was based upon history, and restoration of a natural photoperiod resulted in new hair growth. Cutaneous lupus erythematosus was diagnosed in a captive gray seal that for 9 years had continuous ulcerative nasal dermatitis and intermittent ulcerative dermatitis of the nail beds and dorsum of the body (Burns, 1993). Diagnosis was based on characteristic histological lesions, which included a lymphoplasmocytic interface dermatitis, hydropic degeneration of basal epithelial cells, acanthosis, and hyperkeratosis. Intralesional immunoglobulins were identified by immunofluorescence. Treatment with systemic prednisone, antibiotics, and antifungals, and with topical steroids and protection from ultraviolet radiation were unsuccessful, and the seal died during the second week of treatment. A pruritic allergic dermatitis with loss of guard hairs over the dorsum was described in a captive sub-Antarctic fur seal (Arctocephalus tropicalis) (Bodley et al., 1999). Diagnosis was based on positive reactions to allergens prepared from weed, grass, tree pollens, and some insects. Symptomatic treatment with oral antihistamines was only partially successful, but specific allergen immunotherapy using ten allergens was effective, despite side effects. A skin condition characterized by hyperkeratosis, alopecia, and ulceration has been well described in northern elephant seals, but its etiology remains obscure (Figure 1G) (Beckmen et al., 1997). Another ulcerative skin disease of obscure etiology has been described by Anderson et al. (1974) in gray seals.
Musculoskeletal System Numerous bacteria can cause deep abscesses, myositis, osteomyelitis, and arthritis (see Chapter 16, Bacterial Diseases) (Thornton et al., 1998). Most of these are opportunistic, occurring following trauma, introduction through contaminated hypodermic needles or surgical instruments, or hematogenous spread as a result of generalized sepsis. Clostridium perfringens has
0839_frame_C41 Page 916 Tuesday, May 22, 2001 2:07 PM
916
CRC Handbook of Marine Mammal Medicine
been isolated from cases of severe myositis following poor injection technique (Greenwood and Taylor, 1978). Trauma is especially common in free-ranging, stranded pinnipeds (see Chapter 23, Noninfectious Diseases). Osteomyelitis affecting the extremities is common as a result of trauma (Thurman et al., 1982) or secondary to more superficial infections such as calicivirus (see above). Some neoplastic diseases can manifest clinical signs in the musculoskeletal system. Carcinomas of possible transitional cell origin may erode the lumbar spine, affecting neurological function as well as potentially resulting in pathological fractures. Animals may exhibit hindend paresis or paralysis (Gulland et al., 1996a). Lymphosarcoma has also been seen to affect the bone marrow in a harbor seal (Stroud and Stevens, 1980). Parasites, including the filariid Acanthocheilonema odenhali, the protozoan Sarcocystis, and inactive Uncinaria larvae may be found in muscles and fascia, but do not usually cause clinical signs of disease. Physical examination, fine-needle aspiration with cytology and culture of the aspirate, radiographs, computerized tomography, and ultrasound (see Chapters 24 through 28) will all facilitate diagnosis of musculoskeletal problems. Treatment is dictated by the diagnosis. Examples of fracture repair are given in Bennett et al. (1994).
Digestive System Specific etiologies of gastrointestinal disease include both infectious and noninfectious agents. An adenovirus was reported as a cause of viral hepatitis in California sea lions (Dierauf et al., 1981). Clinical signs of affected animals included diarrhea, anorexia, abdominal pain, posterior paresis, polydipsia, and photophobia. Post-mortem findings included hepatic necrosis and intranuclear inclusions in Kupffer’s cells. Other infectious causes of hepatitis are bacteria, and several types have been isolated from the livers of pinnipeds (Thornton et al., 1998). Mycotic agents including Coccidioides immitis have also affected the liver of California sea lions (Fauquier et al., 1996). Neoplastic diseases may manifest themselves in the liver of pinnipeds (Stroud and Stevens, 1980; Gulland et al., 1996a). Hemochromatosis was recently observed in captive California sea lions and northern fur seals, but the etiology is still obscure (Garcia et al., 2000). Several bacteria have been implicated as causing enteritis in different pinnipeds, including Clostridium spp. and Salmonella spp. The interpretation of culture of these organisms from fecal samples is difficult, as they have been cultured from both clinically normal animals and those with severe hemorrhagic enteritis. Although most gastrointestinal parasites are part of the normal flora of free-ranging pinnipeds and do not significantly affect the host, there are some instances where they can be responsible for causing clinical signs and even mortality. Acanthocephalans have caused gastrointestinal perforation and peritonitis in gray and harbor seals. Gastric nematodes may potentiate malnutrition in already compromised animals, especially young animals, and high burdens may cause increased gastric ulceration and chronic emesis. Gastric nematodes have also caused duodenal perforations leading to peritonitis and death in stranded California sea lions (Fletcher et al., 1998). High cestode burdens may also obstruct the intestinal lumen of nutritionally challenged young pinnipeds (Gulland, unpubl. obs.). Small amounts of rectal bleeding, irregular in frequency and of varying duration, in harbor seals and California sea lions have been associated with a post-mortem finding of ileo-cecocolic intussusception (Lair and Lamberski, pers. comm.). Congenital abnormalities, including cleft palate (Suzuki et al., 1992) and hiatial herniation, have been seen in stranded pinnipeds. Ingestion of foreign bodies occurs in both free-living and captive pinnipeds (see Chapter 23, Noninfectious Diseases).
0839_frame_C41 Page 917 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
917
Iatrogenic causes of gastrointestinal disease include feeding inappropriate formulas or spoiled fish, using poorly designed feeding tubes, or feeding at an inappropriate rate or volume. Young animals, especially when debilitated, often go through a period of regurgitation if formula is introduced too quickly. Adequate rehydration and a gradual introduction to complex diets may aid in decreasing the frequency of emesis. Impactions caused by solidifying formula in neonates has been seen and may also be prevented by feeding appropriate formulas (see Chapter 37, Hand-Rearing), monitoring hydration, and a gradual introduction to complex diets. The clinical signs associated with diseases of the digestive tract in pinnipeds include inappetence, emesis, regurgitation, icterus, melena, diarrhea, and steatorrhea. Abdominal pain or discomfort is often manifested as inappetence, lethargy, or depression. Otariids with abdominal discomfort will often tuck their flippers to their abdomen. In the water, they may float with tucked flippers and a hunched back. Physical examination may help to detect and differentiate broken, missing, or worn teeth, oral ulcers, oral foreign bodies such as fish spines and fishhooks, abdominal distension, abdominal masses, a palpable fluid wave, perineal swelling, or prolapsed rectum. A complete blood count may identify an infectious cause. Clinical chemistry findings may indicate specific organ involvement, hypoproteinemia, gastrointestinal hemorrhaging, and electrolyte imbalances associated with chronic emesis or diarrhea (see Chapter 19, Clinical Pathology). Abdominocentesis of a distended abdomen can differentiate peritonitis and hemoperitoneum. As many pinnipeds have large intra-abdominal vessels and large spleens, aspiration of frank blood does not necessarily indicate hemoperitoneum. However, a case of hemoperitoneum and a gastric intramural hematoma was recently described in a captive northern fur seal (Frasca et al., 2000). Further diagnostics may include radiographs to detect gastric foreign bodies, gastric impaction, or constipation. Although it is often difficult to achieve good contrast in pinniped abdominal radiographs because of the blubber layer, in young and thin animals it is possible to achieve some indication of organ size, displacement, and intra-abdominal masses. Ultrasound may be more helpful in detecting ascites, ileus, or organ abnormalities. Endoscopy has been beneficial in diagnosing gastric ulcers, gastritis, colitis, and gastric foreign bodies, as well as in obtaining gastric and colonic biopsies. Laparoscopic examination can enable direct visualization of the gastrointestinal serosa as well as the liver, pancreas, and associated structures. Biopsies of the liver may be obtained either laparoscopically or by ultrasoundguided biopsy (see Chapters 24 through 28, Diagnostic Imaging). Supportive therapy of gastrointestinal disease is important, because fluid, electrolyte, and protein abnormalities can quickly result in mortality if they are not resolved. Since many animals with gastrointestinal disease vomit, fluids, medications, and potentially even nutrition must be provided parenterally (see above).
Respiratory System Phocine distemper virus (PDV) and canine distemper virus (CDV) have caused epizootics of pneumonia and death in harbor and Baikal seals (Phoca sibirica), respectively (see Chapter 15, Viral Diseases, and reviews by Kennedy, 1998, and Duignan, 1999). Ocular and nasal discharges, cough, cyanosis of mucous membranes, dyspnea, diarrhea, fever, and central nervous signs are observed in affected seals. Subcutaneous emphysema of the neck and thorax occurs as a sequel to pulmonary damage, and seals have difficulty swimming and diving (Krogsrud et al., 1990). Diagnosis is based on detection of eosinophilic intranuclear and intracytoplasmic inclusions in epithelial cells, macrophages, and syncytia, immunohistochemistry, electron microscopy, demonstration of virus in tissue using polymerase chain reaction, and increasing antibody titers in serum. Virus isolation is difficult, yet necessary to confirm identification of the virus. Treatment is directed at supportive care, and controlling secondary bacterial infections that
0839_frame_C41 Page 918 Tuesday, May 22, 2001 2:07 PM
918
CRC Handbook of Marine Mammal Medicine
commonly cause death in infected seals (Baker and Ross, 1992). Antibiotics effective against Bordetella bronchiseptica, Corynebacterium spp., and Streptococcus spp. are recommended. Although clinical recovery is documented, CDV has been isolated from asymptomatic carriers (Lyons et al., 1993). No commercially available vaccine for PDV currently exists, but commercially available attenuated CDV vaccine has been used to immunize stranded gray and harbor seals (Carter et al., 1992). Experimental inoculation of harbor seals with inactivated and subunit CDV vaccines has provided some protection from clinical disease (Visser et al., 1989; 1992; Van Bressem et al., 1991). Influenza virus has also caused epizootics in harbor seals, with clinical signs similar to those in seals with PDV and CDV (Geraci et al., 1982). These included dyspnea, lethargy, bloodstained nasal discharge, and subcutaneous emphysema, with pneumonia as the predominant post-mortem lesion. Phocine herpesvirus-1 (PhHV-1) has caused pneumonia in neonatal harbor seals in rehabilitation (Borst et al., 1986), while another herpesvirus was isolated from a California sea lion with acute hemorrhagic pneumonia (Kennedy-Stoskopf et al., 1986). Diagnosis of both infections is based on viral isolation, and treatment is supportive. Harbor seals with pneumonia associated with influenza virus were also infected with a mycoplasma, so therapy with antibiotics such as tetracyclines may be beneficial (Geraci et al., 1984). Bacterial pneumonias are common in seals and sea lions, both as primary infections and secondary to viral and lungworm infections. A variety of organisms may be involved, although Gram-negative organisms are most common (Keyes et al., 1968; Sweeney, 1986a; Spraker et al., 1995; Thornton et al., 1998). Clinical signs include tachypnea, dyspnea, lethargy, and cough. Diagnosis is based upon auscultation of the chest, radiography of the lung fields, and bronchoscopy. Treatment with the appropriate systemic antibiotic may be based upon prediction of the likely organism, or culture and sensitivity of organisms from tracheal or bronchial washes (Johnson et al., 1998). Oral mucolytics such as acetylcysteine and bronchodilators such as aminophylline have been used regularly on stranded harbor seals and California sea lions in rehabilitation. Pneumonia in otariids may occur with heavy infestation of Parafilaroides decorus, although asymptomatic infection is common in young animals. Parafilaroides gymnurus infects alveoli of phocids, and Otostrongylus circumlitus may cause obstructive bronchitis and bronchiolitis in harbor and ringed seals, and yearling northern elephant seals (see Chapter 18, Parasitic Diseases). The degree of inflammatory response to Parafilaroides infections varies from none to marked suppurative and granulomatous pneumonia. Reaction may be more severe to dead and degenerate worms. Diagnosis depends upon detection of larvae in feces or sputum. Treatment with fenbendazole or ivermectin removes infection, but in severe cases simultaneous treatment with antibiotics and corticosteroids is recommended to control secondary bacterial infections and reduce the inflammatory response to parasites. Interestingly, a Brucella spp. isolate was recently obtained from the lung of a harbor seal with Parafilaroides spp. infestation (Garner et al., 1997). Histological examination revealed most of the inflammation and Brucella to be around the dead parasites. The role of Parafilaroides in the epidemiology of Brucella infections remains unclear. Pulmonary granulomas due to infection with Mycobacterium tuberculosis have been reported in captive and wild pinnipeds (Cousins et al., 1990; Forshaw and Phelps, 1991; Bastida et al., 1999). An enzyme-linked immunosorbent assay (ELISA) test and intradermal tuberculin tests have been used for diagnosis of infection in live pinnipeds, although interpretation of results is difficult (Cousins, 1987; Needham and Phelps, 1990), and successful treatment of clinical cases has not been documented. Similar lesions may also result from infection with Coccidioides immitis and Cryptococcus neoformans. Differential diagnosis is usually made post-mortem based on histological detection of organisms and culture.
0839_frame_C41 Page 919 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
919
Cardiovascular Anemia is common in young otariids as a consequence of hookworm (Uncinaria spp.) infestation, or secondary to malnutrition. Affected animals are weak, occasionally dyspneic, and have pale mucous membranes. Diagnosis of hookworm infestation is based on detection of ova in feces (see Chapter 18, Parasitic Diseases), although animals may remain anemic for weeks after patent infection ceases. Treatment with anthelmintics and supplementation with iron and vitamin B12 is usually effective. Nonregenerative anemia is seen in California sea lions as a consequence of chronic renal damage usually as a result of leptospirosis (see below). Erythropoetin has been used to treat several California sea lions successfully, although the dose of choice is still unclear (Gulland and Haulena, unpubl. obs.). Disseminated intravascular coagulation (DIC), characterized by bleeding from the nares, hematoma formation, thrombocytopenia, hypofibrinogenemia, and extended clotting times is relatively common in stranded northern elephant seals (Gulland et al., 1996b). It may occur with septicemia or vasculitis associated with migrating Otostrongylus larvae (Gulland et al., 1997a). Diagnosis of Otostrongylus infestation in the live animal during the prepatent period, when young elephant seals are commonly clinically affected, is not currently possible, although clinical signs and season of occurrence are highly suggestive of infection. Treatment of DIC is rarely successful, but should be directed at removing the underlying cause, as well as supportive care. Cardiac insufficiency as a consequence of heartworm infestation has been documented in captive and wild pinnipeds (see Chapter 18, Parasitic Diseases). Infection by either the canine heartworm Dirofilaria immitis or the phocid parasite Acanthocheilonema spirocauda may cause dilatation of the pulmonary artery and right ventricle, and can be detected radiographically. Microfilaria observed in blood smears must be distinguished from those of A. odenhali (see Chapter 18, Parasitic Diseases). Successful treatment of documented cases has not been described. Preventive treatment of captive animals in D. immitis endemic regions with ivermectin at 0.6 mg/kg every month during the mosquito season is recommended, as well as removal of lice from stranded animals, as the seal louse Echinophthirius horridus has been shown to transmit A. spirocauda (Geraci et al., 1981).
Urogenital System The most common cause of renal disease in pinnipeds, especially California sea lions, is leptospirosis (Gulland, 1999). The infection is well recognized in California sea lions, but has also been reported in northern fur seals and harbor seals (Stamper et al., 1998; Stevens et al., 1999). Typical clinical signs include depression, anorexia, polydipsia, dehydration, vomiting, abdominal pain, and muscular tremors. Hematological changes include elevations in blood urea nitrogen, phosphorus, globulin, sodium, creatinine, and neutrophil count. Diagnosis is based on rising titers to Leptospira pomona by macroscopic slide agglutination test. Although single titers over 1:100 are considered positive in domestic animals, confirmed cases of L. pomona in sea lions usually have titers greater than 1:3200. Leptospires may also be demonstrated by immunofluorescence in urine and in kidneys of dead animals, and by Warthin–Starry, Steinert, or Levaditis stain in histological sections. Culture is difficult, but possible from blood and urine of live animals, and kidney and liver of dead ones. Treatment with tetracycline at 22 mg/kg every 8 hours, and potassium penicillin G (44,000 U/kg every 12 hours) for 10 days, combined with supportive therapy is usually effective. It is unknown whether or not surviving animals become carriers. Trials with hamsters infected with L. pomona have shown that infection was cleared by dihydrostreptomycin/penicillin G at 25 mg/kg, tylosin at 50 mg/kg, and erythromycin at 25 mg/kg, all for 3 days, or oxytetracycline at 40 mg/kg and ampicillin at 25 mg/kg
0839_frame_C41 Page 920 Tuesday, May 22, 2001 2:07 PM
920
CRC Handbook of Marine Mammal Medicine
for 5 days (Alt and Bolin, 1996). There have been no trials on marine mammals to determine effective dosages for clearing infection. Supportive care should include fluid therapy, oral antacids to reduce gastric ulceration, and phosphorus binders such as aluminum hydroxide to lower blood phosphorus levels. Renal disease may also occur as a consequence of renal calculi or congenital renal aplasia (see Chapter 23, Noninfectious Diseases). Diagnosis of these rare conditions requires radiography and ultrasound, and treatment has not been reported. Tumors of the urinary and reproductive tracts are common in California sea lions (see Chapter 23, Noninfectious Diseases). Clinical signs in these animals usually result from pressure on ureters and invasion of local organs. Diagnosis is based on abdominocentesis and cytology, radiology, ultrasound, and laparoscopic biopsy techniques. Treatment has not been attempted. Abortions and stillborn pups are frequently observed on pinniped rookeries. Leptospires (Gilmartin et al., 1976), herpesviruses (Dietz et al., 1989), caliciviruses (Smith and Boyt, 1990), Coxiella burnetii (Lapointe et al., 1999), and high levels of DDTs (Gilmartin et al., 1976) have all been reported in aborting pinnipeds or their placentae; yet, their relative roles in causing abortion are still unclear. Vaginal prolapse has been observed in California sea lions (Haulena and Gulland, unpubl. obs.) and Australian sea lions (Neophoca cinerea) (Read et al., 1982). Treatment of the latter by ovariohysterectomy was successful. Uterine torsions and ruptures have been observed in California sea lions with domoic acid intoxication, and were believed to be consequences of severe convulsions (Gulland, 2000).
Endocrine System Few primary endocrine disorders have been documented in pinnipeds. Both hyper- and hyponatremia are common in stranded animals (see Chapter 10, Endocrinology; Chapter 36, Nutrition), and may be consequences of inappropriate stress responses. Adrenal necrosis resulting from infection by a herpesvirus, PhHV-1, has been associated with severe electrolyte and glucose abnormalities in stranded neonatal harbor seals undergoing rehabilitation (Gulland et al., 1997b). Hypothyroidism has been suspected as attributing to obesity in captive California sea lions and seems to be responsive to treatment with exogenous thyroid hormone.
Eyes Pinniped eyes are characterized by a large globe, prominent tapetum lucidum, rounded lens, and a slitlike pupil (Wartzok and Ketten, 1999). The pinniped retina is predominantly composed of rods, and most pinnipeds have good vision below the surface of the water in low light and above the surface in bright light (Wartzok and Ketten, 1999). Pinnipeds have very active lacrimal glands producing constant tearing that may protect the cornea. Lack of tearing is often used as an indication of dehydration. Eye lesions appear to be common in both free-ranging (Stoskopf et al., 1985; Schoon and Schoon, 1992) and captive pinnipeds (Hirst et al., 1983). There may be an increased frequency of eye lesions in animals that are maintained in freshwater environments (Sweeney, 1986b; Dunn et al., 1996). However, as some species of free-ranging phocids live entirely in fresh water, it is likely that the etiology of eye lesions in captive pinnipeds is multifactorial. By-products of chlorine disinfection (chloramines and other oxydizing agents), opportunistic pathogens, microtrauma, ultraviolet hypersensitivity, and pH imbalance may be important factors (Geraci, 1986). Animals that develop eye lesions in captive environments may develop corneal opacities, edema, erosions, ulcers, uveitis, and cataracts (Hirst et al., 1983). Cataracts frequently lead to synechiae formation, anterior prolapse, and rupture of the globe. Traumatic lesions, either
0839_frame_C41 Page 921 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
921
anthropogenic or from conspecifics, are common in free-ranging animals. Secondary bacterial infections may be responsible for exacerbating the lesions. Various bacteria have been cultured from traumatic lesions, conjunctivitis, and keratitis in pinnipeds (Thornton et al., 1998). Visually impaired pinnipeds will thrust their vibrissae forward if investigating noises or new surroundings. Although normal pinnipeds will also do this, visually impaired animals tend to exaggerate the action and extend their vibrissae for prolonged periods of time. The menace response is difficult to evaluate, as the vibrissae are very sensitive to movement (Dutton, 1991). Visually impaired animals may not avoid obstacles well if placed into a new surrounding, but accommodate very quickly using tactile and acoustic cues, thus making diagnosis of blindness difficult. Ophthalmic examination is difficult in pinnipeds because of a prominent nictitating membrane, strong eyelids, and the ability to retract the globe into the ocular cavity (Hirst et al., 1983). Very narrow pupils limit visualization of internal eye structures such as the lens and retina, and pinnipeds do not tend to dilate their pupils very well when topical mydriatic agents are applied to the cornea (Dutton, 1991). Intraocular injection of epinephrine has been required to dilate the pupil sufficiently for such procedures as cataract removal by phaecoemulsification. Cataracts have also been removed by incising 180° of the cornea, discission, and extracapsular removal of the lens (Dutton, 1991). Treatment of ocular lesions of pinnipeds is similar to that of domestic animals. However, the use of saline washes, most readily given as saltwater baths or in spray bottles, appears to help decrease corneal edema. Analgesics (such as aspirin or flunixin meglumine) may be necessary to reduce pain.
Nervous System Encephalitis has been attributed to morbilliviruses, herpesviruses, rabies, bacteria, fungi, Toxoplasma gondii, and Sarcocystis neurona infections (see Chapters 15 through 18, Infectious Diseases). Clinical signs are nonspecific and include depression, muscular tremors, ataxia, seizures, and occasionally loss of pupillary reflex. There may be differences in likelihood of different diseases in different age classes of animals: Sarcocystis neurona–associated encephalitis was the most common cause of death in adult harbor seals stranded along the central California coast from 1980 to 1998 (LaPointe et al., 1998), and domoic acid intoxication of California sea lions in the same region occurred in adults but not in yearlings (Gulland, 2000). Morbilliviruses and protozoal infections may be diagnosed on the basis of rising antibody titers, although confirmation is usually made post-mortem on histological examination of brain tissue, using immunoperoxidase and immunofluorescent techniques. Treatment is supportive, although clindamycin has been used on a Hawaiian monk seal (Monachus schauinslandi) showing neurological signs with a rising antibody titer to T. gondii (Braun, pers. comm.). The animal recovered. Neuronal necrosis in the hippocampus of California sea lions is caused by domoic acid exposure (Scholin et al., 2000) (see Chapter 22, Toxicology). Neurological signs include seizures, tremors, and ataxia. Diagnosis depends on detection of domoic acid in serum, urine, or feces of affected animals, coupled with detection of Pseudonitzschia australis in the environment and prey of affected sea lions. As domoic acid is water-soluble and rapidly excreted in urine following ingestion, urine is the most useful fluid for diagnostic purposes. Intracranial space-occupying lesions, usually tumors, have caused seizures in captive sea lions. Although hydrocephalus occurs in young stranded elephant seals (Trupkiewicz et al., 1997), sudden death usually occurs, rather than neurological signs. Electrolyte imbalances associated with renal disease and/or nutritional deficiencies may also cause neurological signs (see Chapter 36, Nutrition; Chapter 10, Endocrinology).
0839_frame_C41 Page 922 Tuesday, May 22, 2001 2:07 PM
922
CRC Handbook of Marine Mammal Medicine
Thiamine deficiency and hyponatremia are diagnosed by a combination of history of lack of supplementation, plasma biochemistry, and response to treatment with sodium chloride or thiamine (Geraci, 1972). Treatment of encephalitides is essentially supportive, although appropriate antibiotic, antifungal, and antiprotozoal drugs are recommended for infectious causes. Control of seizures with diazepam, lorazepam (longer antiseizure effect), and phenobarbitone is beneficial, and can facilitate administration of fluids for rehydration. Successful control of idiopathic seizures in a captive adult California sea lion has been achieved with 1 mg/kg SID oral phenobarbitone (Gage, pers. comm.).
Acknowledgments The authors thank Laurie Gage, Tom Reidarson, Shawn Johnson, Claudia Grili, James Stanton, and Larry Dunn for reviewing this chapter; Karina Acevedo, Bob Braun, Stéfanie Lair, Nadine Lamberski, and Terry Spraker for personal communications; and the animals, volunteers, and staff of The Marine Mammal Center, Sausalito, California, for teaching us all they know.
References Alt, D.P., and Bolin, C., 1996, Preliminary evaluation of antimicrobial agents for treatment of Leptospira interrogans serovar pomona infection in hamsters and swine, Am. J. Vet. Res., 57: 59–62. Anderson, S.S., Bonner, W.N., Baker, J.R., and Richards, R., 1974, Grey seals, Halichoerus grypus, of the Dee Estuary and observations on a characteristic skin lesion in British seals, J. Zool., 174: 429–440. Baker, J.R., and Ross, H.M., 1992, The role of bacteria in phocine distemper, Sci. Total Environ., 115: 9–14. Bastida, R., Loureiro, J., Quse, V., Bernardelli, A., Rodriguez, D., and Costa, E., 1999, Tuberculosis in a wild subantarctic fur seal from Argentina, J. Wildl. Dis., 35: 796–798. Beckmen, K.B., Lowenstine, L.J., Newman, J., Hill, J., Hanni, K., and Gerber, J., 1997, Clinical and pathological characterization of northern elephant seal skin disease, J. Wildl. Dis., 33: 438–449. Bennett, R.A., Dunker, F.H., and Gage, L., 1994, Subtotal radial ostectomy in a California sea lion, in Proceedings of the American Association of Zoo Veterinarians, Pittsburg, PA, October 22–27, 144–145. Bodley, K., Monaghan, C., and Mueller, R.S., 1999, Treatment of allergic dermatitis (atopy) in a subantarctic fur seal (Arctocephalus tropicalis) using immunotherapy, in Proceedings of the American Association of Zoo Veterinarians, Columbus, OH., 129–130. Borkowski, R., Moore, P.A., Mumford, S., Patchett, K., Rice, J., and Rubinstein, B., 1999, Extended use of subpalpebral lavage systems for treatment of keratitis in a harbor seal (Phoca vitulina), in Proceedings of the International Association for Aquatic Animal Medicine, 30: 39–40. Borst, G.H.A., Walvoort, H.C., Reijnders, P.J.H., van der Kamp, J.S., and Osterhaus, A.D.M.E., 1986, An outbreak of herpesvirus infection in harbour seals (Phoca vitulina), J. Wildl. Dis., 22: 1–6. Burns, R., 1993, Cutaneous lupus in a grey seal (Halichoerus grypus), in Proceedings of the American Association of Zoo Veterinarians, St. Louis, MO, 141. Carter, S.D., Hughes, D.E., Taylor, V.J., and Bell, S.C., 1992, Immune responses in common and grey seals during the seal epizootic, Sci. Total Environ., 115: 83–91. Cornell, L., 1986, Capture, transportation, restraint, and marking, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 764–770. Cousins, D.V., 1987, ELISA for detection of tuberculosis in seals, Vet. Rec., 121: 305. Cousins, D.V., Francis, B.R., Gow, B.L., Collins, D.M., McGlashan, C.H., Gregory A., and Mackenzie, R.M., 1990, Tuberculosis in captive seals: Bacteriologic studies on an isolate belonging to the Mycobacterium tuberculosis complex, Res. Vet. Sci., 48: 196–200.
0839_frame_C41 Page 923 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
923
Dierauf, L.A., Lowenstine, L.J., and Jerome, C., 1981, Viral hepatitis (adenovirus) in a California sea lion, J. Am. Vet. Med. Assoc., 179: 1194–1197. Dietz, R., Heide-Jorgensen, J., and Harkonen, T., 1989, Mass death of harbour seals (Phoca vitulina) in Europe, Ambio, 18: 258–264. Duignan, P.J., 1999, Morbillivirus infections of marine mammals, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 497–501. Dunn, J.L., Abt, D.A., Overstrom, N.A., and St. Aubin, D.J., 1996, An epidemiologic survey to determine risk factors associated with corneal and lenticular lesions in captive harbor seals and California sea lions, in International Association for Aquatic Animal Medicine 27th Annual Conference Proceedings, Chattanooga, TN, May 11–15, 100–102. Dutton, A.G., 1991, Cataract removal in a fur seal, J. Am. Vet. Med. Assoc., 198: 309–311. Fauquier, D.A., Gulland, F.M.D., Trupkiewicz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996, Coccidioidomycosis in free-living California sea lions (Zalophus californianus) in central California, J. Wildl. Dis., 32: 707–710. Fletcher, D., Gulland, F.M.D., Haulena, M., Lowenstine, L.J., and Dailey, M., 1998, Nematodeassociated gastrointestinal perforations in stranded California sea lions (Zalophus californianus), International Association for Aquatic Animal Medicine 29th Annual Conference Proceedings, San Diego, CA, 59. Forshaw, D., and Phelps, G.R., 1991, Tuberculosis in a captive colony of pinnipeds, J. Wildl. Dis., 27: 288–295. Frasca, S., Van Kruiningen, H., Dunn, J.L., and St. Aubin, D.J., 2000, Gastric intramural hematoma and hemoperitoneum in a captive northern fur seal, J. Wildl. Dis., 36: 565–569. Gage, L.J., Amaya-Sherman, L., Roletto, J., and Bently, S., 1990, Clinical signs of San Miguel sea lion virus in debilitated California sea lions, J. Zoo Wildl. Med., 21: 79–83. Garcia, A.R., Montali, R.J., Dunn, J.L., Torres, N.L., Centeno, J.A., and Goodman, K., 2000, Hemochromatosis in captive otariids, in Proceedings of the Joint Conference of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine, New Orleans, LA, September 17–21, 197. Garner, M.M., Lambourn, D.M., Jeffries, S.J., Hall, B.P., 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. Gentry, R.L., and Casanas, V.R., 1997, A new method for immobilizing otariid neonates, Mar. Mammal Sci., 13: 155–157. Gentry, R.L., and Holt, J.R., 1982, Equipment and techniques for handling northern fur seals, U.S. Department of Commerce, NOAA Technical Report NMFS SSRF-758, 15 pp. Geraci, J.R., 1972, Experimental thiamine deficiency in captive harp seals, Phoca groenlandica, induced by eating herring, Clupea harengus, and smelts, Osmerus mordax, Can. J. Zool., 50: 179–195. Geraci, J.R., 1981, Dietary disorders in marine mammals: Synthesis and new findings, J. Am. Vet. Med. Assoc., 179: 1183–1191. Geraci, J.R., 1986, Husbandry, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 757–760. Geraci, J.R., Fortin, J.F., St. Aubin, D.J., and Hicks, B.D., 1981, The seal louse, Echinophthirius horridus: An intermediate host of the seal heartworm, Dipetalonema spirocauda (Nematoda), Can. J. Zool., 59: 1457–1459. Geraci, J.R., St. Aubin, D.J., Barker, I.K., Webster, R.G., Hinshaw, V.S., Bean, W.J., Ruhnke, H.L., Prescott, J.H., Early, G., Baker, A.S., Madoff, S., and Schooley, R.T., 1982, Mass mortality of harbor seals: Pneumonia associated with influenza A virus, Science, 215: 1129–1131. Geraci, J.R., St. Aubin, D.J., Barker, I.K., Hinshaw, V.S., Webster, R.G., and Ruhnke, H.L., 1984, Susceptibility of gray (Halichoerus grypus) and harp (Phoca groenlandica) seals to the influenza virus and mycoplasma of epizootic pneumonia of harbour seals (Phoca vitulina), Can. J. Fish. Aquat. Sci., 41: 151–156.
0839_frame_C41 Page 924 Tuesday, May 22, 2001 2:07 PM
924
CRC Handbook of Marine Mammal Medicine
Gilmartin, W.G., DeLong, R.L., Smith A.W., Sweeney, J.C., De Lappe, B.W., Risebrough, R.W., Griner, L.A, Dailey, M.D., and Peakall, D.B., 1976, Premature parturition of the California sea lion, J. Wildl. Dis., 12: 104–115. Goldstein, T., Johnson, S.P., Werner, L.J., Nolan, S., and Hilliard, B.A., 1998, Causes of erroneous white blood cell counts and differentials in clinically healthy young northern elephant seals, Mirounga angustirostris, J. Zoo Wildl. Med., 29: 408–412. Goldstein, T., Johnson, S.P., Phillips, A.V., Hanni, K., 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. Greenwood, A.G., and Taylor, D.C., 1978, Clostridial myositis in marine mammals, Vet. Rec., 103: 54–55. Gulland, F.M.D., 1999, Leptospirosis in marine mammals, in Zoo and Wild Animal Medicine: Current Therapy 4, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 469–471. Gulland, F.M.D., 2000, Domoic acid toxicity in California sea lions (Zalophus californianus) stranded along the central California Coast, May–October 1998, NOAA Technical Memorandum, NMFSOPR, 44 pp. Gulland, F.M.D.., Trupkiewicz, J.G., Spraker, T.R., and Lowenstine, L.J., 1996a, Metastatic carcinoma of probable transitional cell origin in 66 free-living California sea lions (Zalophus californianus), 1979–1994, J. Wildl. Dis., 32: 250–258. Gulland, F.M.D, Werner, L., O’Neill, S., Lowenstine, L.J., Trupkiewicz, J., Smith, D., Royal, B., and Strubel, I., 1996b, Baseline coagulation assay values for northern elephant seals (Mirounga angustirostris), and disseminated intravascular coagulation in this species, J. Wildl. Dis., 32: 536–540. Gulland, F.M.D., Beckmen, K., Burek, K., Lowenstine, L., Werner, L., Spraker, T., and Harris, E., 1997a, Otostrongylus circumlitus infestation of northern elephant seals (Mirounga angustirostris) stranded in central California, Mar. Mammal Sci., 13: 446–459. Gulland, F.M.D., Lowenstine, L.J., Lapointe, J.M., Spraker, T., and King, D.P., 1997b, Herpesvirus infection in stranded Pacific harbor seals of coastal California, J. Wildl. Dis., 33: 450–458. Gutter, A.E., Wells, S.K., and Spraker, T.R., 1987, Generalized mycobacteriosis in a California sea lion (Zalophus californianus), J. Zoo Anim. Med., 18: 118–120. Hastings, B.E., Lowenstine, L.J., Gage, L.J., and Munn, R.J., 1989, An epizootic of seal pox in pinnipeds at a rehabilitation center, J. Zoo Wildl. Med., 20: 282–290. Hirst, L.W., Stoskopf, M.K., Graham, D., Green, W.R., and Dickson, J.B., 1983, Pathologic findings in the anterior segment of the pinniped eye, J. Am. Vet. Med. Assoc., 183: 1226–1231. 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. Kennedy, S., 1998, Morbillivirus infections in aquatic mammals, J. Comp. Pathol., 119: 201–225. Kennedy-Stoskopf, S., Stoskopf, M.K., Eckhaus, M.A., and Strandberg, J.D., 1986, Isolation of a retrovirus and a herpesvirus from a captive California sea lion, J. Wildl. Dis., 22: 156–164. Keyes, M.C., Crews, F.W., and Ross, A.J., 1968, Pasturella multocida isolated from a California sea lion (Zalophus californianus), J. Am. Vet. Med. Assoc., 153: 803–804. Klontz, K.C., Mullen, R.C., Corbyons, T.M., and Barnard, W.P., 1993, Vibrio wound infections in humans following shark attack, J. Wilderness Med., 4: 68–72. Krogsrud, J., Evensen, Ø., Hølt, G., and Markussen, N.H., 1990, Seal distemper in Norway in 1988 and 1999, Vet. Rec., 126: 460–461. Lapointe, J.-M., Duignan, P.J., Marsh, A.E., Gulland, F.M., Barr, B.C., Naydan, D.K., King, D.P., Farman, C.A., Huntington, K.A., and Lowenstine, L.J., 1998, Meningoencephalitis due to a Sarcocystis neurona-like protozoan in Pacific harbor seals (Phoca vitulina richardsi), J. Parasitol., 84: 1184–1189. Lapointe, J.-M., Gulland, F.M., Haines, D.M., Barr, B.C., and Duignan, P.J., 1999, Placentitis due to Coxiella burnetii in Pacific harbor seal (Phoca vitulina richardsi), J. Vet. Diagn. Invest., 11: 541–543.
0839_frame_C41 Page 925 Tuesday, May 22, 2001 2:07 PM
Seals and Sea Lions
925
Lyons, C., Welsh, M.J., Thorsen, J., Ronald, K., and Rima, B.K., 1993, Canine distemper virus isolated from a captive seal, Vet. Rec., 132: 487–488. Mo, G., Gili, C., and Ferrando, P., 2000, Do photoperiod and temperature influence the molt cycle of Phoca vitulina in captivity? Mar. Mammal Sci., 16: 570–578. Needham, D.J., and Phelps, G.R., 1990, Interpretation of tuberculin tests in pinnipeds, in Proceedings of the American Association of Zoo Veterinarians, South Padre Island, TX, 115. Pavia, A.T., Bryan, J.A., Maher, K.L., Hester, T.R., Jr., and Farmer III, J.J., 1989, Vibrio carchariae infection after shark bite, Ann. Intern. Med., 111: 85–86. Read, R.A., Reynolds, W.T., Griffiths, D.J., and Reilly, J.S., 1982, Vaginal prolapse in a south Australian sea lion (Neophoca nove hollandia), Aus. Vet. J., 58: 269–271. Reynolds, J.E., and Rommel, S.A. (Eds.), 1999, Biology of Marine Mammals, Smithsonian Institution Press, Washington, D.C., 578 pp. Riedman, M., 1990, The Pinnipeds. Seals, Sea Lions, and Walruses, University of California Press, Berkeley, 439 pp. 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. Schoon, H.A., and Schoon, D., 1992, Lenticular lesions in harbour seals (Phoca vitulina), J. Comp. Pathol., 107: 379–388. Smith, A.W., and Boyt, P.M., 1990, Caliciviruses of ocean origin: A review, J. Zoo Wildl. Med., 21: 3–23. Spraker, T.R., Bradley, D., Antonelis, G., DeLong, R., and Calkins, D., 1995, Fibrinous pneumonia of neonatal pinnipeds associated with β-hemolytic E. coli, in Proceedings of the American Association of Zoo Veterinarians/American Association of Wildlife Veterinarians, August 12–17, East Lansing, MI, 504. Stamper, M.A., Gulland, F.M.D., and Spraker, T., 1998, Leptospirosis in rehabilitated Pacific harbor seals from California, J. Wildl. Dis., 34: 407–410. Stevens, E., Lipscomb, T.P., and Gulland, F.M.D., 1999, An additional case of leptospirosis in a harbor seal, J. Wildl. Dis., 35: 150. Stoskopf, M.K., Zimmerman, S., Hirst, L.W., and Green, R., 1985, Ocular anterior segment disease in northern fur seals, J. Am. Vet. Med. Assoc., 187: 1141–1144. Stoskopf, M.K., Moench, T., Thoen, C., and Charache, P., 1987, Tuberculosis in pinnipeds, in Proceedings of the American Association of Zoo Veterinarians, September 6–11, Oahu, HI, 393. Stroud, R.K., and Stevens, D.R., 1980, Lymphosarcoma in a harbor seal (Phoca vitulina richardsi), J. Wildl. Dis., 16: 267–270. Suzuki, M., Kishimoto, M., Hayama, S., Ohtaishi, N., and Nakane, F., 1992, A case of cleft palate in a Kuril seal (Phoca vitulina stejnegeri), from Hokkaido, Japan, J. Wildl. Dis., 28: 490–493. Sweeney, J., 1986a, Infectious diseases, in Zoo and Wild Animal Medicine, 2nd ed., Fowler, M.E. (Ed.), W.B. Saunders, Philadelphia, 777–781. Sweeney, J., 1986b, 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. Thornton, S.M., Nolan, S., and Gulland, F.M.D., 1998, Bacterial isolates from California sea lions (Zalophus californianus), harbor seals (Phoca vitulina), and northern elephant seals (Mirounga angustirostris) admitted to a rehabilitation center along the central California coast, 1994–1995, J. Zoo Wildl. Med., 29: 171–176. Thurman, G.D., Downes, S.J., and Barrow, S., 1982, Anaesthetization of a Cape fur seal (Arctocephalus pusillus) for the treatment of a chronic eye infection and amputation of a metatarsal bone, J. S. Afr. Vet. Assoc., 53: 255–257. Trupkiewicz, J.G., Gulland, F.M.D., and Lowenstine, L.J., 1997, Congenital defects in northern elephant seals stranded along the central California coast, J. Wildl. Dis., 33: 220–225.
0839_frame_C41 Page 926 Tuesday, May 22, 2001 2:07 PM
926
CRC Handbook of Marine Mammal Medicine
Van Bonn, W., Jensen, E.D., House, C., House, J.A., Burrage, T., and Gregg, D.A., 2000, Epizootic vesicular disease in captive California sea lions, J. Wildl. Dis., 36: 500–507. Van Bressem, M.F., De Meurichy, J., Chappuis, G., Spehner, D., Kieny, M.P., and Pastoret, P.P., 1991, Attempt to vaccinate orally harbour seals against phocid distemper, Vet. Rec., 129: 362. Visser, I.K.G., van de Bildt, M.W.G., Brugge, H.N., Reijnders, P.J.H., Vedder, E.J., Kuiper, J., de Vries, P., Groen, J., Walvoort, H.C., Uyt de Haag, F.G.C.M., and Osterhaus, A.D.M.E., 1989, Vaccination of harbour seals (Phoca vitulina) against phocid distemper with two different inactivated canine distemper virus vaccines, Vaccine, 7: 521–526. Visser, I.K.G., Vedder, E.J., and van de Bildt, M.W.G., Orvell, C., Barrett, T., Osterhaus, A.D.M.E., 1992, Canine distemper virus ISCOMS induce protection in harbour seals (Phoca vitulina) against phocid distemper but still allow subsequent infection with phocid distemper virus-1, Vaccine, 10: 435–438. Wartzok, D., and Ketten, D.R., 1999, Marine mammal sensory systems, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 117–175.
0839_frame_C42 Page 927 Tuesday, May 22, 2001 2:08 PM
42 Walruses Michael T. Walsh, Brad F. Andrews, and Jim Antrim
Introduction As with all wild animals, to understand the medicine and husbandry of the walrus, one must understand its biology and ecology. Fay (1982) reviewed the ecology and biology of the Pacific walrus (Odobenus rosmarus), and additional reports describe its physical and biological characteristics (Brooks, 1954; Mansfield, 1958; Reeves, 1978; Ridgway and Harrison, 1981). Relatively few walruses are maintained in display facilities. In the past, walruses on display at zoological parks and aquaria were obtained as young orphans from the Arctic. During the spring migration, a number of calves are orphaned by accidents, weather, and hunting, and the majority of these die as a result of starvation and predation. A small number are recovered and sent to zoological facilities (Samansky and Sieswerda, 1995). Veterinary care has focused around hand-rearing (see Chapter 37) and treatment for tusk and gastrointestinal problems usually associated with ingestion of foreign bodies.
Biology There is only one living walrus species, which is divided into approximately six major populations in the Arctic. Fay describes four of these locations as containing “Atlantic” walrus (subspecies rosmarus); one group in the Bering and Chukchi Seas, as the “Pacific” walrus (O. r. divergens); and a sixth population in the Laptev Sea as a possible third subspecies (O. r. laptevi), although the relationship of this third group as a separate subspecies is undecided. The Pacific walrus tends to be a larger animal with longer tusks and a smaller nose compared with the Atlantic animal. While body weights of wild animals are difficult to obtain, weights of male walruses maintained in captivity at Marineland of the Pacific reached 1900 kg (4180 lb). Mature male walruses at SeaWorld are now weighed weekly and average 1350 to 1500 kg (2970 to 3300 lb). The skin of the walrus is thick, with neck and shoulder areas achieving maximum thickness of up to 5 cm (2 in.) in some males. In histological section, the epidermis of the walrus is about 1 cm thick with sebaceous and apocrine glands associated with each hair. Larger sweat glands are present in the area of the snout, possibly involved with scent production and recognition. Adult walruses in the wild generally molt in May and June, with variation between animals. Fay (1982) noted that animals maintained in the temperate zone begin to molt earlier, suggesting a thermal or photoperiod influence.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
927
0839_frame_C42 Page 928 Tuesday, May 22, 2001 2:08 PM
928
CRC Handbook of Marine Mammal Medicine
Cobb (1933) described and revised the information relating to early work on dentition. This was refined by Fay with the formula for complete secondary dentition at the time of birth as follows: 1 to 3 1 1 to 4 I -------------- C -- PC -------------- × 2 ≅ 30 1 to 3 1 2 to 4
(1)
where I = incisors, C = canines, and PC = postcanines. This formula is variable between individuals, with resorption of some incisors. Additional variation in adults may occur from trauma or disease. Not including the tusks (canines), there are 16 adult teeth that are consistently present: 3 1 1 to 4 I -- C -- PC -------------- × 2 = 16 0 1 2 to 4
(2)
Fay (1982) discusses in detail the form, development, and abrasion of tusks. For a long time, it was popularly believed that tusks were used for digging up mollusks from the seafloor. Fay concluded from his observations of wear patterns in tusk specimens, together with abrasion patterns of facial vibrissae, that walruses probably root using the upper edge of the snout rather than digging with the tusks. He felt that the primary role of the tusks is more social, as they are utilized in dominance and threat displays. Other functions may include fighting, formation and maintenance of holes in the ice, and as anchors in the ice to prevent drift with the current. When multiple animals are maintained in zoological institutions, it is common for them to spar with their tusks, especially during the breeding season. Locomotion of walruses out of water varies with the size of the animal and the surface it is on. The walrus can pull itself along by alternate movements of its pectoral flippers (usually done by smaller walrus on slippery surfaces), or by pulling forward with both (pectoral) flippers simultaneously in a rowing motion (in larger walrus on rough surfaces). In addition, the walrus is able to stand on all fours and walk in an ungainly fashion to avoid abdominal contact on rough or uncomfortable surfaces.
Reproduction Male walruses begin to reach sexual maturity by 7 years of age, with most males becoming sexually mature by 10 (Fay, 1982). The months of February, March, and April are the rut period for male walruses. Walruses maintained at Marineland of the Pacific sired their first calves at 10 years of age. Testicular enlargement in the bulls was observed in March and April. Some females may ovulate as early as 4 to 6 years of age, with 100% of females becoming fertile by 10 years (Fay, 1982). The latter age correlates with the attainment of maximum size. Females that gave birth at Marineland of the Pacific did so at 11 years of age. Breeding in wild populations occurs primarily from January to March. There is likely a delayed implantation until May to July (see Chapter 11, Reproduction). With this period added to gestation, pregnancy lasts 15 to 16 months. Most births seem to occur from mid-April to mid-June, with occasional calves born at any time of year (Fay, 1982). Females that have given birth nurse their calves for approximately 9 to 11 months, with the calves consistently taking fish on their own at 8 to 10 months of age (Fay, 1982).
0839_frame_C42 Page 929 Tuesday, May 22, 2001 2:08 PM
Walruses
929
Diet Prior to the early 1960s, attempts at rearing young walruses to adulthood achieved mixed success. Brown and Asper (1966) at Marineland of the Pacific felt many problems were related to nutritional imbalances. Artificial formulas (Chapter 37, Hand-Rearing) may differ depending on the experience of the staff, with the basic goal being the provision of a formula similar in composition to natural mother’s milk. Pathological fractures, seizures, and fatty bone marrow occasionally observed in young walruses may be related to excessive fat intake and insufficient calcium supplementation. The diet of the Pacific walrus consists of more than 60 genera of marine organisms, including mollusks, soft-shelled crabs, shrimp, sea cucumbers, tunicates, tube worms, soft coral, and occasionally portions of other pinnipeds (Lowry and Frost, 1981; Fay, 1982; Oliver et al., 1985). Walruses in captivity are fed a variety of whole fish and invertebrates (herring, smelt, capelin, sardines, squid, clams, and mackerel). A great deal of variation exists between diets at different institutions. The amount of food consumed by adult walruses varies with size and activity. The staff should be aware of the caloric makeup of each fish, in case bulk or caloric adjustments become necessary (see Chapter 36, Nutrition). Caution must be exercised to avoid overfeeding older animals resulting in obesity and poor health. Once a week fasting, a practice used for many carnivores held in zoological collections, or a 50% diet decrease 2 days a week, is recommended. Adult walruses should be weighed at regular intervals. Sexually active male walruses will increase food intake prior to their rut, but during the rut (January to April) food intake may decrease dramatically. The walrus may eat sporadically or not at all for days. During the rut, adequate time must be taken daily by the staff to observe the animal’s behavior, since loss of appetite at other times of the year is often the first sign of illness. Attitude, change in swimming pattern, change in haul-out time, change in posture or vocalizations, appearance of eyes, listlessness, and excessive nasal mucus should be noted. As with all animals on fish diets, care is taken to provide the best quality fish possible. Defrosting should take place under controlled conditions, with daily allocations maintained on ice to avoid short-term spoilage. Clams should be checked for signs of spoilage, such as odors and foreign material. Multivitamins with additional thiamine are given daily by placing them inside whole fish. Hand-feeding is recommended to discourage the “rooting” behavior along hard surfaces, which can cause abnormal tusk wear.
Physical Examination Physical examination of a walrus can be limited by animal size and accessibility. Animals that are trained for medical access, such as those who can calmly lie flat and open their mouths on demand, can be more closely examined than untrained individuals. Important information can be gained by monitoring daily activity, attitude, and animal interaction. Cardiac monitoring with a stethoscope is usually limited to trained, small to medium-sized animals, but should still be attempted in larger or untrained animals if feasible. Heart rates and respirations can be visually observed in animals lying out sunning. Caretakers should be adequately trained in physical evaluation so that they can distinguish between a walrus in good flesh vs. a thin animal or one that is obese. Individuals with excessive weight loss show an excess of loose folds of skin. Attitude and appetite are important indicators of potential medical problems. Depression, lethargy, loss of appetite, and nonresponsiveness to trainers may be early indicators of disease.
0839_frame_C42 Page 930 Tuesday, May 22, 2001 2:08 PM
930
CRC Handbook of Marine Mammal Medicine
Keepers and trainers should be taught to monitor stool and urine production daily and learn what normal respirations sound like. Staff provide important daily input on animal status upon which the veterinarian must depend. Unfortunately, noting changes in respiratory rate, or character, for signs of illness is not as productive in pinnipeds as in cetaceans, since respiratory cycles are not as regular. It is recommended that institutions housing walruses have a method available for obtaining periodic weights.
Restraint Manual Restraint of walruses, as with any large mammal, is potentially dangerous for both the animal and the handler. The agility of the animal in the water makes it necessary that any restraint be accomplished in a dry area or at the bottom of an empty pool. All staff involved in the procedure should be briefed about techniques and goals, and escape routes should be planned in the event of problems. During a netting procedure, handlers must be aware of their proximity to the animal’s head, which, with or without tusks present, can be very dangerous. In addition, handlers should avoid becoming placed between the animal and a wall, or entwining their fingers in the netting where they can become injured. Small- and medium-sized walruses can be adequately restrained for blood sampling by an experienced group of handlers with proper nets. Nets with a draw rope are very useful in limiting mobility. Squeeze cages have also been used but are difficult to apply to older animals. Larger animals can be walked onto a cargo net, which is pursed up by a crane and the animal suspended. This restraint method should be used only for short procedures, such as blood sampling or administration of injections. One or more individuals should monitor the animal’s responses, including attitude, respirations, and any signs of overheating. Water from a hose or sprayer must be available for spraying on the animal to avoid hyperthermia. All manual handling should be attempted only when the ambient temperature is relatively cool, such as early morning or evening. Walruses can also be trained to hold still for blood sampling, which can decrease the amount of physical restraint needed. Sick animals may not hold still for blood collection, so an alternative handling plan may be needed. Maintaining walruses without adequately training the personnel for handling procedures reduces the likelihood that a sick animal will receive the best treatment.
Sedation and General Anesthesia Sedation and anesthesia of walrus have been attempted with various classes of chemicals (DeMaster et al., 1981; Walsh et al., 1988; Tuomi et al., 1996), and are described in Chapter 29 (Anesthesia).
Specimen Collection and Diagnostic Techniques When walruses are individually trained, some samples can be obtained without restraint. However, individuals that are ill, even if trained, are not as cooperative, so training techniques cannot be entirely relied upon for sample acquisition. The diagnostic advantages of obtaining samples must be weighed against the possible stress and damage resulting from restraint. Normal ranges for walrus hematology are presented in Chapter 19 (Clinical Pathology), and are available in the MedArks database (and Wolk and Kosygin, 1979). Blood samples may be obtained from the vertebral sinus (similar to phocids) from trained walruses, or less reliably from interdigital vessels in the hind flippers (see Chapter 19, Clinical Pathology).
0839_frame_C42 Page 931 Tuesday, May 22, 2001 2:08 PM
Walruses
931
Swabs for bacterial and fungal cultures should be taken when clinically indicated. Common sites of clinical concern include the skin, tusk, fistulous tracts associated with tusk infection, the colon, and abscesses. Deep scrapings of the skin for fungus are more productive if the sample is cut directly into a fungal media plate. Cytology should also be performed on clinical samples (see Chapter 20, Cytology). Biopsies have been taken from skin lesions in trained animals by first injecting a local anesthetic then using a punch biopsy. Ultrasound has only been used to a limited degree because of animal size and the thickness of the skin (see Chapter 26, Ultrasonography). Ultrasound was used to detect an abdominal hernia in a young male walrus, and also the presence of fluid associated with severe intestinal disease. A fetus can be detected during an ultrasound examination if the mother is not too large, but in general the technology has not been adequately tested in the species. Radiography has been used for examination of the musculoskeletal system, such as the head and tusk area. The density of the tusk enamel is similar to the surrounding bone, so it is difficult to use this technique to examine the tusk unless there is infection present. Walruses have been trained to position themselves for radiographs of the head for evaluation of tusk complications. Abdominal radiographs can help to evaluate presence and location of foreign bodies. Dalton et al. (1990) describe radiographic techniques for the head and abdomen of walruses. Gastroscopy has been performed on a number of walruses under anesthesia and should be part of a standard evaluation for foreign bodies (see Chapter 27, Endoscopy). Laparoscopy has been performed on a walrus to evaluate the liver and obtain biopsies (Dover et al., 1998). It may also be an important evaluation before committing to a laparotomy.
Medical Problems Fay (1985) noted the following pathological conditions of walruses in the wild: intraspecific trauma (trampling, tusk injury), parasites (Trichinella spp., lice), tusk pulpitis (7 of 2000 examined), urethritis, pleuritis, pneumonia, subdermal bullae, aneurysm (one case), cystic ovaries, umbilical hernia (one case), renal calculi (one case), frostbite, exhaustion, and malnutrition. Brown (1962) summarized the health problems of four walruses being raised at Marineland of the Pacific. These included parasites (marine louse Antarctophthirius trichechi), epidermal abscesses (Staphylococcus, Streptococcus, and Pseudomonas spp. cultured), ophthalmic conditions (blepharitis, conjunctivitis), and a central nervous system disorder (seizure). The seizuring animal recovered after 6 days. Clinical description of the neuromuscular problems of the animal resembled that seen with calcium deficiency or hyponatremia, although no blood was taken for diagnosis.
Dermatology Walruses have light brown hair that gradually becomes patchy until the next molt. The condition of the coat may be impacted by the swimming patterns of the individual. Animals maintained at SeaWorld have been observed with oval areas of inflammation of varying sizes scattered diffusely over the body. A dermatophyte, Trichophyton spp., has been cultured from these cases, although biopsies did not always indicate that the fungus was invading the skin. Although histologically these lesions suggest a possible insect hypersensitivity, they have been clinically responsive to the azole antifungals (see Chapter 17, Mycotic Diseases). The lesions have not been responsive to superficial application of solutions containing iodine. Dalton (pers. comm.) has found viral particles associated with similar lesions. Pustular folliculitis has been observed in three walrus. Biopsies indicated a bacterial etiology, with Staphylococcus and Corynebacterium cultured from the lesions. Bacterial and fungal isolates from skin of walruses at SeaWorld of Florida are listed in Table 1. Interdigital ulcerative
0839_frame_C42 Page 932 Tuesday, May 22, 2001 2:08 PM
932
CRC Handbook of Marine Mammal Medicine
TABLE 1 Bacterial, Yeast, and Fungal Isolates from Walrus Sites and Specimens Skin Streptococcus faecalis Staphylococcus xylosus Klebsiella pneumoniae Streptococcus faecium Streptococcus spp. β-Hemolytic Streptococcus Shewenella putrifaciens Citrobacter koseri Corynebacterium pseudotuberculosis Staphylococcus Candida albicans Trichophyton rubrum Aspergillus Tusk-Fistulous Tract Streptococcus avium S. faecalis S. viridans Escherichia coli Proteus mirabilis Morganella morganii Yersinia enterocolitica Vibrio metschnikovii Corynebacterium-like Bacteroides Arcanobacterium haemolyticum
Fecal Citrobacter spp. Escherichia coli Streptococcus, Group D Proteus mirabilis Pseudomonas (fluorescent group) Enterococcus feacalis Salmonella spp. Morganella morgani Yersinia pseudotuberculosis Aeromonas hydrophila Edwardsiella tarda Actinobacter calco Yersinia enterocolitica Candida parapsilosis Ear Cultures Escherichia coli β-Hemolytic Streptococcus Corynebacteria haemolyticus Actinomyces naeslundii Eubacteria Candida spp.
dermatitis associated with yeast infection has been responsive to antifungals. When treating skin conditions, it is important to evaluate the environment for complicating factors, such as retention of fecal debris and urine in commonly used haul-out areas. Water quality parameters should also be reviewed (see Chapter 35, Water Quality). Superficial skin lacerations are usually from tusks of other animals, since sparring is a constant activity between males and is increased during breeding. Females may be constantly harassed and have numerous bruises and lacerations that are very evident by thermography (see Chapter 28, Thermography).
Ophthalmology Diseases of the walrus eye are common, so it is important that the animal be trained for treatment when problems arise. Walruses maintained in water of low salinity (<3%) tend to develop more corneal problems than those in more saline pools. If not corrected, minor eye problems can lead to chronic conditions. Low-salinity environments can also lead to eye problems secondary to injury. Corneal edema often resolves when the eye is treated with hypertonic saline (5%) applied three to five times a day. Corneal ulcers, cataracts, and lens luxations have been seen. Cataract removal in a juvenile individual has been successful (Briggs, pers. comm.). One walrus at SeaWorld currently appears to have allergies that are expressed as swollen eyelids that respond to topical steroids. Traumatic eye lesions may occur, especially if animals are of breeding age.
0839_frame_C42 Page 933 Tuesday, May 22, 2001 2:08 PM
Walruses
933
An ophthalmologist must be involved in the early stages of an ocular condition, so he/she may become familiar with the ocular anatomy of the walrus. Most diagnostic approaches and treatments are similar to those used in domestic animals, although the animal may need to be maintained out of water for a few minutes after administration of medication. The medical staff should help the caretaker staff become familiar with ophthalmic conditions to monitor.
Tusk Infections and Trauma Tusk pulpitis, associated with worn tusks and fistulous tracts in the nasal area, has been observed in numerous walruses in captivity (Sweeney, 1985). Etiological factors that may be involved include the individual’s behavior, facility design, and tusk anatomy. The tusk has a central core of vascular globular dentine. When the distal portion of the tusk is rubbed close to the pulp (usually within a few inches), bacteria may gain access to the pulp cavity through the central core, resulting in pulpitis. Bacteria isolated from tusk pulpitis or secondary fistulous tracts are given in Table 1. This condition may proceed unnoticed, until swelling is observed over the nasal area. The presence of swelling indicates that the dorsal bone over the tusk base has been compromised and that osteomyelitis has started. At this point, the walrus may become lethargic and lose its appetite. Temporary relief can be obtained by incising the swelling to establish drainage. However, this can be dangerous to veterinarian and animal alike. Often these individuals may not cooperate to allow close access for drainage establishment, so may require sedation. Determination of the best point for drainage may be difficult. Once opened, aerobic and anaerobic bacterial cultures are taken to guide possible antibiotic therapy. Although this therapy may temporarily result in resolution of symptoms, the clinician should prepare for possible surgical intervention. The treatment of choice for pulpitis with fistulous tracts is removal of the tusk. Since the tusk is continually growing, a root-canal procedure is not indicated. In the past, root-canal procedures were attempted after infection was already grossly present. A previously reported technique for tusk removal (Cornell and Joseph, 1985) had the disadvantage of potentially causing multiple fractures in the lateral alveolar bone. An improved technique, a modification of an elephant tusk removal technique, has been used on 11 walruses (Welch et al., 1988). This technique involves hollowing out the tusk with a drill bit, leaving only a thin wall of cementum and dentine. This ring of tusk material is longitudinally split into three to four sections with a saw blade, and each section is separated from the surrounding bone with a bone gouge. This technique avoids fracturing the bone and aims at preventing chronic complications of fracture and sequestrum formation. It is important that the pulp tissue be completely removed after the tusk is pulled to avoid tusk regrowth (Cook et al., 1990). The authors use a strong betadyne solution (10%) and thoroughly clean the tusk cavity with a wire shotgun-cleaning brush to ensure removal. The cavity is also visually inspected and flushed repeatedly. Tusk regrowth after extraction is usually disorganized, and difficult to remedy. The separation between bone and dentine is irregular, so the standard removal technique is not as easy. Additionally, the regrowth usually occurs behind the healing maxillary bone, interfering with normal elongation of the tusk. Many are also chronically infected with secondary fistulous tracts. One walrus died secondary to a chronic tusk infection that migrated to the brain. Prevention of tusk wear has been attempted somewhat successfully by placing ticonium caps over the end of the worn tusk before it becomes contaminated (Briggs et al., 1998). Potential disadvantages include need for constant vigilance and replacement, traumatic removal of the tusk from entrapment on facility material, and increased abrasion of acrylic panels. The time between loss of a cap and replacement can result in increased abrasion, infection, and the need for surgical removal.
0839_frame_C42 Page 934 Tuesday, May 22, 2001 2:08 PM
934
CRC Handbook of Marine Mammal Medicine
Foreign Bodies Walruses are prone to ingesting many objects, including leaves, sticks, rocks, hardware, and anything that may be tossed or dropped into an exhibit. The intestinal tract of the walrus narrows at the junction of the small and large intestines, so foreign objects can lodge there. Clinical signs include depression, abdominal distress, lack of appetite, vomiting (with or without debris presented), decrease or lack of feces, and staying out of water. Sometimes the authors will give a pain relief drug such as flunixin meglumine (0.5 mg/kg IM) as a diagnostic tool. Typically, the animal will eat better on the medication, but relapse into anorexia once treatment ceases. Radiographs may be helpful, but do not detect common objects such as balls or baby pacifiers. Ultrasound may not be helpful or possible, so the diagnosis is often based on clinical judgment and symptoms. The treatment of choice is surgery, which can be a difficult consideration for the inexperienced clinician who must master diagnosis, anesthesia, and surgery in a short amount of time. Exploratory laparotomies have been performed on at least six walrus at various facilities to date. The basic approach is to include large-animal surgeons in the procedure, to help reduce the risk of surgical error. Often animals may be toxic because of delays in choosing surgery, so it is important to maintain an open intravenous line. This can sometimes be performed peripherally at the interdigital veins of the flippers, but the authors use the vertebral sinus. A sialastic catheter is threaded through a larger-bore needle with a distal curvature (Cook Medical, Winston-Salem, NC) into the vertebral sinus. The needle is removed and the catheter sutured in place and connected to a fluid setup. One animal died shortly after relieving an obstruction, possibly because of release of toxic compounds. On necropsy of this walrus it was noted that the heart might have impinged on the caudal vena cava if the animal had been placed straight on its back in dorsal recumbency. As a result, the authors choose to lay walruses at a slightly oblique angle during prolonged procedures. Another problem may be dehiscence of the surgical site. This can occur because of inadequate closure for the pressure exerted, as this species drags its abdomen along the ground. To avoid dehiscence, the skin incisions are closed with 20-gauge wire in two patterns. The first pattern is a single row of simple, interrupted sutures placed 2.5 cm apart. The second pattern, also of simple interrupted sutures, is used as a tension-relieving pattern, with a single suture placed between two sutures of the first pattern. Since walrus skin is thick and difficult to penetrate any substantial distance with a suture needle, an 18-gauge, 3in. spinal needle is preplaced 3.5 cm from each side of the incision. The wire is guided through the needle with each suture, spanning 7 cm across the incision. Prevention of foreign body involvement can be partially achieved by proper facility design. The four main sources of foreign bodies in walrus facilities are construction materials that are not walrus-proof, materials presented by the general public, materials presented by caretaker staff, and materials presented by the surrounding natural environment (such as leaves). The walrus environment should have limited public access to decrease material being dropped or thrown into the habitat. It should have netting to eliminate leaves and debris if surrounded by natural vegetation. It should have recessed caulking to avoid removal and ingestion by the very strong suction ability of walrus, and all bolts must be engineered to avoid removal. With walrus, any material that can fit into their mouths will most likely be swallowed.
Intestinal Disease Nonspecific gastroenteritis has been observed in a number of walruses. Clinical signs include lethargy, staying dry out of water, lying in a tucked fetal position on land, or floating high in the water in a tucked position, suggesting intestinal gas. Because of the lack of specific signs, diagnosis is difficult, but hematology, serum chemistry, aerobic and anaerobic stool cultures, and fecal cytology may be helpful. Treatment is often symptomatic, utilizing antigas compounds
0839_frame_C42 Page 935 Tuesday, May 22, 2001 2:08 PM
Walruses
935
(simethicone), coating solutions, and possibly antibiotics (see Chapter 31, Pharmaceuticals). These symptoms can also relate to foreign body ingestion, so if feces are not observed, mineral oil (500 ml for a 450-kg animal) is given two to three times a day for up to 5 days. If appetite is totally lost, flunixin meglumine can be administered intramuscularly to improve appetite and maintain caloric and hydration needs. This drug should not be used in dehydrated individuals, or for extended periods of time, without periodic evaluation of renal status. If no response is seen after two doses and blood results are nonspecific, foreign body involvement is a possibility. Feces of animals showing signs of unexplained enteritis must be cultured in an attempt to establish a link between illness and organisms present. Salmonella spp. have been observed in three juvenile walruses (Calle et al., 1995). Clinical signs included partial to complete anorexia, regurgitation, diarrhea, lethargy, depression, and weight loss. Salmonella muenchen, S. anatum, S. alachua, S. istanbul, S. hadat, and an untypable Salmonella sp. were all recovered at different times in these three juveniles, and two older walruses during the course of treatment. Gulls might have been the source of the original infection.
Miscellaneous Diseases Clostridium perfringens has been associated with the death of a walrus caused by trauma and contamination of an injection site. The effects of the type A toxin were severe myositis, toxininduced necrosis of the liver, brain, and heart, and respiratory compromise (Murnane et al., 1997). Pseudomonas aeruginosa has been isolated from a severe case of mastitis in a walrus. This walrus showed anorexia, anemia, weakness, discomfort, and swelling of the left rear flipper. The left mammary quadrants were severely affected and appeared to influence the swelling of the left leg. Caliciviruses (see Chapter 15, Viral Diseases) (San Miguel sea lion virus) have been recovered from walrus feces, and serology has indicated a wide exposure to this group of viruses (Barlough et al., 1985; Smith et al., 1983, 1989; Poet and Smith, 1991). Morbillivirus is a potential disease of walruses. Calle et al. (1998) tested 158 walruses for the presence of antibodies to phocine morbillivirus, Calicivirus, Brucella, and Leptospira, and found this group negative for antibodies to these pathogens. Advanced renal disease was observed in an aged walrus by Joseph and Torgerson (1989). Walruses maintained in areas where heartworm disease is endemic should be placed on heartworm prevention, as it is known that other pinnipeds can develop clinical heartworm (Beusse et al., 1977). Oral ivermectin is currently used once a month, although diethylcarbamazine was the standard for many years. At present, no vaccines are currently recommended for use in walruses. As veterinarians document diseases seen in walruses, the level of care given to the species will continue to improve. This will also lead to better understanding of walruses in the wild.
Acknowledgments The authors thank Dr. Danise Martinez for her help with the manuscript, and all those who appreciate the uniqueness of this special species.
References Barlough, J.E., Berry, E.S., Skilling, D.E., and Smith, Q.W., 1985, The AYKOVO Collections: Marine calicivirus antibodies in the Pacific walrus, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive.
0839_frame_C42 Page 936 Tuesday, May 22, 2001 2:08 PM
936
CRC Handbook of Marine Mammal Medicine
Beusse, D.O., Asper, E.D., Baucom, J.N., and Searles, S.W., 1977, Diethylcarbamazine citrate for prevention of heartworm (Dirofilaria immitis) in the California sea lion (Zalophus californianus), Vet. Med. Small Anim. Clin., 470. Briggs, M.B., Meehan, T.P., Zdziarski, J., Messinger, D., and Willis, G.P., 1998, A technique for preventing wear and maintaining the integrity of young walrus (Odobenus rosmarus) tusks, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Brooks, J.W., 1954, A contribution to the life history and ecology of the Pacific walrus, Special Report 1, Alaska Cooperative Wildlife Research Unit, University of Alaska, Fairbanks, 103. Brown, D.H., 1962, The health problems of walrus calves, and remarks on their general progress in captivity, Int. Zoo Yearb., 4: 13. Brown, D.H., and Asper, E.D., 1966, Observations on the Pacific walrus (Odobenus rosmarus divergens) in captivity, Int. Zoo Yearb., 6: 78. Calle, P.P., Stetter, M.D., Cook, R.A., McClave, C.A., Massucci, S., and Walsh, K.M., 1995, Enteric salmonellosis of captive Pacific walrus (Odobenus rosmarus divergens), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Calle, P.P., Seagers, D.J., McClave, C., Senne, D., House, C., and House, J.A., 1998, Infectious disease serology of free-ranging Alaskan Pacific walrus (Odobenus rosmarus divergens), in Proceedings American Association of Zoo Veterinarians and the American Association of Wildlife Veterinarians Joint Conference. Cobb, W.M., 1933, The dentition of walrus (Odobenus obesus), Proc. Zool. Soc. London, 103: 645. Cook, R.A., Klein, L., Welch, B.B., and Walsh, M.T., 1990, Tusk regrowth following surgical removal in a female Pacific walrus (Odobenus rosmarus divergens), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Cornell, L.H., and Joseph, B.E., 1985, Anesthesia and tusk extraction in walrus, in Proceedings of the American Association of Zoo Veterinarians, 97. Dalton, L.M., Mathey, S.W., and Hines, R.S., 1990, Radiology as a diagnostic aid in marine animal medicine, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CDROM Archive. DeMaster, D.P., Faro, J.B., Estes, J.A., Taggart, J., and Zabel, C., 1981, Drug immobilization of walrus (Odobenus rosmarus), Can. J. Fish. Aqua. Sci., 38: 365. Dover, S.R., Kolata, R., and Walsh, M.T., 1998, The development of laparoscopic techniques for use in marine mammals, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Fay, F.H., 1982, Ecology and biology of the Pacific walrus Odobenus rosmarus divergens (Illiger), North Am. Fauna (Washington, D.C.), 74: 1. Fay, F.H., 1985, Odobenus rosmarus, Am. Soc. Mamm. Species, 238: 7. Joseph, B.E., and Torgerson, R.W., 1989, Advanced renal disease in an aged Atlantic walrus (Odobenus rosmarus): Emphasizing clinical presentation and therapy, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Lowry, L.F., and Frost, K.J., 1981, Feeding and tropic relations of phocid seals and walruses in the eastern Bering Sea, in The Eastern Bering Sea Shelf: Oceanography and Resources, Vol. 2, Hood, D.W., and Calder, J.A. (Eds.), Office of Marine Pollution Assessment, NOAA, University of Washington Press, Seattle. Mansfield, A.W., 1958, The biology of the Atlantic walrus, Odobenus rosmarus (Linnaeus), in the eastern Canadian Arctic, Fisheries Research Board of Canada, Manuscript, Rep. Serv. (Biol.), 653. Murnane, R.D., Kinsel, M.J., and Briggs, M.B., 1997, Clostridium perfringens type A-induced myositis with probable toxemia in an adult walrus (Odobenus rosmarus), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Oliver, J.S., Kvitek, R.G., and Slattery, P.N., 1985, Walrus feeding disturbance: Scavenging habits and recolonization of the Bering Sea benthos, J. Exp. Mar. Biol. Eco., 91: 233.
0839_frame_C42 Page 937 Tuesday, May 22, 2001 2:08 PM
Walruses
937
Poet, S.E., and Smith, A.W., 1991, A DNA hybridization probe for detecting caliciviruses from the sea, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Reeves, R.R., 1978, Atlantic walrus (Odobenus rosmarus): A literature survey and status report, U.S. Fish Wildlife Research Report, 10: 1. Reeves, R.R., 1979, The Atlantic walrus in retreat, Nat. Parks Conserv. Mag., 53: 10. Ridgway, S.H., and Harrison, R.J., 1981, Handbook of Marine Mammals, Vol. 1, The Walrus, Sea Lions, Fur Seals and Sea Otter, Academic Press, London, 1. Samansky, T., and Sieswerda, P., 1995, The recovery, care, and transport of orphaned walrus calves at St. Lawrence Island, Alaska, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Smith, A.W., Ritter, D.G., Ray, G.C., Skilling, D.E., and Wartzok, D., 1983, New calicivirus isolates from feces of walrus (Odobenus rosmarus), J. Wildl. Dis., 19: 86–89. Smith, A.W., Skilling, D.E., and Poet, S.E., 1989, Marine caliciviruses and diseases of pisces, pigs, pinnipeds and people, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Sweeney, J.C., 1985, Clinical management of maxillary abscess, cellulitis, and osteomyelitis due to tusk wear in a juvenile walrus, Odobenus rosmarus, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Tuomi, P.A., Mulcahy, D.M., and Garner, G.W., 1996, Immobilization of Pacific walrus (Odobenus rosmarus divergens) with carfentanil, naltrexone reversal and isoflurane anesthesia, in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Walsh, M.T., Webb, A.I., Beusse, D.O., Brock, K.A., Robertson, S.A., Abou-Madi, N., Cook, R.A., and Klein, L., 1988, Sedation and general anesthesia of four Arctic walrus (Odobenus rosmarus), in Proceedings of the International Association for Aquatic Animal Medicine, IAAAM CD-ROM Archive. Welch, B., Walsh, M.T., and Buesse, D.O., 1988, Tusk extraction in the walrus, in Proceedings of the International Association for Aquatic Animal Medicine, 186. Wolk, E., and Kosygin, G.M., 1979, A hematological study of the walrus, Odobenus rosmarus, Acta Theriol., 24: 99.
0839_frame_C42 Page 938 Tuesday, May 22, 2001 2:08 PM
0839_frame_C43.fm Page 939 Tuesday, May 22, 2001 2:08 PM
43 Manatees Gregory D. Bossart
Introduction The Florida manatee (Trichechus manatus latirostris), with an estimated population of about 2300 animals, is one of the most endangered marine mammals in the coastal waters of the United States (U.S. Marine Mammal Commission, 1999) (see Chapter 3, Manatee Case Study). Manatee medicine has progressed rapidly during the past 10 years. Critical-care facilities in Florida use the latest diagnostic and treatment techniques for manatees in rescue and rehabilitation conservation programs. Artificial milk formulas have been developed for raising orphaned manatee calves (see Chapter 37, Hand-Rearing), and baseline clinicopathological reference ranges (see Chapter 19, Clinical Pathology) have been established for adults and calves. Manatees have been maintained in captivity since 1875 (Crane, 1881; True, 1884). Based on long-term observations of captive manatees and necropsy results from over 20 years, it appears that adult manatees are extremely hardy, with successful captive maintenance directly related to good water quality and nutrition (White and Francis-Floyd, 1990). In one study of pathology in 16 manatees, only one case of natural disease was described (Buergelt et al., 1984).
Natural History The Florida manatee is a large, herbivorous, totally aquatic mammal that is one of four living species in the order Sirenia (Odell, 1982) (Figure 1). Sirenians are believed to have evolved from quadripedal land mammals more than 60 million years ago. The closest living terrestrial relatives of the Sirenia are the Proboscidea (elephants) and Hyracoidea (hyraxes) (Domning, 1994). The Florida manatee, because of its evolutionary development, shares many unusual hematological and immunological features with the elephant. Sirenians live in tropical and subtropical regions around the world, and include one species of dugong (Dugong dugon), generally found in coastal regions of the Indopacific; the West African manatee (Trichechus senegalensis), found in coastal waters, rivers, and lakes of western and west central Africa; the Amazonian manatee (Trichechus inunguis), which is restricted to the freshwater rivers and lakes of the Amazon basin; and the West Indian manatee (Trichechus manatus) (Lefebvre et al., 1989). The West Indian manatee has two subspecies: the Antillean manatee (Trichechus manatus manatus), found in the West Indies, the Caribbean, and coastal waters and rivers of Mexico, Central America, and northeastern South America, and the Florida manatee, found in the coastal waters and rivers of the southeastern United States and Gulf of Mexico, as far west as the Texas coast. Florida is essentially the northernmost range for the
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
939
0839_frame_C43.fm Page 940 Tuesday, May 22, 2001 2:08 PM
940
CRC Handbook of Marine Mammal Medicine
FIGURE 1 An adult female Florida manatee. (Photo credit: G. Bossart.)
West Indian manatee, as manatees are intolerant of cold weather, although some animals may range farther north to the mid-Atlantic Coast of the United States during the summer. During the winter, Florida manatees can be found in warm-water natural springs, southern Florida canals and lakes, and warm-water effluents of electric power plants and other industrial sources (Irvine, 1983). The Florida manatee is unusual in that it can move freely between salinity extremes. It can live for extended periods in freshwater, brackish, and marine habitats, and can be found in clear, muddy, and heavily polluted water. Manatees usually prefer water depths of 0.9 to 2.1 m (3 to 7 ft) for resting and grazing on various seagrasses and freshwater plants. Typical Florida manatee habitat is shallow coastal waters, lakes, and rivers on both the east and west coasts of Florida. Tracking studies indicate that manatees often migrate large distances along the East Coast of the United States. Most manatees migrate seasonally between winter gathering sites and summer distribution areas. Unlike many marine mammal species, manatees are considered minimally social (Hartman, 1979). Except for cow–calf relationships that may last for 2 years, and cold weather–related aggregations, most relationships appear temporary. Additionally, the Florida manatee is not territorial and does not display apparent intraspecific or interspecific aggression. Temporary social interactions may include behaviors such as mouthing, bumping, chasing, body surfing, group somersaulting, barrel rolling, and upside-down gliding. It is difficult to estimate the manatee population accurately because of the unique characteristics of the manatee and the environment it inhabits. Manatee counts are highly variable, and, to date, an accurate way to estimate numbers of animals during population surveys has not been found (U.S. Marine Mammal Commission, 1999). In 1996, a winter population survey identified 2639 manatees. In 1997, two population surveys were conducted during the winter; a January survey yielded a count of 2229, whereas a February survey resulted in a count of
0839_frame_C43.fm Page 941 Tuesday, May 22, 2001 2:08 PM
Manatees
941
1709 manatees. An increasing number of documented manatee deaths since 1978, including a large proportion of human-related deaths, have caused serious concerns about the long-term survival of this species.
Anatomy, Physiology, and Behavior A basic understanding of the manatee’s biologic character is helpful, and an in-depth anatomical review is provided by Rommel and Lowenstine in Chapter 9, Anatomy. Manatees are large, fusiform mammals with flat, rounded, spatulate tails. Adults can reach a mean length of 2.7 to 3 m (9 to 10 ft) and weigh between 410 and 545 kg (900 and 1200 lb). Maximum length of 3.6 m (12 ft) and weight of up to 1775 kg (3900 lb) have been reported. Female manatees tend to be larger and heavier (Odell, 1982). The manatee has a low metabolic rate (Irvine, 1983), which may seriously restrict this large, tropical mammal’s ability to maintain body temperature in the colder winter weather of northern Florida and which may account for the manatee’s susceptibility to cold. The head of the manatee is unique. A flexible, prehensile upper lip acts as a “shortened trunk” that is similar to the trunk of its terrestrial cousin, the elephant. The lip is used to gather, manipulate, and direct plant matter into the mouth. Both lips are highly tactile and contain many modified vibrissae (perioral bristles) (Reep et al., 1998). In addition to their use in food gathering, the lips are important in social interaction and communication. The nostrils are located dorsally on the snout, and have valves that close when the manatee dives. Eyes are small and widely spaced with eyelids that close in a circular manner. Histological examination of the retina suggests that manatees see color. Depth perception is poor, but long-distance acuity appears to be good (Hartman, 1979). Manatees can emit a wide range of sounds that are used for communication, rather than echolocation as in cetaceans. These sounds are typically used to maintain contact with other adults, especially during sexual and play behaviors. Sound communication is especially prominent between cows and dependent calves. In addition to sound, sight, and touch, manatees may communicate through taste and smell. The digestive system of the manatee is extraordinary, and probably reflects its herbivorous diet and the incidental ingestion of water of varying salinity (Burn, 1986; Burn and Odell, 1987; Reynolds and Rommel, 1996). Incisors are absent and are replaced by horny gingival plates. The molar teeth are uniformly shaped, but are of different sizes and are continually replaced in a forward direction when worn. The manatee has a gastrointestinal tract characterized by an enlarged hindgut, as do other nonruminant herbivores, such as horses. Other gross anatomical adaptations include a simple, saccular stomach with a discrete accessory digestive gland (the cardiac gland), a large duodenal ampulla with paired duodenal diverticulae, and a large cecum with paired cecal diverticulae (see Chapter 9, Anatomy). Hindgut digestion of cellulose produces abundant gas and flatulence. Indeed, the presence (healthy) or absence (ill) of flatulence can be used as a prognostic indicator for manatees. Manatees graze 5 or more hours/day, and typically consume 4 to 10% of their body weight in wet vegetation per day (Best, 1981). The reproductive physiological characteristics of manatees are largely unknown. Anatomically, male manatees have a genital opening just caudal to the umbilicus. Females have a genital opening just cranial to the anus. Females have prominent axillary teats. Manatees have low reproductive rates. Typically, a single calf is produced with a birth interval of approximately 3 to 5 years. The gestation period is about 13 months, and calves are dependent on their dams for approximately 2 years after birth. Newborn calves are about 1.2 m (4 ft) long and weigh
0839_frame_C43.fm Page 942 Tuesday, May 22, 2001 2:08 PM
942
CRC Handbook of Marine Mammal Medicine
approximately 32 kg (70 lb). Calves nurse underwater for 3 to 5 min every 1 to 2 hours around the clock. Manatee milk contains more fat and protein when compared with milk of other mammals, and contains little or no lactose (Bachman and Irvine, 1979; Walsh and Bossart, 1999) (see Chapter 36, Nutrition). Contrary to popular assumption, manatees are capable of understanding discrimination tasks, and they show signs of complex associated learning and advanced long-term memory (Gerstein, 1994). This has been partially demonstrated through successful transfer of generalized tasks. In behavioral tests, manatees exhibit complex discrimination and task-learning abilities on a par with dolphins and pinnipeds in similar acoustic and visual studies.
Husbandry Habitat Requirements Space requirements based on physical, behavioral, social, or psychological factors are poorly understood for the manatee. Manatees have been successfully maintained in pools with a wide range of depths and volumes at the Miami Seaquarium (Florida) for over 43 years and successfully reproduced in such environments (Zeiler, 1978; White, 1984). Manatee rehabilitation programs in the Colombian Amazon, Belize, and Mexico have successfully reared orphan manatee calves in freshwater cisterns or molded plastic child swimming pools measuring approximately 2.5 m (8 ft) in diameter and less than 1.0 m (3.3 ft) deep (Bossart and Menchaca, 1998; Bossart, unpubl. data). However, in most situations larger pools should be available to accommodate physiological, developmental, and behavioral needs. Ideally, a medical treatment pool should be included in the habitat design. This pool should have a lift system for “stranding” the manatee for examination and treatment or a pump system to drain water rapidly for treatment on the pool bottom. Minimum housing requirements for manatees are described in the Animal Welfare Act (see Chapter 33, Legislation).
Water Requirements Free-ranging Florida and Antillean manatees can live for extended periods in fresh, brackish, and/or seawater habitats, although in captivity they are usually maintained in fresh water. However, the Miami Seaquarium has maintained manatees in seawater (a salinity of approximately 32 parts per thousand, ppt) since 1957 (White, 1984), but provides fresh water at all times as a drinking source. No physiological or pathological abnormalities have been observed in these captive manatees. Manatees maintained in fresh water show an increase in skin sloughing after being moved from fresh water to salt water (Walsh and Bossart, 1999). Chlorine in concentrations of less than 1 part per million (ppm) has been used as a disinfectant in some manatee habitats. Chlorine can be used with ozone to reduce total chlorine levels. However, chemical treatment of the water should not replace the absolute need for filtration systems that can effectively and rapidly handle the extraordinary bioload produced by vegetable food material and abundant fecal production. Captive manatees are sensitive to cold temperatures and should be maintained in water ranging from 23 to 30°C (74 to 86°F). The first signs of “cold stress syndrome” can be seen in manatees exposed to water temperatures below 20°C (68°F) for 2 or 3 days. These signs may include shivering, partial or complete anorexia, and changes in swimming, resting, and respiration patterns. Prolonged exposure to cold water may result in opportunistic diseases and compromised immune function (Bossart, 1995). Facilities should have water heaters available for use if the ambient water temperature drops below 23°C (74°F). Adult Florida manatees
0839_frame_C43.fm Page 943 Tuesday, May 22, 2001 2:08 PM
Manatees
943
can adapt to high water temperatures of greater than 32°C (90°F). Antillean manatees at a zoo in Guyana live in summer water temperatures of 40°C (104°F) (Bossart and Menchaca, 1998). These values likely represent the extremes, and water temperatures should not exceed 30°C (86°F), if at all possible.
Nutrition Manatees often are indiscriminate and opportunistic feeders that graze on a wide variety of plant materials (Hartman, 1979; Best, 1981; Bengston, 1983). In fresh water, free-ranging manatees will feed on hydrilla (Hydrilla verticillata), water hyacinth (Eichhornia crassipes), and water lettuce (Pistia philoxeroides); and in salt water on manatee grass (Syringodium filiforme) and turtle grass (Thalassia testudinum). Manatees have also been observed grazing on leaves, acorns, nuts, and tree branches overhanging the water (Hartman, 1979). In Guyana, manatees have been seen partially hauling out of the water and eating grass on the shoreline (Bossart, unpubl. obs.). In the United States, captive manatees have been fed a wide variety of foods, including fresh, hand-collected manatee and turtle seagrasses, lettuce (romaine, iceburg, and others), cabbage, kale, carrots, sweet potatoes, bananas, apples, timothy and alfalfa hays, hydroponically grown wheat and oats, various commercially available monkey chow biscuits (Monkey Diet, PMI International, Inc., Brentwood, MO) and other feeds designed for terrestrial species (Caron, pers. comm.; White and Francis-Floyd, 1990; Walsh and Bossart, 1999). In Mexico and South America, captive manatees in rescue/rehabilitation programs and zoos have been fed fresh, hand-collected natural water grasses (e.g., water hyacinth), fresh green cattle pasture grasses, locally-grown greens and vegetables for human consumption (e.g., spinach, squash, melon), and lawn grass clippings (Bossart and Menchaca, 1998). The exact nutritional requirements of the manatee are unknown. However, many of the above dietary regimens have provided nourishment for animals from weaning to reproductive adulthood with concomitant successful rearing of healthy calves. A varied diet with calcium/ phosphorus supplementation correlated with dramatic improvements in reproductive success at the Miami Seaquarium during the early 1980s (White and Francis-Floyd, 1990). In this study, vitamins were added in bananas or capelin (Mallotus villosus). A general guideline is to provide a diet that incorporates 70 to 85% leafy green vegetables (e.g., romaine and iceburg lettuce, cabbage, kale) or hydroponic wheat, oats, and/or sprouts; 10 to 20% dried forage such as timothy or alfalfa hay (filtration must be upgraded to handle dried matter), about 5% of vegetables and fruits (carrots, yams, apples, bananas, etc.), and various commercially available dry pelleted feeds (monkey chow, elephant “flakes,” etc.) (Walsh and Bossart, 1999). The specific vitamin and mineral requirements for manatees are unknown. Some facilities with captive manatee management programs use empirical oral vitamin supplements. Adult males and nonlactating females consume from 7 to 9% of their body weight/day (Best, 1981; Bengston, 1983; Etheridge et al., 1985); calves (>175 cm, or 68 in., length) consume about 15% of body weight/day (Etheridge et al., 1985); and lactating females eat from 10 to 13% of their body weight/day (Best, 1981). Manatees that are in rehabilitation/release programs must not, if at all possible, be fed diets that are vastly different from natural diets. However, approaching natural diets in captivity may be difficult. A possible solution to this problem has been to wean these manatees gradually off artificial diets and replace the diets with a natural diet of various fresh and seawater grasses (Caron, pers. comm.). This process involves the daily field collection of fresh grass. Additionally, weaned manatees that will be released should not be hand-fed. It has been reported that free-ranging manatees feed primarily on submerged plants and tend to ignore floating and emergent vegetation (Hartman, 1979). However, captive manatees
0839_frame_C43.fm Page 944 Tuesday, May 22, 2001 2:08 PM
944
CRC Handbook of Marine Mammal Medicine
are generally fed by surface food distribution. Underwater bottom devices, which trap or contain food for bottom feeding, may also be used. Free-ranging manatees graze 5 or more hours/day (Best, 1981), so captive daily feeding frequencies should be high enough to allow for this behavior. Smaller quantities of food given more frequently during the day will permit normal grazing behavior and help prevent filtration systems from being overwhelmed. From a nutritional standpoint, orphan manatee calves are a unique challenge for veterinarians. Nutritional support for calves has usually been based on commercial products with varying degrees of success. Manatee milk contains about 74% moisture, 8% protein, 16% fat, 1% ash, and 1% carbohydrates supplying about 189 cal /100 g (Walsh and Bossart, 1999). Further analyses have demonstrated that manatee milk has high levels of taurine and methionine, as well as short- and medium-chain fatty acids (Offtedal, pers. comm.). Many different artificial nursing formulas have been used to raise manatees. Formula standardization may prove useful in minimizing the frequent occurrence of intestinal disease in these animals. Formulas have included various combinations of milk and soy-based commercial products, goat’s milk, and bitch’s milk mixed with additives such as Lactobacillus, vitamins, canola oil, taurine, and short-chain fatty acids (see Chapter 36, Nutrition; Chapter 37, Hand-Rearing). Manatee calves in Guyana have been successfully raised on a nursing formula of exclusively cornmeal and cow’s milk (Bossart and Menchaca, 1998). Calves can be offered formula via a bottle with a lamb’s or calf ’s nipple and should be fed every 2 to 3 hours around the clock. The incidence of intestinal complications seems to be reduced if special attention is paid to the cleaning and sterilization of nursing equipment prior to each use. Observations of calves born at the Miami Seaquarium indicate that manatees will nurse until 18 to 24 months of age and start nibbling on lettuce as early as 11 days of age (Bossart, unpubl. data; Odell, 1978). If necessary, manatee calves can be weaned as early as 8 months of age.
Restraint, Handling, and Transport Manatees are extremely powerful and will thrash dorsoventrally, laterally, and roll violently during restraint procedures. The fluke is especially dangerous during struggling. These movements can easily cause serious injury to handlers and animal caregivers. In water rescue situations, handlers must use extreme caution to avoid becoming entangled in the net and pulled into the water (Geraci and Lounsbury, 1993). Manatee calves also are strong and should be restrained with caution. Injury to the manatee or handlers can be minimized by removing the manatee completely from the water. This can be accomplished by draining the water from the pool and stranding the manatee on the pool bottom, or by placing the manatee in a stretcher in low water and moving it to a dry work area. While restrained, the manatee should be placed on thick, closed-cell foam that is at least the length and about twice the width of the animal. At least four or five experienced people are required to restrain an adult manatee during this procedure. Manatee calves can be restrained and supported in the water by a single person. However, calves must be placed in a stretcher before removal from the water, because one person cannot safely lift a manatee calf. Most diagnostic and therapeutic procedures can be accomplished using these stranding methods. Additionally, when an animal is in a stretcher, its body weight can be determined using a mechanically assisted hoist, crane, or block-andtackle system. If a manatee becomes too fractious during restraint, a restraint board and/or sedation should be considered. While out of the water, manatees should be sprayed with water to prevent skin desiccation and overheating. Rescued manatees are generally secured in a stretcher and transported in an enclosed truck on thick, 15- to 20-cm (6- to 8-in.), open- or closed-cell foam pads or in specially designed
0839_frame_C43.fm Page 945 Tuesday, May 22, 2001 2:08 PM
945
Manatees
TABLE 1 Commonly Used Drugs in the Florida Manatee Drug
Dose
Frequency
Route
SID BID SID BID BID BID TID
IM IM/PO IM PO IM PO PO
BID
PO
TID
PO
Once Once Once
PO PO PO
See text
IM
See text See text
IM IM
Antibiotics Ceftriaxone Amikacin Penicillin (G and benzathine) Metronidazole Tetracycline a Sulfasalazine a Gentamicin
22 mg/kg 7 mg/kg 22,000 U/kg 7 mg/kg 55 mg/kg 10 mg/kg 2.5 mg/kg Antifungal Drugs 2.5 mg/kg
Itraconazole
Gastrointestinal Drugs ®
Pepto-Bismol
30 ml/animal b
Parasiticidal Drugs Fenbendazole Ivermectin Praziquantel
10 mg/kg 200 µg/kg 8–16 mg/kg Sedative Drugs
Midazolam hydrochloride Diazepam hydrochloride Meperidine hydrochloride
c
0.045 mg/kg d 0.080 mg/kg c 0.066 mg/kg e 0.5–1.0 mg/kg
a
For calf enterocolitis only. Florida manatees that are destined for release to a free-ranging state are not routinely deparasitized unless complications associated with parasitism (e.g., diarrhea, etc.) are present. c For tranquilization with effects lasting 60 to 90 min; may be reversed with flumazenil (IM) on an equal-volume basis. d To facilitate intubation for general anesthesia; may be reversed with flumazenil (IM) on an equal-volume basis. e To provide sedation/analgesia for minor surgical procedures. Key: SID = once a day; BID = twice a day; TID = three times a day; IM = intramuscularly; PO = per os; orally. b
and padded transport boxes. Cranes and forklifts are generally needed to lift adults. During transport, the manatee should be kept moist and shaded. Ambient air temperature should be kept at 22 to 26°C (72 to 79°F). Manatees have been transported for 16 hours in this manner with no apparent ill effects beyond transitory increases in serum lactate dehydrogenase and creatinine phosphokinase (see Chapter 39, Transport). Chemical restraint can also be safely used in manatees. Generally, chemical restraint is used to calm fractious, healthy, adult manatees. Midazolam hydrochloride (0.045 mg/kg IM) and diazepam (0.066 mg/kg IM) have been used for manatee sedation (Table 1) (Walsh and Bossart, 1999). With these drugs, movement and respiration are decreased, and the manatee may appear asleep, although it will respond to stimuli and have a palpebral reflex.
0839_frame_C43.fm Page 946 Tuesday, May 22, 2001 2:08 PM
946
CRC Handbook of Marine Mammal Medicine
FIGURE 2 Thoracic auscultation in a cold stress syndrome manatee. (Photo credit: G. Bossart.)
Physical Examination As with domestic animal species, the most important initial clinical tool for formulating a differential diagnosis is the physical examination. The approach is similar to that for terrestrial species with a few notable exceptions. If possible, a history should be obtained. The manatee should be visualized in the water to determine position, attitude, and swimming and diving capabilities. Breathing rate and breathing excursion characteristics should be determined, and an overall subjective assessment of nutritional status should be made. The normal respiratory rate is variable, but three to four breaths per 5-min period is typical. The breath excursion is relatively slow compared with cetaceans. The breath excursion should be forceful, constant, and slightly crisp sounding, with no rasping sounds. The breath in a healthy manatee should have a rumen-like “sweet” odor. Malnourished manatees typically have a distinct neck and caudal peduncle with a “sunken-in” appearance of the ventral abdomen. The “hands-on” physical examination should include body weight determination and thorough inspection of the oral cavity for foreign bodies and digital evaluation of the molars. Oral mucous membrane color and capillary refill time are similar to that in domestic mammals. Auscultation and percussion can be utilized to evaluate the lungs, heart, and gastrointestinal tract (Figure 2). Because of the breath-holding nature of the manatee, patience is required for thoracic auscultation. The hindgut fermentation process produces abundant colonic gas. This gas and peristaltic gut sounds are normal findings on abdominal auscultation and percussion in a healthy manatee. For anatomical landmarks for auscultation, the reader is referred to Chapter 9 (Anatomy). Traumatic injuries should be characterized regarding the extent of the injury and the presence of musculoskeletal or neurological compromise. The physical examination should also include digital rectal palpation to determine the presence of constipation, diarrhea, or parasites. The remaining physical examination closely follows the procedures used for domestic animals.
Diagnostic Techniques Blood samples are taken from the medial interosseous space of the radius and ulna (Figure 3). This site constitutes the brachial vascular bundle. Blood vessels cannot be visualized; hence, practice is required to become proficient at venipuncture. The middle portion of the medial surface of the pectoral flipper is surgically scrubbed with a commercially available iodine
0839_frame_C43.fm Page 947 Tuesday, May 22, 2001 2:08 PM
Manatees
947
FIGURE 3 Blood is collected from the brachial vascular bundle following a 3-min surgical scrub. (Photo credit: G. Bossart.)
surgical solution for at least 3 min prior to venipuncture. The flipper is firmly restrained while the manatee is stranded, and an 18- to 20-gauge, 1- to 1.5-in. needle (with attached syringe or butterfly set) is inserted between the palpable medial edges of the radius and ulna (see Chapter 19, Clinical Pathology, for venipuncture sites). The vessels of the brachial vascular bundle are arranged in a small plexus, and are difficult to isolate for catheterization. Laboratory tests performed immediately should include a spun hematocrit, plasma protein (via refractometer), and blood glucose (via a reagent test strip or point-of-care glucose analyzer) determinations. The last is particularly important for orphaned manatee calves and cold stress syndrome cases. Blood testing should include a complete blood count (CBC), a standard panel of serum analytes, and serum protein electrophoresis (see Chapter 19, Clinical Pathology). Specialized clinicopathological tests may include aspirate and exudate diagnostic cytology, fecal/nasal parasite examinations, urinalysis, serological testing for infectious agents (Duignan et al., 1995; Geraci et al., 1999), and testing for biotoxins (Bossart et al., 1998b). Imaging techniques that can be utilized in manatees include radiography, ultrasonography, endoscopy, thermography, and magnetic resonance imaging (MRI) (see Chapters 24 through 28). Each technique has limitations that are primarily related to the large size and unique anatomy of manatees. For example, radiographic interpretation of the thorax and abdomen can be difficult, because of the overlap of the respiratory and alimentary tracts in this species. MRI studies are generally limited to manatee calves because of the size restrictions of MRI equipment. Complete ultrasound examinations are often limited by the normally large amount of intestinal gas. However, ultrasonography has been used successfully for subcutaneous and abdominal abscess evaluation, kidney and urinary bladder examination, and echocardiograms. Ultrasonography is particularly useful for pregnancy diagnosis and should be used when pregnancy is suspected (e.g., abdominal distension, enlarged and swollen vulva, and elevated serum progesterone). Gastric and colonic endoscopy can be used but may be limited by the slow gut transit time of approximately 7 to 10 days. Urine collection by catheterization is difficult because of the small, tight urogenital opening, the distance of the urinary opening from the surface, and animal resistance to the procedure. One can have limited success by manually stimulating urine flow; this is done by applying pressure on the abdomen anterior to the vulva in females or posterior to the genital opening in males. Patience is required. Urine collection from captive manatees can be successfully accomplished using animals trained to urinate on command (Manire, pers. comm.).
0839_frame_C43.fm Page 948 Tuesday, May 22, 2001 2:08 PM
948
CRC Handbook of Marine Mammal Medicine
Electrocardiography (ECG) has been performed successfully in manatees of various ages. Standard domestic animal methodology is used. Additionally, commercially available, battery-powered, portable ECG units with built-in electrodes designed for humans have been successfully used in research studies. Electrocardiogram abnormalities have not been reported in manatees.
Therapeutics Oral medication is best administered by stomach tube. Some manatees may take oral medication concealed in food such as bananas or monkey chow; however, the natural crushing action of the molars usually results in drug loss. In most instances, oral antibiotic therapy is not recommended, because it can result in loss of normal enterocolic flora, diarrhea, and hypermotility. Parenteral drug administration is recommended in most instances. Intravenous drugs can be administered via the brachial vascular bundle. However, this site is difficult to catheterize for long-term intravenous drug therapy, and care must be taken to avoid inadvertent arteriolar injection. Intramuscular drug administration is the preferred route in manatees. Intramuscular injection sites include the caudal epaxial muscles and shoulder muscles. The injection site is disinfected with a commercially available iodine surgical solution for at least 3 min to minimize iatrogenic contamination of internal tissues. Needles recommended for intramuscular injection in adults are 2 to 3.5 in. and 18 to 20 gauge. Needles for calves are 1 to 1.5 in. and 20 to 22 gauge. Common antibiotics and other medications are listed in Table 1. Combination antibiotic use, such as penicillin or ceftriaxone combined with amikacin sulfate, is used frequently with life-threatening bacterial infections. Pharmacokinetic studies have not been conducted in manatees; therefore, drug dosages are generally based on doses recommended for domestic animals or humans. Pharmacokinetic studies are needed to guide drug dosages safely in this species. Stomach intubation is recommended for fluid therapy and nutritional supplementation (Figure 4), using a foal or small equine size soft plastic stomach tube by either the nasal or oral route. The length of the inserted tube is predetermined by measuring from the outer upper lip to the level of the caudal tip of a “tucked” pectoral flipper. At this point, the tube is externally marked with tape or a felt-tip pen, which then designates the point of maximum tube insertion. The tube should then be thoroughly lubricated with petroleum jelly. A bite block can be used when using the oral route. Intubation should be done by patient and experienced personnel to avoid inadvertent airway intubation and damage to the easily traumatized caudal oropharynx. Nasogastric intubation is well tolerated by most manatees and prevents the tube from being chewed, potential tube leakage, and/or tube fractures. Rehydration is best achieved by gastric intubation. Precise fluid therapy formulas have not been determined for this species. However, domestic animal fluid therapy guidelines can be used with a few notes of caution. First, extremely ill or dehydrated manatees typically have alimentary tract stasis. Therefore, these manatees should be given only water initially, and only about 50 to 75% of the calculated total fluid volume should be given to minimize the chance of fluid reflux. A 350-kg (770-lb) manatee may require up to 2.5 l of water twice daily to improve its hydration status. However, for the first two to three treatments, only 1.0 to 1.5 l of water should be given and then this volume slowly increased. Severely dehydrated or malnourished manatees may be tube-fed three times a day. As with fluid therapy in domestic animals, hematocrit and electrolytes should be closely monitored, and the fluids should be administered by gravity feed only. Inappetent or malnourished manatees can also be given nutritional supplementation by gastric tube. Manatees that have sustained human-related traumatic injury and cold stress
0839_frame_C43.fm Page 949 Tuesday, May 22, 2001 2:08 PM
Manatees
949
FIGURE 4 Orogastric intubation for fluid therapy and nutritional supplementation. Nasogastric intubation is an alternative route. (Photo credit: G. Bossart.)
syndrome often develop life-threatening dehydration and gastrointestinal stasis, which, if prolonged, results in cachexia. Reestablishment of gut function and motility is critical for a successful outcome. This can be accomplished by tube feeding the manatee a gruel composed of commercial primate pellets, lettuce, spinach, and water two to three times a day in volumes similar to those for rehydration. The gruel should be thin initially to allow the gastrointestinal tract to adapt to the presence and type of digesta. The gruel should be thickened, gradually, when flatulence and fecal production are noted. The presence of flatulence is generally a favorable prognostic indicator. If there is excessive reflux of gruel into the tube, the gruel volume should be reduced. However, a small amount of gruel reflux from the previous feeding is normal. As gut function improves, manatees will start to graze on offered vegetation, but tube feedings should not be discontinued until appetite and daily food intake are normal. Cessation of gruel feedings during the recovery process is a common mistake. Prolonged tube-feeding nutritional support is not uncommon. One manatee at the Miami Seaquarium with severe boat-related, traumatic injuries required tube feeding twice a day for 6 months. Constipation is often a problem in the initial attempts to reestablish gut function. Mineral oil may be administered for constipation via a stomach tube at 2 ml/kg up to 1 l total volume. It is important to keep in mind that normal gut transit time may be as long as 10 days, so three or four doses of oil may be required. Enemas are particularly helpful in removing dried and impacted fecal material. Warm freshwater enemas for constipation are recommended for 2 to 3 days in adult manatees. Approximately 3 to 5 l of water can be used per treatment. Mineral oil can also be added in small amounts. Saline enemas should be used if treatment longer than 3 days duration is necessary. Extremely thin, comatose, or abnormally buoyant (e.g., pneumothorax cases) manatees may require support with a flotation device made from diver wet suit (neoprene) material (Walsh and Bossart, 1999) or a human water-flotation vest (Murphy, pers. comm.). Flotation devices
0839_frame_C43.fm Page 950 Tuesday, May 22, 2001 2:08 PM
950
CRC Handbook of Marine Mammal Medicine
FIGURE 5 Isoflurane inhalation anesthesia in a manatee. Note intranasal route of tracheal intubation. (Photo credit: G. Bossart.)
often help injured or sick manatees regain a normal appetite. However, some manatees resent these devices and may recover faster without their application.
Anesthesia Midazolam or diazepam combined with meperidine hydrochloride has been used for minor surgical procedures such as removing bony sequestra from vertebral injuries, and removing deeply embedded and entangled monofilament fishing line from pectoral flippers (see Table 1). Drug onset time is approximately 15 to 20 min. Flumazenil administered IM on an equalvolume basis can be used to reverse the effects of diazepam and midazolam after procedure completion. Naloxone hydrochloride given IM can be used to reverse the effects of meperidine hydrochloride. Reversal is indicated if the manatee shows signs of disorientation or incoordination. Following any anesthetic protocol, it is recommended that the manatee be maintained in water at a depth of less than 1 m (2 to 3 ft) until recovery is complete (i.e., until normal swimming, diving, breathing, and feeding behaviors are present). Inhalation anesthesia with isoflurane can be used in manatees (see Chapter 29, Anesthesia) (Figure 5). Midazolam (0.08 mg/kg IM) and an isoflurane reservoir system consisting of a 5gal plastic water jug head mask with the bottom removed (cut out of the jug) are used for preparation for intubation. The head mask should cover the head and be form fitting to prevent gas from escaping. Because of the manatee’s unique elongated soft palate and small oral cavity, tracheal intubation is achieved intranasally with the required visual assistance of an endoscope. The endoscope is inserted through one nostril until the larynx is observed and an elongated foal endotracheal tube with stylet is then inserted through the other nostril and visually guided to insertion. The trematode Cochleotrema cochleotrema may interfere with nasal intubation, but the endotracheal tube will usually dislodge these parasites, facilitating intubation. Isoflurane maintenance levels are similar to those in domestic animals. The use of a ventilator is recommended as spontaneous breathing typically results in hypoxia and/or hypercapnia. Additionally, pulse oximeter monitoring and determination of end-tidal carbon dioxide levels and blood gases are also recommended. Endotracheal tubes generally require frequent suctioning during the course of the procedure. Following the procedure, the midazolam or diazepam can be reversed with flumazenil. Postanesthesia water depth restrictions should be followed as described above.
0839_frame_C43.fm Page 951 Tuesday, May 22, 2001 2:08 PM
Manatees
951
Environmental Diseases Brevetoxicosis Red tide–associated mortality may represent an emerging problem for the Florida manatee (see Chapter 2, Emerging and Resurging Diseases). Red tides are composed of dinoflagellates that produce potent neurotoxins. The dinoflagellate Gymnodinium breve produces potent neurotoxins known as brevetoxins. During a 2-month period in the spring of 1996, at least 150 manatees died along the west coast of Florida (U.S. Marine Mammal Commission, 1996). A novel immunohistochemical technique developed at the University of Miami School of Medicine (Florida) showed that brevetoxicosis was a primary component of this unprecedented epizootic (Bossart et al., 1998b). This permitted, for the first time, the determination of the presence, abundance, and distribution of brevetoxins in tissues. Results indicated that brevetoxins were likely inhaled and disseminated widely throughout the body in lymphoid tissues. The hemotoxic and possible long-term effects on the immune system were also demonstrated. Utilization of this methodology in a retrospective study indicated that a manatee epizootic in 1982 was also caused by brevetoxicosis. Toxicosis in these cases was likely a result of incidental ingestion of toxin-containing ascidians (tunicates). Neurotoxic signs of clinical brevetoxicosis reported from SeaWorld (in Orlando, Florida) and the Lowry Park Zoo (in Tampa, Florida) include seizure, disorientation, incoordination, hyperflexion, muscle fasciculations, flacid paralysis, and dyspnea (Murphy, pers. comm.; Walsh and Bossart, 1999). Murphy also reported a type of consumption coagulopathy in manatees from this epizootic and from a smaller epizootic in 2000. Manatees with clinical signs of brevetoxicosis are treated symptomatically with steroids and nonsteroidal anti-inflammatory drugs. Supportive care includes providing fluids, nutritional supplementation, and water buoyancy devices to prevent drowning (see Therapeutics, above). In many instances, the manatees survived following this treatment regimen and were eventually released.
Cold Stress Syndrome Water temperatures below 20°C (68°F) for extended periods initiate a cascade of clinical signs and disease processes that constitute the manatee cold stress syndrome. Adult manatees appear to handle the effects of cold temperatures better than juveniles or calves. This may be due to surface-to-volume body relationships and a nutritional plane resulting in lowered capacity for heat production. The more acute lethal effects of cold temperature probably involve lethargy, anorexia, and terminal hypothermia through metabolic drains to the environment (Buergelt et al., 1984). The chronic lethal effects of cold likely trigger a cascade of physiological changes that predispose these animals to various opportunistic pathogens involving multiple organ systems (Bossart, 1995). Infectious bronchopneumonia, generalized infectious dermatitis, and enterocolitis are common sequelae of the chronic cold stress syndrome. These conditions may require months of pathogen-specific and supportive therapy for favorable outcomes. Bacteria that have been isolated from these disease conditions include Staphylococcus aureus, Morganella morganii, Edwardsiella tarda, Aeromonas hydrophila, and various species of Pseudomonas, Vibrio, and Clostridium. Additionally, secondary fungal infections of the skin (especially Mucor spp.) and lungs can occur as this syndrome progresses. Skin lesions may become severe and generalized with ulceration resembling a toxic contact dermatitis (Walsh and Bossart, 1999). Treatment is based on culture and sensitivity results, and appropriate parenteral antibiotic therapy. Skin lesions should be treated with daily povidone–iodine scrubs, and special attention should be directed to maintaining a clean and bacteria-free water environment.
0839_frame_C43.fm Page 952 Tuesday, May 22, 2001 2:08 PM
952
CRC Handbook of Marine Mammal Medicine
Supportive therapy is of paramount importance. Nonspecific clinical signs include shivering, cachexia, anorexia, constipation, and absence of gut sounds and flatulence. Hematological and serum analyte findings may include dehydration, leukocytosis, and elevated lactate dehydrogenase (LDH), creatinite kinase (CPK), and creatinine concentrations. Regaining normal gut function and treating initial dehydration involves gastric intubation with water only. This is followed by nutritional support via gastric intubation of a gruel mixture that is gradually increased in consistency and volume (see Therapeutics, above). Gruel feeding may be done twice daily as long as feces are produced. Constipation, characterized by absent or hard, firm feces, can be treated with oral mineral oil and warm-water enemas.
Infectious Diseases Juvenile and adult manatees may be unusually prone to chronic abscessation, which is apparently not secondary to trauma. Abscesses may be dermal to subcutaneous, and relatively superficial, or lying deep within skeletal muscle fascial planes. Staphylococcus aureus is usually isolated from these lesions. Treatment typically involves surgical drainage and daily flushes with a dilute hydrogen peroxide and povidone–iodine solution, often followed by an antibiotic and/or proteolytic enzyme solution. Surgical drains or Foley catheters can be used to prevent premature closure of the skin. The pathogenesis of this predisposition is unknown. Fatal septic metritis secondary to dystocia and subsequent fetal maceration have been reported in three manatees (Walsh and Bossart, 1999). In three other cases of dystocia, calves were manually extracted (two cases) or delivered by cesarean section with favorable outcomes. Lesions consistent with brucellosis have not been reported in manatees, although antibodies to Brucella spp. have recently been reported in 5 of 71 Florida manatees (Geraci et al., 1999). Other infectious diseases reported in adult manatees include systemic mycobacteriosis due to Mycobacterium marinum (Morales et al., 1985) and M. chelonei (Boever et al., 1976), mycotic dermatitis (Dilbone, 1965; Tabuchi et al., 1974), hemorrhagic enteritis (O’Shea et al., 1985), pleuritis, lung abscessation, and various nonspecific dermatopathies (Bossart, unpubl. data; Walsh and Bossart, 1999). The first virus to cause clinical disease in manatees was only recently described (Bossart et al., 1998a). Cutaneous papillomatosis was documented in a group of captive manatees (Lowe, pers. comm.). Virions consistent with a papillomavirus were found with transmission electron microscopy in characteristic verruciform cutaneous lesions. Positive immunohistochemical staining was demonstrated using a polyclonal antibody developed from bovine papillomavirus Type 1 (Bossart et al., 1998a). Polymerase chain reaction analysis of these tissues demonstrated a virus consistent with Type 1 bovine papilloma virus (Lipscomb, pers. comm.). Serological evidence of morbillivirus has been demonstrated in manatees (Duignan et al., 1995). Additionally, serological evidence of pseudorabies, San Miguel sea lion virus type 1, eastern, western, and Venezuelan equine encephalitis has recently been reported in Florida manatees (Geraci et al., 1999). However, signs of clinical disease or active infection due to these viruses have not been described. Therefore, these data likely reflect exposure and an immunological response to these pathogens without accompanying clinical disease. Additional investigations are needed to associate these test results with natural history and clinical and pathological data.
Parasites A few species of endoparasites are commonly found in manatees; however, pathological signs or clinical disease are rarely associated with these parasites (Buergelt et al., 1984; Beck and Forrester, 1988) (see Chapter 18, Parasitic Diseases).
0839_frame_C43.fm Page 953 Tuesday, May 22, 2001 2:08 PM
Manatees
953
The nasopharyngeal trematode Cochleotrema cochleotrema can cause signs of chronic rhinitis and make nasogastric or endotracheal intubation difficult. A trematode (likely Nudacotyle undicola) has been associated with a severe enterocolitis (Walsh and Bossart, 1999). Toxoplasmosis due to Toxoplasma gondii has been reported to cause fatal encephalitis (Buergelt and Bonde, 1983) and myocarditis (Bossart, unpubl. obs.) in Florida and Antillean manatees, respectively. Dermal parasites described in manatees include copepoda (Harpacticus pulex) (Humes, 1964), cirripedia (Chelonibia manatii), and some nematode species (Walsh and Bossart, 1999). These parasites likely cause opportunistic infestations rather than being primary pathogens. Manatees scheduled for release are not routinely dewormed, unless clinical disease associated with parasitism is present. Commonly used parasiticidal drugs are listed in Table 1. No controlled studies to determine the safety, efficacy, or dosage of parasiticidal drugs in this species have been reported.
Miscellaneous Conditions Neoplasia Neoplasia in manatees is rare. Benign viral papillomas have recently been reported (Bossart et al., 1998a). Disseminated malignant lymphoma has been described in one manatee (Murphy, pers. comm.; Bossart, unpubl. data).
Neonatal Disease Orphaned neonatal manatees comprise a large percentage of cases presented to manatee rehabilitation facilities in the United States, Mexico, Belize, Colombia, and Brazil (Bossart and Menchaca, 1998). In the United States, there are a number of possible causes of premature maternal separation, including death of the cow from human-related or other factors, and/or recruitment of inexperienced cows into the breeding population, because of a loss of experienced breeders. Compared with adult manatees, neonates can develop severe medical problems that are labor-intensive and expensive to treat. Congenital disease is rarely reported, although congenital malformations of the pectoral flipper (Watson and Bonde, 1986; Walsh and Bossart, 1999) and umbilical hernias (Walsh and Bossart, 1999) have been reported in manatee neonates. Orphaned neonates are usually presented to rehabilitation facilities in a critical medical state. Orphans typically are cachectic, hypothermic, and in a state of metabolic exhaustion. Blood studies can indicate severe life-threatening hypoglycemia, hypernatremia, hypoproteinemia, hypoalbuminemia, and hypogammaglobulinemia. Orphaned manatees have also been described with Gram-negative bacterial omphalitis and secondary peritonitis (Walsh et al., 1987). Orphaned manatees can develop enterocolitis, which is also life-threatening. Pneumotosis intestinalis has been reported as a sequela to enterocolitis (Walsh et al., 1999). The etiology of the enterocolonic inflammation is likely multifactorial involving dietary (e.g., artificial formulas), infectious (e.g., Pseudomonas aeruginosa, Salmonella spp., Clostridium difficile), and immunological factors. Critical care treatment is based on clinicopathological findings. Environmental treatment should involve providing an ultraclean water habitat with water temperature maintained around 29°C (84°F). Elemental tube-fed diets consisting of Criticare HN (Criticare HN, Mead Johnson and Co., Evansville, IN), Nutramigen (Nutramigen, Mead Johnson and Co., Evansville, IN), and medium-chain triglycerides have been used successfully (Walsh et al., 1999).
0839_frame_C43.fm Page 954 Tuesday, May 22, 2001 2:08 PM
954
CRC Handbook of Marine Mammal Medicine
One notable exception to oral antibiotic use occurs with calf enterocolitis. In this case, the recommended treatment typically involves combination oral antibiotic therapy in an attempt to “sterilize” the alimentary tract. Combination therapy includes oral sulfasalazine, gentamycin, or amikacin, and metronidazole with bismuth salicylate (see Table 1) given by stomach tube for 10 days. This treatment is combined with oral fluid and nutritional therapy as needed and followed by “reseeding” the alimentary tract with Lactobacillus and fecal material from a healthy manatee.
Human-Related Traumatic Injuries A well-organized manatee carcass salvage program has existed in Florida since the early 1970s, in an effort to identify and quantify mortality factors. Necropsies are conducted at the Marine Mammal Pathobiology Laboratory in St. Petersburg, Florida. Human-related activities seriously threaten the future of this species. Up to 33% of annual manatee deaths, for which a cause of death can be determined, are directly related to human activities (O’Shea et al., 1985; U.S. Marine Mammal Commission, 1999). Deaths that are indirectly related to human activities account for more than 60% of the human-related category of manatee mortalities annually (Bossart, 1999). Boat trauma consists of sharp (propeller) and blunt (hull strike) injury subcategories. Blunt trauma also can result from the effects of injuries due to floodgate crushing. Sharp trauma is usually the result of a boat propeller injury. Propeller wounds have a wide variety of characteristics and may be focal to multifocally extensive and range from mild to severe. The extent and severity of the lesion(s) are related to propeller size and acceleration. Anatomical characteristics and the manatee’s normal position in the water column predispose propeller injuries to the head, dorsal thorax, caudal-dorsal peduncle, and dorsal fluke. Dorsal thoracic injury may penetrate the pleural cavity due to the long horizontal axis of the lungs. If the wound is of sufficient depth, exteriorization of the lung may result (Figure 6).
FIGURE 6 Boat propeller sharp trauma with resultant ante-mortem lung exteriorization. (Photo credit: G. Bossart.)
0839_frame_C43.fm Page 955 Tuesday, May 22, 2001 2:08 PM
Manatees
955
Boat propeller wounds that penetrate the pleural cavity are usually fatal due to compromised pulmonary function, secondary bacterial infection, and sepsis. Head trauma may also be lethal depending on the damage to sensory, musculoskeletal, or central nervous system functions. Propeller laceration of the caudal-dorsal peduncle with subsequent subcutaneous and skeletal muscle loss has a better prognosis for survival. However, these injuries may result in permanent neuromuscular compromise, vertebral osteomyelitis, and/or chronic fistulous tracts that preclude complete rehabilitation and release. The medical treatment protocols for propeller wounds involve parenteral antibiotics based on bacterial culture and sensitivities, wound care management, and nutritional support. Ceftriaxone, or procaine penicillin combined with amikacin, can be administered prior to receiving results of hematological, serum chemistry, and culture submissions (see Table 1). Wound care is complicated by the aquatic environment. Pool water must be kept as clean as possible with special attention to a high water turnover rate and water treatment to minimize the growth of microorganisms (e.g., addition of chlorine to approximately 1 ppm, ultraviolet sterilization). A standard surgical povidone–iodine solution diluted with saline is applied to the wound two to four times a day. The wound should be thoroughly scrubbed and flushed with the iodine solution. A dental pick or nozzled hose helps with deep wound flushing and debridement. The strength of the iodine solution used is indirectly proportional to solution contact time available (i.e., higher iodine concentrations are used with a shorter available contact time). Petroleum jelly–based antibiotic ointments and proteolytic enzyme preparations can be used as a postiodine treatment. Wounds can be covered with custom-made wet suit body wraps or left open to facilitate a granulation tissue response. With rare exceptions, all wounds are left to heal by second intention in manatees, and a crude prognostic indicator is the rapidity and extent of the granulation tissue response. A more favorable prognosis is given to manatees that exhibit a faster and proliferative granulation tissue response. However, the manatee granulation response may become unusually exuberant, resulting in premature wound closure. This can lead to the formation of bony sequestra and fistulous tracts. In these cases, debridement must be performed daily to prevent premature wound closure. These manatees also require intensive nutritional support. A juvenile female manatee with severe propeller injury to the caudal-dorsal peduncle required tube feeding twice a day for almost 6 months at the Miami Seaquarium (Figure 7). Tube feeding should not be stopped until normal grazing behavior, daily food consumption, and fecal production have occurred. One of the most severe nonlethal boat propeller injuries observed at the Miami Seaquarium involved an adult male manatee (Dougherty et al., 2000a). A propeller blade estimated to be 2 m (6 ft) in length caused a complete thoracolumbar fracture, cranioventral thoracolumbar dislocation, and complete spinal cord transection. The propeller trauma also resulted in the loss of significant amounts of epaxial skeletal muscle, subcutis, and skin. The tail fluke was immobile, but anal sphincter tone and penile sensation were present. Blood parameters indicated acute renal failure with profound myoglobulinemia (3630 µ g/l). Interactive and social behaviors were exhibited with other manatees. A favorable response to the acute renal failure was seen with intensive oral fluid administration and furosemide (1 mg/kg IV). Fibrin glue (Hemaseel, Fibrin Glue, Haemacure, Inc., Sarasota, FL) was used to close the spinal canal following infusion with 750 mg amikacin sulfate diluted in saline. Ceftriaxone was given parenterally (5 mg/kg IM, once daily). To reduce and stabilize the fracture and dislocation, the manatee was sedated with midazolam (0.044 mg/kg IM) and meperidine (0.5 mg/kg IM). Epidural anesthesia was provided by infusing 1% xylocaine via a central venous catheter placed in the spinal canal approximately 20 cm (∼8 in.) cranial to the fracture. A partial corpectomy of the first lumbar vertebra was performed, because it was displaced beneath the last thoracic vertebra. Stainless-steel pedicle screws were bilaterally placed into T18, L2, and L3. Stainless
0839_frame_C43.fm Page 956 Tuesday, May 22, 2001 2:08 PM
956
CRC Handbook of Marine Mammal Medicine
FIGURE 7 A juvenile female manatee with a severe propeller injury. This manatee survived following 2 years of medical and supportive care. (Photo credit: G. Bossart.)
FIGURE 8 Successful surgical reduction of a thoracolumbar fracture and dislocation using pedicle screws and stainless steel rods. This massive sharp trauma injury was due to a propeller blade estimated to be just under 3 m (6 ft) in length. (Photo credit: G. Bossart.)
FIGURE 9 Blunt boat hull trauma resulting in pneumothorax. This injury typically results in positive buoyancy with listing and/or difficulty in diving. (Photo credit: G. Bossart.)
0839_frame_C43.fm Page 957 Tuesday, May 22, 2001 2:08 PM
Manatees
957
steel rods were sequentially tightened to the pedicle screws to facilitate complete fracture reduction (Figure 8). Postsurgery, the manatee was bright, alert, and active. The fracture remained rigidly fixed and reduced. After 26 days, the manatee died from myocardial failure, thromboembolic disease, and bacterial sepsis. Blunt trauma from the impacts of boat hulls or floodgate crushing injuries can result in thoracic injury, including multiple rib fractures, vertebral fractures, lung perforation and torsion, pulmonary hemorrhage, and pneumothorax. External injury may be minimal. These manatees may exhibit positive buoyancy and are unable to dive or maintain a horizontal position in the water column (Figure 9). The traumatic injuries resulting from blunt trauma are directly related to hull mass and acceleration. Blunt trauma can often cause pneumothorax resulting in an inability of the animal to dive and listing to one side, with or without dyspnea. Lung torsion may develop and may be reduced by physically manipulating the manatee in a head-up, tail-down position to encourage the lung to a normal position (Walsh and Bossart, 1999). Use of buoyancy jackets or maintenance of a shallow water depth may provide breathing assistance. The surgical placement of chest tubes or drains for pneumothorax treatment is not recommended unless dyspnea is severe and life-threatening. This is based on experience from critical-care facilities at the Miami Seaquarium and Sea World. The manatee lung is normally under constant positive pressure; hence, the placement of drains results in the maintenance of the parenchymal rent and slows healing. The time for healing with pneumothorax without therapeutic drainage ranges from 4 to 18 months. Occasionally, the sites may be drained using a spinal needle and syringe with a threeway stopcock to evaluate the degree of healing. Common sequelae to blunt impact thoracic injury include focally extensive pyothorax, suppurative pleuritis, necrosuppurative pneumonia, rib and vertebral proliferative osteomyelitis, and subcutaneous emphysema. These sequelae may result in chronic debilitating disease, which precludes release. Treatment and diagnostic regimens (e.g., thoracic radiography, ultrasound, MRI) parallel those used in domestic animal medicine. Another common human-related traumatic injury is entanglement with monofilament fishing line, crab trap line, or discarded plastic loops used to bind parcels or boxes (Figure 10). Strangulation with amputation of the pectoral flipper commonly occurs. In some cases, monofilament line may entangle and penetrate deep to periosteal bony structures resulting in a proliferative, suppurative osteomyelitis. Therefore, radiographs should be taken in all cases of pectoral
FIGURE 10 Monofilament fishing line entanglement resulting in severe distal edema of the pectoral flipper. (Photo credit: G. Bossart.)
0839_frame_C43.fm Page 958 Tuesday, May 22, 2001 2:08 PM
958
CRC Handbook of Marine Mammal Medicine
flipper line entanglement. Entanglement may become so severe that amputation is required. Surgical amputation using a harmonic scalpel and electrocautery, followed by thorough wound irrigation with antibiotics and topical application of a cyanoacrylate spray (to act as a temporary water-resistant cover) has been used for one of these difficult cases (Dougherty et al., 2000b). Additionally, tape stirrups using standard veterinary tape and waterproof adhesive tape anchored with cyanoacrylate glue can be used (Walsh and Bossart, 1999). Another unusual form of entanglement was seen at the Miami Seaquarium. A rescued juvenile manatee had a discarded circular plastic band loop wrapped tightly around the midthorax. The manatee apparently swam through this loop as a calf and literally grew into it. This loop had penetrated through the skin to the rib periosteum circumferentially around the thorax caudal to the pectoral flippers. The band was cut and removed. The channel created by the loop was flushed daily with a dilute povidone–iodine solution for 8 weeks until the wound closed by second intention. Other types of human-related mortality factors include fishhook foreign bodies of the alimentary tract and lips, entrapment in culvert pipes and canals, and the destruction and degradation of habitat.
Acknowledgments The author thanks Ruth Francis-Floyd, Rose Manduca, and Charles Manire for taking the time to review this chapter.
References Bachman, K.C., and Irvine, A.B., 1979, Composition of milk from the Florida manatee, Trichechus manatus latirostris, Comp. Biochem. Physiol., 62: 873–878. Beck, C., and Forrester, D.J., 1988, Helminths of the Florida manatee, Trichechus manatus latirostris, with a discussion and summary of the parasites of sirenians, J. Parasitol., 74: 628–637. Bengston, J.L., 1983, Estimated food consumption of free-ranging manatees in Florida, J. Wildl. Manage., 42: 1186–1192. Best, R.C., 1981, Foods and feeding habits of wild and captive Sirenia, Mammal. Rev., 11: 3–29. Boever, W.J., Theon, C.O., and Wallach, J.D., 1976, Systemic Mycobacterium chelonei infection in a Natterer manatee, J. Am. Vet. Med. Assoc., 169: 927–929. Bossart, G.D., 1995, Immunocytes of the Atlantic Bottlenose Dolphin (Tursiops truncatus) and West Indian Manatee (Trichechus manatus latirostris): Morphologic Characterizations and Correlations between Healthy and Disease States under Free-Ranging and Captive Conditions, Ph.D. dissertation, Florida International University, Miami. Bossart, G.D., 1999, The Florida manatee: On the verge of extinction? J. Am. Vet. Med. Assoc., 214: 1178–1182. Bossart, G.D., and Bigger, C.H., 1994, Cellular components of the non-specific, humoral and cellmediated immune systems in the peripheral blood of the West Indian manatee (Trichechus manatus latirostris): Cellular population dynamics in health and disease, in Proceedings of the 1st International Manatee and Dugong Research Conference, Gainesville, FL, 148. Bossart, G.D., and Menchaca, M., 1998, Emerging manatee rehabilitation/reintroduction programs in Latin America, in Proceedings, First Captive Manatee Reintroduction/Release Workshop, Florida Marine Research Institute, St. Petersburg, FL, 10. Bossart, G.D., Ewing, R., Lowe, M., Murphy, D., and Sweat, M., 1998a, Cutaneous viral papillomatosis in manatees, in Proceedings, First Captive Manatee Reintroduction/Release Workshop, Florida Marine Research Institute, St. Petersburg, FL, 22.
0839_frame_C43.fm Page 959 Tuesday, May 22, 2001 2:08 PM
Manatees
959
Bossart, G.D., Baden, D.G., Ewing, R.Y., Roberts, B., and Wright, S.D., 1998b, Brevetoxicosis in manatees (Trichechus manatus latirostris) from the 1996 epizotic: Gross, histologic and immunohistochemical features, Toxicol. Pathol., 26: 276–282. Buergelt, C.D., and Bonde, R.K., 1983, Toxoplasmic meningoencephalitis in a West Indian manatee, J. Am. Vet. Med. Assoc., 185: 1294–1296. Buergelt, C.D., Bonde, R.K., Beck, C.A., and O’Shea, T.J., 1984, Pathologic findings in manatees in Florida, J. Am. Vet. Med. Assoc., 185: 1331–1334. Burn, D., 1986, The digestive strategy and efficiency of the West Indian manatee, Comp. Biochem. Physiol., 85: 139–142. Burn, D., and Odell, D., 1987, Volatile fatty acid concentrations in the digestive tract of the West Indian manatee, Comp. Biochem. Physiol., 88: 47–49. Crane, A., 1881, Notes on the habits of the manatees (Manatus australis) in captivity in the Brighton Aquarium, Proc. Zool. Soc. London, 30: 457–460. Dilbone, R.P., 1965, Mycosis in a manatee, J. Am. Vet. Med. Assoc., 147: 1095–1097. Domning, D.P., 1994, Paleontology and evolution of sirenians: Status of knowledge and research needs, in Proceedings of the 1st International Manatee and Dugong Research Conference, Gainesville, FL, 1–5. Dougherty, M., Bossart, G.D., Guest, J.D., Belanger, E., and Rommel, S., 2000a, Thoracolumbar fracture dislocation repair in a Florida manatee (Trichechus manatus latirostris) presented with a propeller wound, Abstr., in Proceedings of the Joint Meeting of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine Conference. Dougherty, M., Bossart, G.D., Guest, J.D., and Harris, D., 2000b, Entanglements in the Florida manatee (Trichechus manatus latirostris): A case report of amputation following multiple surgeries in a pregnant female, Abstr., in Proceedings of the Joint Meeting of the American Association of Zoo Veterinarians and the International Association for Aquatic Animal Medicine Conference. Duignan, P.J., House, C., Walsh, M.T., Bossart, G.D., Duffy, N., Fernandes, P.J., Rima, B.K., and Geraci, J.R., 1995, Morbillivirus infection in manatees, Mar. Mammal Sci., 11: 441–451. Etheridge, K., Rathbun, G.B., Powell, J.A., and Kochman, H.I., 1985, Consumption of aquatic plants by the West Indian manatee, J. Aquat. Manage., 23: 21–25. Florida Fish and Wildlife Conservation Commission, 2000, Tallahassee, Florida, press release. Geraci, J.R., and Lounsbury, V.J., 1993, Marine Mammals Ashore. A Field Guide to Strandings, Texas A&M Sea Grant College Program, Galveston, 305 pp. Geraci, J.R., Arnold, J., Schmitt, B.J., Walsh, M.T., Wright, S.D., Bossart, G.D., and Lounsbury, V.J., 1999, A serologic survey of manatees in Florida, in Marine Mammals Ashore, A Field Guide for Strandings, Geraci, J.R., and Lounsbury, V.J. (Eds.), Texas A&M Sea Grant Program, Galveston, 145–158. Gerstein, E.R., 1994, The manatee mind: Discrimination training for sensory perception testing of West Indian manatees (Trichechus manatus), Mar. Mammals, 1: 10–21. Hartman, D.S., 1979, Ecology and Behavior of the Manatee in Florida, the American Society of Mammalogists, Special Publication, No. 5, Pittsburgh, PA, 69: 95–120. Humes, A.G., 1964, Harpacticus pulex, a new species of copepod from the skin of a porpoise and a manatee in Florida, Bull. Mar. Sci. Gulf Caribb., 14: 517–520. Irvine, A.B., 1983, Manatee metabolism and its influence on distribution in Florida, Biol. Conserv., 25: 315–334. Lefebvre, L.W., O’Shea, T.J., and Rathbun, G.B., 1989, Distribution, status and biogeography of the West Indian manatee, in Biogeography of the West Indies: Past, Present, and Future, Woods, C.A. (Ed.), Sandhill Crane Press, Gainesville, FL, 567–610. Marmontel, M., Humphrey, S.R., and O’Shea, T.J., 1997, Population viability analysis of the Florida manatee (Trichechus manatus latirostris) 1976–1991, Conserv. Biol., 11: 467–481. Morales, P., Madin, S.H., and Hunter, A., 1985, Systemic Mycobacterium marinum infection in an Amazonian manatee, J. Am. Vet. Med. Assoc., 187: 1230–1232.
0839_frame_C43.fm Page 960 Tuesday, May 22, 2001 2:08 PM
960
CRC Handbook of Marine Mammal Medicine
Odell, D.K., 1978, Growth of a West Indian manatee, Trichechus manatus, born in captivity, in Proceedings, The West Indian Manatee in Florida, Brownell, R.L., and Ralls, K. (Eds.), Florida Audubon Society, Maitland, 131–140. Odell, D.K., 1982, The West Indian manatee Trichechus manatus Linnaeus, in Wild Mammals of North America, Chapman, J.A., and Feldhamer, G.A. (Eds.), Johns Hopkins University Press, Baltimore, MD, 828–837. O’Shea, T.J., 1995, Waterborne recreation and the Florida manatee, in Wildlife and Recreationists: Coexistence through Management and Research, Knight, R.I., and Gutzwiller, K.J. (Eds.), Island Press, Washington, D.C., 297–311. 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, Wildl. Manage., 49: 1–11. Powell, J.A., and Reynolds, J.E., 1999, The status of Florida’s manatees—A measured approach, Manatee News Q., 3: 1–2. Ray, C.E., and Domning, D.P., 1986, Manatees and genocide, Mar. Mammal Sci., 2: 77–78. Reep, R.I., Marshall, C.D., and Stoll, M.L., 1998, Distribution and innervation of the facial bristles and hairs in the Florida manatee (Trichechus manatus latirostris), Mar. Mammal Sci., 14: 257–273. Reynolds, J.E., and Rommel, S.A., 1996, Structure and function of the gastrointestinal tract of the Florida manatee, Trichechus manatus latirostris, Anat. Rec., 245: 539–558. Tabuchi, K., Muku, T., Satomicxhi, T., Hara, M., Imai, N., and Iwamoto, Y., 1974, A dermatosis in a manatee (Trichechus manatus): Mycological report of a case, Bull. Azabu Vet. Coll., 28: 127–129. True, F.W., 1884, The sirenians or sea cows, Fisheries and Fisheries Industries of the U.S., Sect. 1: 114–136. U.S. Fish and Wildlife Service, 1995, Florida Manatee Recovery Plan, 2nd rev., Atlanta, GA, 1–33. U.S. Marine Mammal Commission, 1996, Annual Report to Congress, Bethesda, MD, 6–14. U.S. Marine Mammal Commission, 1999, Annual Report to Congress, Bethesda, MD, 85–95. Walsh, M.T., and Bossart, G.D., 1999, Manatee medicine, in Zoo and Wildlife Medicine, Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 507–516. Walsh, M.T., Bossart, G.D., Young, W.G., and Rose, P.M., 1987, Omphalitis and peritonitis in a young West Indian manatee (Trichechus manatus latirostris), J. Wildl. Dis., 23: 702–704. Walsh, M.T., Murphy, D., and Innis, S.M., 1999, Pneumatosis intestinalis in orphan manatees (Trichechus manatus latirostris): Diagnosis, pathological findings and potential therapy, in Proceedings of the 30th Annual International Association for Aquatic Animal Medicine, 1. Watson, A.G., and Bonde, R.K., 1986, Congenital malformations of the flipper in three West Indian manatees, Trichechus manatus, and a proposed mechanism for development of ectrodactyly and cleft hand in mammals, Clin. Orthop., 202: 294–301. White, J.R., 1984, Man can save the manatee, Nat. Geogr., 166: 414–418. White, J.R., and Francis-Floyd, R., 1990, Manatee biology and medicine, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 601–623. Zeiler, W., 1978, The management of West Indian manatees (Trichechus manatus) at the Miami Seaquarium, in The West Indian Manatee in Florida, Brownell, R.L., and Ralls, K. (Eds.), Florida Audubon Society, Maitland, 103–108.
0839_frame_C44 Page 961 Tuesday, May 22, 2001 2:09 PM
44 Sea Otters Pamela Tuomi
Introduction The sea otter (Enhydra lutris) is a charismatic marine mammal that has received considerable attention from biologists, ecologists, physiologists, and the public (Rotterman and Simon-Jackson, 1988; Riedman and Estes, 1990; Brennan, 1996). Veterinary understanding of sea otters is comparatively limited, and derived from care of small numbers of sea otters held in display facilities, rehabilitation of stranded otters, care of otters exposed to the Exxon Valdez oil spill, and clinical support for researchers monitoring free-living otters. A number of novel diseases have recently been documented in sea otters (see Chapter 2, Emerging and Resurging Diseases; Chapter 18, Parasitic Diseases); however, their significance at the population level is still unknown. As the sea otter differs considerably from the other marine mammals discussed in this book, this chapter will include sections on its history, anatomy, and life history. Tagging and tracking of sea otters is described in Chapter 38 (Tagging and Tracking) and toxicology in Chapter 22 (Toxicology).
History The sea otter has always been limited to the coastal North Pacific. The original distribution extended from Hokkaido Island in northern Japan, through the Kuril Islands, along the Kamchatka Peninsula, the Commander Islands, the Aleutian Islands, the Alaska Peninsula and Kodiak Island, through Prince William Sound, south along the Pacific Coast of Canada and the United States, to about two thirds of the extent of the Baja California Peninsula of Mexico. The northern extent of the sea otter distribution is limited by sea ice. Colonization of the northern side of the Alaska Peninsula occurs, but is occasionally pushed back in years when the Bering Sea pack ice extends to the Alaska Peninsula (Schneider and Faro, 1975). There have been anecdotal reports of sea otters in the summer ranging as far north as Barrow, Alaska. The total population of sea otters prior to exploitation by modern humans was estimated to range from 150,000 (Kenyon, 1969) to 300,000 (Johnson, 1982) animals. Aboriginal hunting occurred throughout the sea otter’s range, and there is evidence from middens in the Aleutian Islands that such hunting was capable of severely depressing local populations of sea otters (Simenstad et al., 1978). Commercial hunting of sea otters was based on the value of their pelts, and began in the mid-18th century. Hunting ended in 1911, with the adoption of the International Fur Seal Treaty (see Chapter 33, Legislation), which extended protection to sea otters. The population may have been reduced to as few as 1000 to 2000 individuals remaining in 13 colonies, largely located along the eastern coast of Russia, the Aleutian Islands, southwestern Alaska, Kodiak
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
961
0839_frame_C44 Page 962 Tuesday, May 22, 2001 2:09 PM
962
CRC Handbook of Marine Mammal Medicine
Island, Prince William Sound, British Columbia, California, and Mexico (Ogden, 1941; Kenyon, 1969). The populations in British Columbia and Mexico eventually disappeared (Kenyon, 1969; Estes, 1980). Early in the 20th century, 32 sea otters were discovered near Point Sur, California (Bryant, 1915), and a group of 50 sea otters was reported off Bixby Creek (south of Monterey), California, in 1938 (Bolin, 1938). This small group is the basis for the present southern sea otter population. It is unlikely that the exact number of animals harvested during the exploitation period will ever be known accurately. Estimates of the numbers of sea otters killed from the mid1700s to 1911 range from a half million to a million (Lensink, 1960; 1962; Kenyon, 1969). With the halting of commercial hunting and the availability of previously unutilized prey resources, sea otter populations have proved to be amazingly resilient. Translocated sea otter populations in southeastern Alaska, British Columbia, and Washington State have increased in size at rates of 17 to 20% per year (Estes, 1990). However, the Californian sea otter population had a much slower rate of increase of 4 to 5% per year (Estes, 1990), with a 4% per year decline reported from 1995 to 1999 (Jameson, 1999). Sporadic reports of sea otters in Mexico suggest that recolonization of Baja California is possible (Gallo-Reynoso and Rathbun, 1997).
Classification The sea otter is the largest representative of the Family Mustelidae. Attempts to classify subspecies of sea otters have been based on geographic distribution or on morphological variables such as coat color or skull morphometrics. Using skull morphometrics and distinctions of pelage, Roest (1973) argued that California sea otters and Alaskan sea otters were the same species, and concluded that only two subspecies of sea otters exist: E. l. lutris, found from the Commander Islands through the Aleutian Islands and down the coast of North America, including California, and E. l. gracilis, found in the Kurile Islands and southern Kamchatka. However, his opinion was not universally accepted (Davis and Lidicker, 1975; Roest, 1976). Wilson et al. (1991) used univariate and multivariate statistical analyses of 20 skull measurements done on 304 skulls from adult sea otters to classify them as three subspecies: E. l. lutris from Japanese and Russian waters, E. l. kenyoni from the Aleutian Islands east and south to the Oregon coast, and E. l. nereis, found in California. Enhydra l. lutris skulls are large and wide, with short nasal bones. Enhydra l. nereis skulls are narrow with a long rostrum and small teeth, and lack a characteristic notch in the postorbital region found in skulls from the other two subspecies. Enhydra l. kenyoni skulls are intermediate in most characteristics, and have longer mandibles than either of the other two subspecies. The sea otter in California (E. l. nereis, called the “southern” sea otter) is listed as a threatened species (Greenwalt, 1979). Molecular genetic techniques have only recently been used to examine the classification of sea otters into subspecies. Cronin et al. (1996) used restriction enzyme analysis of polymerase chain reaction–amplified mitochondrial DNA to determine that there are two or three haplotypes of mitochondrial DNA and that the diversity of haplotypes is 0.1376 to 0.5854 in each population of otters. Enhydra l. nereis has monophyletic haplotypes of mitochondrial DNA, which are not shared with other populations, but E. l. lutris and E. l. kenyoni do not. Cronin et al. (1996) and Sanchez (1992) found four haplotypes of mitochondrial DNA in California sea otters and suggested that many haplotypes were preserved in sea otter populations despite the isolation that occurred following human intervention. Ralls et al. (1983) reached a similar conclusion by applying the concepts of population genetics to sea otters in California, and estimated that the present population retains about 77% of the genetic diversity that existed in the original population. They determined that a new population could be established by translocation of 50 breeding animals, and the present genetic diversity would be preserved in the new colony by introducing one or two individuals per generation.
0839_frame_C44 Page 963 Tuesday, May 22, 2001 2:09 PM
Sea Otters
963
Anatomy Sea otters have loose skin, very little body fat, and no scent glands. They are moderately sexually dimorphic. Males can exceed 45 kg (99 lb) and 148 cm (57 in.) in length and females can be 32.5 kg (71.5 lb) and 140 cm (54.6 in.) in length. Adult males are 34% heavier and 8% longer than females on average, and males tend to have a heavier head and neck than females (Kenyon, 1969; Estes, 1980). Gender can be distinguished visually only by the observation of two abdominal mammae or, more reliably, by the presence of a penile bulge, caused by the presence of a substantial baculum (Barabash-Nikiforov et al., 1947; Kenyon, 1969). Age estimates of wild sea otters have been based on observation of tagged individuals. Examination of incremental lines in the cementum of the first premolar tooth has also been used (Schneider, 1972; Garshelis, 1984; Bodkin et al., 1997). Life expectancy in the wild in Alaska appears to be about 15 to 20 years (Estes et al., 1996) but may exceed 20 years in captivity. Sea otters have 38 chromosomes. The dental formula is I 3/2, C 1/1, P 3/3, M 1/2; total 32. The sea otter is the only species of fissiped carnivore with two pairs of lower incisors. All other species of Lutrinae have three incisors on each lower jaw, and a total of 34 (Amblyonyx cinerea) or 36 teeth. The lower incisors protrude and are chisel-shaped to scrape meat from the shells of prey (Hildebrand, 1954). Molars and premolars (postcanine teeth) are strong and flattened for crushing hard-shelled prey (Kenyon, 1969; Reidman and Estes, 1988). Mature dentition is achieved in 1 year. Sea otter pups are born with 26 teeth erupted (I 3/2, C 1/1, P 3/3). The canines are wider in males than in females (Scheffer, 1951). The gender difference in canine width has been used to distinguish sex of beach-cast skulls. The vertebral column consists of 7 cervical, 14 thoracic, 6 lumbar, 3 sacral, and 20 to 21 caudal vertebrae. Great flexibility of the spine is permitted by reduction of the vertebral processes, shortening and heightening of the centra, and enlargement of the intervertebral foramina (Taylor, 1914). The absence of a clavicle may allow for greater flexibility of the pectoral girdle (Estes, 1980). Bones are sometimes dyed a pale violet due to absorption of polyhydroxynaphthoquinone from ingested purple sea urchins (Strongylocentrotus purpuratus). The nasal turbinates of sea otters are well developed, and olfactory sensitivity typical of terrestrial carnivores has apparently been retained. Scent recognition and sexual pheromones may play important roles in sea otter social behavior (Reidman and Estes, 1990). Increased turbinate surface may also allow for better prewarming of inspired air, and assist in reducing body heat loss during respiration. The thoracic cavity is large relative to the rest of the body, reflecting the importance of lung volume to the buoyancy of sea otters (Kooyman, 1973). Mean total lung capacity is 9 l, and, as a proportion of body weight, lung capacity of sea otters (345 ml/kg) is two to four times greater than pinnipeds (84 to 145 ml/kg) (Lenfant et al., 1970). The very large lung volume of sea otters comprises about 66% of its total oxygen store, compared with 16 to 43% in pinnipeds (Lenfant et al., 1970), which may reflect the behavior of sea otters as shallow and brief divers compared with other marine mammals. The right lung has four lobes, and the left lung has two (Tarasoff and Kooyman, 1973a). The trachea is incomplete dorsally with partially calcified rings. Development of the cartilaginous rings is incomplete at birth, and continues while the pup is in the nondiving phase of life (Drabek and Kooyman, 1984). Some cartilage-supported airways extend into the alveoli, to compensate for compression during diving (Tarasoff and Kooyman, 1973b), while others branch into nonsupported airways, which extend up to 1 mm before ending in clusters of alveoli (Denison and Kooyman, 1973; Kooyman, 1973). It is considered an intermediate form of reinforcement, falling between the structure of phocid lungs, whose small airways are reinforced by a slender tube of oblique muscle, and those of otariids, whose small airways are reinforced by much
0839_frame_C44 Page 964 Tuesday, May 22, 2001 2:09 PM
964
CRC Handbook of Marine Mammal Medicine
thicker pieces of cartilage (Denison and Kooyman, 1973; Kooyman, 1973). The reinforcement of terminal airways in both sea otters and in members of the Pinnipedia is a remarkable example of convergent evolution. Sea otters rely on pelage that is the densest of any mammal for thermoregulation. Hair 2 density ranges from 26,413 to 164,662 hairs/cm (Williams et al., 1992). This density of hair is twice that of northern fur seals (Callorhinus) and of the river otter (Lutra canadensis) and more than 60 times greater than that of haired seals (Tarasoff, 1974). The combination of extremely dense fur and sebaceous gland secretions creates a waterproof barrier that greatly reduces conductive heat loss to the water. The sebaceous secretion in the sea otter consists mainly of squalene (C30H50), a fatty acid precursor to cholesterol, which is found only in trace amounts in the fur of other mammals (Williams et al., 1992). Hair length varies from 30 mm for guard hairs to 12 mm for underfur hairs (Tarasoff, 1974). Guard hairs are relatively few in number, but may contribute to insulation by helping to support the underhairs through interlocking with the scales found at their base. In addition, they may overlap sufficiently to prevent disruption of the insulative underfur while swimming (Tarasoff, 1974). Guard hairs are of larger diameter on the abdomen (mean: 106 µm) than on the back (mean: 44 µm), which may be an adaptation to the sea otter’s preference for floating on its back. In contrast, the river otter (Lutra lutra), which swims on its ventrum, has larger guard hairs on its back (Barabash-Nikiforov et al., 1947). Undercoat hairs provide most of the insulation by interlocking to trap air. Both the guard hairs and the undercoat hairs are angled caudally in the skin (61.9° to 84.3°) and lack arrector pili muscles, which helps with streamlining during swimming and diving (Scheffer, 1964; Ling, 1970; Tarasoff, 1974; Williams et al., 1992). Smooth muscle bundles in the dermis may function as de facto arrector muscles, or they may assist in aeration of the pelage by local thickening of the skin (Williams et al., 1992). The cutaneous trunci muscle is well developed along the sides and dorsum of the sea otter, and its rapid contractions assist in pleating of the skin to help trap air in the fur (Tarasoff, 1974). Sea otters molt gradually throughout the year (Kenyon, 1969), although a peak molting period in spring was noted in captive Alaska sea otters. Although the sea otter does not rely on subcutaneous fat for insulation, such fat is present in an animal in good body condition. The weight of body fat in one sea otter was 1.8% of the total body weight, compared with 37.9% for the harp seal (Pagophilus groenlandicus), a marine mammal that uses body fat as its primary insulation (Tarasoff, 1974). During necropsy, measuring the depth of subcutaneous fat in the groin can be used to assess body condition. Normal animals have about 1 cm or more thickness of subcutaneous fat. In a starving animal, fat is quickly lost from the mesentery and around the kidney lobules. 2 2 The surface area of the sea otter averaged 10,856 cm (range: 5206 to 16,380 cm ) for 11 animals. The forefeet and hind feet areas are 0.8 and 5.8%, respectively, of the total surface area (Tarasoff, 1974). Because the palmar and plantar surfaces of the paws are not haired, they represent a potential avenue of thermal loss. The forepaws are mittenlike, with united pads and retractable claws. Sea otters can look behind themselves by arching their backs and rolling in the sagittal axis, while keeping their paws completely out of the water, a movement called highintensity rocking (Packard and Ribic, 1982). When slowly approached on the surface, a resting sea otter only reluctantly reintroduces its paws into the water. First, it tries to move out of the way by sculling with its tail. Then it may paddle using its hind flippers, before finally making a surface dive to escape. The fifth digit of the hind flipper is the longest digit, a characteristic not found in other mammals, which makes walking on land somewhat awkward. The hind digits are webbed to the tips of the toes and the toe pads are only slightly formed. In some animals, metatarsal pads can be faintly distinguished (Kenyon, 1969).
0839_frame_C44 Page 965 Tuesday, May 22, 2001 2:09 PM
Sea Otters
965
Vision Sea otters must be able to see well both above and below water, but have slightly poorer underwater vision than do harbor seals (Phoca vitulina) or California sea lions (Zalophus californianus), as determined by ability to discriminate between subsurface disks of slightly different diameter (Gentry and Peterson, 1967). Sea otters are approximately emmetropic in both air and water with an accommodative range of 59 D (Murphy et al., 1990), a range three times greater than that reported for any other mammal (Sivak, 1980). The large accommodation is accomplished through a highly developed iris musculature, meridional ciliary muscle, and corneoscleral venous plexus (Murphy et al., 1990). Using this tremendous accommodation to compensate for the refractive loss of their corneas upon immersion in water, sea otters can clearly focus on objects both above and below water. The cornea of a sea otter eye has a thickness centrally of 0.3 mm, and the anterior epithelium of the cornea is greatly developed, comprising about one third of the corneal thickness.
Social Organization Free-living sea otters segregate into male and female areas (Kenyon, 1969). Females may form matrilineal groups with resident females, their adult daughters, and young offspring sharing feeding and resting areas within home ranges for several years (Lyons, 1991). Food preferences and foraging strategy appear to be shared vertically within these family groups. While few females are found in male areas, male otters will enter female areas and defend territories against incursions by other males (Kenyon, 1969; Riedman and Estes, 1990). Territorial males then mate with multiple females. Large groups of nonbreeding males may congregate in bachelor groups bordering on female areas. Adult mortality rates are higher in males than in females (Siniff and Ralls, 1991; Monson and DeGange, 1995).
Reproduction Male sea otters reach sexual maturity by age 5 to 6 years (Schneider, 1978), but males under 6 years are not successful breeders (Garshelis, 1984). A 19-year-old male successfully fathered a pup in captivity (Reidman and Estes, 1990). Spermatogenesis occurs throughout the year (Lensink, 1962). Male breeding behavior also can occur throughout the year (Brosseau, et al., 1975). Sea otters have delayed implantation of blastocysts. Unimplanted blastocysts can be found in preserved reproductive tracts, but their detection requires serial sectioning of the entire uterine horn. The copora lutea of implanted animals measure 9 to 17 mm in diameter (Sinha et al., 1966). The chorioallantoic placenta is zonary and endotheliochorial. There can be considerable temporal plasticity in reproductive events in female sea otters based on individual and geographic variations. Most female sea otters have their first estrus at 4 to 5 years of age, but a few individuals may have their first estrus as early as 2 to 3 years old (Bodkin et al., 1993; Kenyon, 1969; Garshelis, 1984). A captive-raised female successfully delivered a live pup when just under 3 years of age (Crossen, pers. comm.). In California, female sea otters give birth after a gestation period of 4 to 6 months (Loughlin et al., 1981; Jameson and Johnson, 1993). In the latter study, the 6-month gestation period was considered to consist of an unimplanted phase of 2 to 3 months, and an implanted phase of 4 months. In Alaskan sea otters, the unimplanted phase may last from 3.5 to 4.5 months (Schneider, 1972; 1973; 1978) to 7 to 8 months (Kenyon, 1969). Twinning occurs
0839_frame_C44 Page 966 Tuesday, May 22, 2001 2:09 PM
966
CRC Handbook of Marine Mammal Medicine
infrequently (<2%) in sea otters (Schneider, 1973), and one of the twin pups is usually lost at birth or shortly thereafter (Williams et al., 1980; Jameson and Bodkin, 1986). Field observations indicate that the female comes into estrus again soon after weaning or the loss of a pup, and typically mates in less than 1 day to several weeks after weaning her pup. The average period of female receptivity is 3 to 5 days (Brosseau et al., 1975; Garshelis 1984). Old female otters may have difficulty in conceiving or maintaining pregnancy, resulting in the repeated recurrence of estrus (Riedman and Estes, 1990). Hormone assays in captive females indicate that they are seasonally polyestrous with estrus occuring in late winter or spring and again in late summer or fall (Larson, unpubl. data). Females with dependent pups rarely copulate (Lensink, 1962; Kenyon, 1969; Schneider, 1972; Calkins and Lent, 1975; Garshelis, 1984). Mating is frequently prolonged and violent. Copulation always occurs in the water, usually after a short period of courtship during which an estrus female becomes increasingly receptive to the male (Kenyon, 1969). In some cases, very aggressive males may forcibly mate with females or separate females from their pups in breeding attempts. During copulation attempts, the male holds the female’s nose and face in his teeth and may forcibly hold the female’s head under water (Riedman and Estes, 1990). Lacerations to the nose and face are common, resulting in characteristic scarring in most mature females (Foott, 1971). These mating injuries may be fatal as a result of the extent of the facial wounds, or of drowning (Staedler and Riedman, 1993). The length of pup dependency is very variable, from 5 to 13 months (Wendell et al., 1984; Rotterman and Simon-Jackson, 1988; Jameson and Johnson, 1993). Annual reproductive rates increase from 22% at 2 years of age to 78% at 5, thence remaining stable at least through 15 years (Bodkin et al., 1993). Only 30% of pups survive their first year (Jameson and Johnson, 1993) and immature females are apparently less capable than experienced adults in successfully raising their young (Kenyon, 1969; Wendell et al., 1984). Newborn pups weigh about 1.4 to 2.3 kg and are covered by a wooly natal pelage that is lost by 10 to 13 weeks of age (Kenyon, 1969; Wendell et al., 1984). Pups are totally dependent on maternal care for food and grooming for the first 2 to 3 months, and are carried almost constantly on the mother’s upturned chest and abdomen. A pup may be left floating on the water or placed on a nearby haul-out area for brief periods while the mother forages or grooms, where it vocalizes loudly until the mother returns (Sandegren et al., 1973). California sea otters spend 20 to 30% of their time grooming and nursing their young, 41% resting with the pup, 16% feeding, and only 10% grooming themselves (Sandegren et al., 1973). Newborns rely solely on their mother’s milk for the first 3 to 4 weeks and then begin to feed on small pieces of prey items offered by their mothers. Diving ability develops between 6 and 10 weeks of age and pups may begin successfully foraging by 14 weeks, but continue to have difficulty opening hard-shelled items until closer to 5 or 6 months of age. Although otters have been observed apparently nursing very large pups (estimated to be 7 to 8 months of age) and grooming even older pups, mean dependency of one study group of 12 knownage pups was 6 months (Payne and Jameson, 1984). Apparent “adoption” (nursing and grooming) and tolerance of approach of orphaned pups has been reported (Kenyon, 1969; Staedler and Riedman, 1989; Williams, 1990). Pupping intervals of 1 to 2 years have been reported (Loughlin et al., 1981; Wendell et al., 1984). Birth may occur on land or in the water (Kenyon, 1969; Antrim and Cornell, 1980; Jameson, 1983). Seasonal peaks of pupping occur and vary with region. In California, pupping peaks between January and March (Fisher, 1940; Sandegren et al., 1973), while it peaks in May in Alaska (Schneider, 1978; Rotterman and Simon-Jackson, 1988), and in May/June in Russia. After birth, the corpus luteum of the ovary rapidly degenerates, and a distinguishable corpus albicans persists for at least 2 years (Sinha and Conaway, 1968).
0839_frame_C44 Page 967 Tuesday, May 22, 2001 2:09 PM
Sea Otters
967
Causes of Mortality in Free-Living Otters Mortality in wild sea otters has been investigated in California by Ames et al. (1983), and documented in other areas by a variety of authors. Barabash-Nikiforov et al. (1947) and Estes et al. (1998) report killer whale predation as important in the North Pacific, although this was considered uncommon by Kenyon (1969). White sharks frequently injure and sometimes kill sea otters in California (Ames and Morejohn, 1980; Ames et al., 1996). Eagles (Sherrod et al., 1975), coyotes, and brown bears (Reidman and Estes, 1990) have also been reported to take sea otters. Starvation can occur rapidly if an otter is unable to forage normally, and may occur secondary to infections, damaged pelage, parasitism, or toxicity (Morejohn et al., 1975). Hemorrhagic gastroenteritis is frequently seen associated with stress, starvation, or parasitic infestations (Barabash-Nikiforov et al., 1947; Stullken and Kirkpatrick, 1955; Lipscomb et al., 1993). Bacterial infection can occur anywhere in the body due to wounds, damaged teeth, or parasite migration (Thomas and Cole, 1996) and may result in valvular endocarditis (Joseph et al., 1990). Adverse weather conditions may separate mothers from dependent young or stress otherwise compromised otters. Mating injuries, male–male aggression, and complications of pregnancy may account for small numbers of mortalities (Ames et al., 1983). Human interactions cause sea otter mortality through net and fishing gear entanglement, shooting, capture operations, oil and other toxic spills, and collisions with boats and props (Ames et al., 1983; Reidman and Estes, 1990). In 1972, the Marine Mammal Protection Act made hunting of sea otters legal for Alaska natives for subsistence, and creation and sale of authentic native articles of handicrafts and clothing, provided that the taking is not wasteful.
Feeding and Metabolism Sea otters occupy an inshore littoral habitat. Although they might be found offshore during emigration movements, their preferred habitat lies within waters shallow enough for them to forage for benthic invertebrates. Generally, otters make foraging dives at depths of 28 to 30 m (Wild and Ames, 1974; Estes, 1980). However, a sea otter drowned in a crab pot set at about 100 m, indicating that deeper dives are possible (Newby, 1975). Although a very wide variety of prey items have been described for sea otters (Table 1), most individual otters tend to focus on one to several food items. Food preferences, likely taught to pups by their mothers, persist for up to 5 years. The presence of a pup adversely influences the selection of prey by female otters, probably by limiting dive depths and durations, as well as limiting the locations for foraging to those suitable for the pups (Garshelis et al., 1986; Van Blaricom, 1988a). Sea otters sometimes use rocks or heavy shells as tools against which to pound prey items, especially bivalves (Hall and Schaller, 1964; Hall, 1965), and have been observed collecting invertebrate prey from discarded bottles and cans. Sea otters ingest about 20% of their body weight in food per day. Daily food energy equivalents have been calculated to be to 189 (Kenyon, 1969), 234 (Costa, 1982), and 307 (Fausett, 1976) kcal/kg/day. Captive sea otters ingested 2.4 times their standard metabolic rate (Morrison et al., 1974) and eight times the standard metabolic rate of similar-sized terrestrial mammals (Costa, 1982). This high metabolic rate can be explained in part by the increased heat loss caused by the high thermal conductivity of water on a relatively small animal that has a high ratio of surface area to volume. In addition, sea otters have the lowest assimilation efficiency (82%) reported for carnivores, possibly due to their rapid (3-hour) gut transit time (Costa, 1982). Sea otters must forage as often as every 5 hours (Loughlin, 1977), but a rapid gut passage
0839_frame_C44 Page 968 Tuesday, May 22, 2001 2:09 PM
968
CRC Handbook of Marine Mammal Medicine
TABLE 1 Prey Items Consumed by Sea Otters throughout Their Range
Classification Phylum Phylum Phylum Class Phylum Subphylum Class Order Class Order Order Order Phylum Class Class Class Class Phylum Class Class Class Class Phylum Class Class Class Family Family Family Family Family
Echiura Nemertea Annelida Polychaeta Arthropoda Crustacea Cirripedia Thoracica Malacostraca Isopoda Amphipoda Decapoda Mollusca Gastropoda Bivalvia Polyplacophora Cephalopoda Echinodermata Echinoidea Asteroidea Ophiuroidea Holothurioidea Urochordata Ascidiacea Osteichthyes Aves Anatidae Gavidae Laridae Phalacrocoracidae Podicipedidae
Common Name
Number of Identified and Unidentified Genera and Species
Spoonworms Ribbonworms
2 1
Polychaete worms
5
Barnacles 3 Pillbugs Beach fleas Shrimps, crabs
2 2 30
Snails Bivalves Chitons Octopus
24 41 7 3
Sea urchins Starfish Brittlestars Sea cucumbers Chordates Sea squirts, tunicates Bony fish Birds Ducks Loons Gulls Comorants Grebes
5 9 2 4 2 17 2 1 1 1 1
Source: Derived from Riedman and Estes (1990).
time means that they lose part of the food energy they ingest. Costa (1982) calculated the net metabolizable energy to be 168 kcal/kg/day or 72% of ingested food (see Chapter 36, Nutrition). The osmolality of sea otter urine (mean ± SE: 1627 ± 104 mOsm/kg; maximum: 2130 mOsm/ kg) is no greater than that of fish-eating pinnipeds (Costa, 1982). However, the lobulated kidney of sea otters is large (2% of body mass) compared with kidneys of bottlenose dolphins (1.1%, Tursiops truncatus), harbor seals (Phoca vitulina) (0.25 to 0.4%), and northern fur seals (Callorhinus ursinus) (0.33 to 0.71%). The greater kidney mass may permit an increased glomerular filtration rate, allowing production of larger urine volumes. Seawater taken in with food, water contained in food, and metabolic water are the principal sources of fluid. The invertebrates that form much of the prey of sea otters have higher electrolyte concentrations than do fish. This may require wild sea otters to drink seawater (estimated 0 to 124 ml/kg/day) to maintain water balance (Costa, 1982). Given a high-protein diet, requiring elimination of large amounts of urea, seawater drinking may assist in increasing urine volume and reducing urine concentration. Captive sea otters will drink fresh water and chew and eat ice, but this does not appear to be a major source of fluid intake in wild otters.
0839_frame_C44 Page 969 Tuesday, May 22, 2001 2:09 PM
Sea Otters
969
Husbandry The basic tenets of captive sea otter care were enumerated by Barabash-Nikiforov et al. (1947), based on observations of a long-term project conducted between 1932 and 1940 in Russia. He stated that sanitation, temperature susceptibility, access to clean water, and frequent feeding of good-quality fish and whole invertebrates were critical to sea otter survival and concluded that “sea otters are quickly and fully tamed.” Transport and long-term holding techniques were explored in the early 1950s at field study sites in Alaska (Stullken and Kirkpatrick, 1955). Based on these experiences, Kenyon (1969) stipulated that sea otters must have an abundant supply of clean (preferably salt) water; adequate clean, dry haul-out space; good air circulation; and food equal to 20 to 25% of their body weight per day. He also reported that, especially during transport, care must be taken to avoid soiling the fur or otters will lose their waterproofing and become susceptible to hypothermia and death. Housing requirements for captive sea otters are governed by the U.S. Department of Agriculture (USDA, 1995) (see Chapter 33, Legislation). Formulas are provided for calculating haulout or dry resting area (DRA) and minimum horizontal tank dimension (MHD) for sea otters. Water must have a depth of not less than 0.91 m (3 ft). Inclined ramps leading to the haul-out areas are useful. Emergency transport and holding facilities used for rehabilitation are exempt from USDA regulations, but do require permitting by the U.S. Fish and Wildlife Service, which has oversight authority for all captive sea otter activities. In a temporary facility, pools holding adult otters should still provide at least 0.91 m (3 ft) of depth as above. Fresh water can be used when temporarily housing sea otters; however, the water must be free of chlorine. If a sea otter is kept in chlorinated water more than 10 days, the ends of the hairs will split and become matted. Salt water must be used in a permanent facility, as the long-term effects of fresh water on the sea otter are not known. Salt water may be artificially created or supplied from natural bodies of water after adequate filtration. Water temperature should be maintained at 7 to 15.5°C (45 to 60°F) and should never exceed 21°C (70°F) (Williams, 1990; Tuomi et al., 1995). Good skimming of the surface water is essential. This is accomplished by jets placed at the waterline or by placing skimming boxes at the surface to collect water and keep it free of scum or debris. Water turnover time should be 0.5 to 1.5 hours. Fecal coliform counts can rise dramatically when sea otters are housed in pools, due to their high food consumption, poor food assimilation, and consequent high-volume fecal output (Van Blaricom, 1988b). Recirculated water should be filtered through sand filtration and disinfected with ultraviolet sterilization or ozone treatment (Nightingale, 1981; Tuomi et al., 1995). Shell and discarded food fragments can rapidly accumulate in the pool bottom, and provision must be made for their frequent removal. Soiling of the fur with food or fecal debris can lead to loss of waterproofing with potentially disastrous results. Salt water may be warmed with a swimming pool “spa” heater (maximum temperature 68°C, or 154°F) and temporarily provided for otters having difficulty thermoregulating because of medical problems. This allows animals to spend more time grooming and feeding in water without becoming hypothermic (Tuttle, pers. comm.). Water temperature is gradually reduced as metabolic rate returns to normal levels. The fur may have decreased loft at water temperatures above 15.5°C (60°F) but regains waterproofing as the otter grooms in increasingly cold water (McBain, pers. comm.). Sea otters are agile climbers. Pools and pens must be constructed to include overhangs, a secure roof or at least 4 ft of smooth vertical wall. Netting or chain link (1-in. mesh) may be used in enclosures and will allow good air circulation and ventilation. Sea otters are notorious for their ability to cause damage by manipulating objects within their enclosures. They will
0839_frame_C44 Page 970 Tuesday, May 22, 2001 2:09 PM
970
CRC Handbook of Marine Mammal Medicine
disassemble exposed fasteners, scratch painted or acrylic surfaces with rocks and shell fragments, and damage their teeth chewing on corners or dividing bars. Consideration should be given in design of enclosures to eliminate risk of loose materials and to round corners and space openings small enough to prevent chewing. Materials for construction of haul-out areas must be easily cleaned and not damaged by chewing. Haul-out surfaces must have no sharp edges and should allow for drainage of water so the fur can dry. Stretched, thick, stranded rope netting, smooth hard plastic, or sealed concrete can be easily cleaned and helps prevent injuries to feet and legs from cuts or pressure sores. Captive otters benefit from access to objects that they can chew and manipulate. Whole food items, blocks of ice, kelp strands, feeder toys, large unbreakable balls, floats, and large dog chew toys provide diversion and encourage exercise. Artificial objects should be large and tough enough to avoid accidental ingestion. Sea otters should be held in social groups of two or more animals of the same sex, or in breeding groups with one male and one or more females. Large groups of bachelor males occur in the wild and can be held for short periods in captive enclosures as long as females are not present (Tuomi, 1990). Calle et al. (1997) reported on the use of depot leuprolide and cyproterone to control aggression in a colony of four male California sea otters. Females with newborn pups may need to be held in an area separate from other otters, especially if the mother is inexperienced. Injury or death of pups has been observed on several occasions when a dominant female or aggressive male attempted to take the pup from its mother (Reidman and Estes, 1990). Transportation of sea otters over short distances may be accomplished using a standard plastic canine airline kennel equipped with a raised slotted floor to prevent soiling of the fur by allowing water, urine, and feces to fall away from the animal. Blocks of ice or wet towels rolled and frozen into “logs” may be placed in the kennel with the otter to prevent overheating while the animal is out of the water. For longer shipment or for prolonged holding out of the water for medical purposes, otters should be placed in open mesh net boxes or carts with slotted floors (Figure 1). The additional open space allows more adequate ventilation, and the animals can be easily rinsed with a hose or garden spray bottle to maintain temperature and cleanliness. Transport vehicles and holding areas should be cooled to no more than 15.5°C (60°F). Proposed USDA regulations would require an experienced handler to travel with a sea otter to ensure adequate conditions and care.
FIGURE 1 Open mesh net box with slotted floor for extended sea otter holding or shipping. Note otter’s normal pelage.
0839_frame_C44 Page 971 Tuesday, May 22, 2001 2:09 PM
Sea Otters
971
Captive Nutrition The typical captive sea otter diet is very high in water content (70 to 78%) with more than 70% of the calories provided by protein. By comparison, cat food is typically 25 to 35% protein and dog food is 18 to 23%. Only 10 to 20% of the calories in a sea otter’s diet comes from fat and almost none from carbohydrate (Tuomi et al., 1990). Clams, squid, mussels, crab, abalone, octopus, scallops, and shrimp may be fed whole or with the hard parts of shell or carapace removed to reduce littering of the enclosure. The ink sac may also be removed from squid to avoid dark discoloration of the feces, which may resemble melena (Williams, 1990). A variety of fish (usually fillets) including pollock, herring, smelt, salmon, flounder, cod, lumpsuckers, and rockfish have also been used. Barabash-Nikiforov et al. (1947) stated that sea urchins and crab or whole young cod (undressed) must be included in the diet to prevent diarrhea. Roughage, in the form of some shell, carapace, or bone does appear to be necessary for a normal, formed stool and may contribute to the mineral nutrition of otters (Tuomi et al., 1995). Sea otters do not digest fish bones well and fatal bony impactions have been found in northern sea otters that apparently had fed on a large volume of whole fish carcasses (Lowenstine, pers. comm.; Tuomi, unpubl. obs.). All food should be fresh-frozen to decrease parasite transmission (Sweeney, 1965) and thawed in a refrigerator or under cold running water to decrease opportunity for bacterial growth. Thawed food may be kept refrigerated at 4 to 6°C (40 to 43°F) or on ice for up to 24 hours (Crissey, 1998). Items may be refrozen into “ice treats” for enrichment. Because of their high metabolic rate and rapid gut transit time, sea otters should be fed at 3- to 4-hour intervals during the day with a total intake of about 20 to 25% of body weight per animal per day. Food may be thrown into pools for free-feeding or offered by hand or with metal tongs as part of operant conditioning or nursing care programs. Alternatively, unbreakable trays may be placed on haul-out surfaces or buckets lowered into the water to allow otters to forage for themselves. Uneaten portions should be collected and discarded before each subsequent feeding (Tuomi et al., 1995). Bacterial pathogens are common in raw seafood and may cause enteritis or death. Enhanced serum antibody titers to Salmonella newport, S. pullorum, S. typhimurium, Vibrio alginolyticus, and Pasturella multocida were detected in sea otters in the Exxon Valdez rehabilitation facilities (Wilson et al., 1990). Multivitamin mineral supplementation should be provided for all captive sea otters. Exact requirements have not been defined, but several commercially prepared marine mammal formulations are available and appear to be safe and efficacious. Tablets may be inserted into the mantle cavity of invertebrates or into partially thawed fish fillets; however, sea otters chew their food well and may refuse or discard food if the taste is disagreeable. Muffins containing krill and vitamins (Otten-Stanger, pers. comm.) and pureés of food items mixed with supplements or medications and then refrozen (Murray, pers. comm.) have been used successfully in some facilities. Newborn sea otter pups that are unable to nurse from their mothers must be fed an artificial formula by bottle or stomach tube for the first 2 to 3 months of age (see Chapter 37, HandRearing). Solid food may be introduced after the first month in the form of small, soft pieces of adult diet. The amount of formula fed is gradually decreased as solid food intake increases. By 20 to 24 weeks of age, most pups can procure their own food, and have begun to open hard-shelled items with rock or shell tools (Payne and Jameson, 1984).
Physical and Chemical Restraint Adult sea otters are large, carnivorous mustelids well equipped with crushing molars, long canine teeth, and retractable front toenails that can inflict serious injury. The loose skin of sea otters
0839_frame_C44 Page 972 Tuesday, May 22, 2001 2:09 PM
972
CRC Handbook of Marine Mammal Medicine
allows them to twist and turn rapidly, even when their body is firmly grasped. Since they can reach all parts of their body to groom, it follows that they can also bite a hand placed almost anywhere on their body. It is easy to be lulled by their typically benign demeanor and cuddly appearance, but an otter can quickly become a bundle of fury when it is touched or restrained. To avoid serious injury to the animal and to personnel, all attempts at handling should be well planned and executed with proper equipment. Only experienced personnel should attempt these procedures, and care must be taken to plan procedures to minimize handling time to prevent overheating and excessive stress. One person should be assigned to observe the otter continuously during restraint. Corrective or preventative measures must be readily available, such as cooling with water or ice, shading from sunshine, and adequate ventilation. Williams and Sawyer (1995) reported on successful techniques used on a large number of otters in the rehabilitation centers during the Exxon Valdez oil spill. Restraint equipment included heavy leather welder’s gloves with long sleeves, a large salmon dip net fitted with an elongated bag of soft 1-in. mesh net, a throw net, blankets, stuff bags, and some sort of squeeze box. Figure 2 illustrates a frequently used squeeze box design (Williams et al., 1990). An otter can be netted from the water or haul-out area and enclosed in the net by a drawstring or by rotating the hoop to close off the open bag. Otters may be carried short distances by suspending the net between two or more people; body weight may be obtained by hanging the net from an overhead scale hook. The animal can be further restrained using a heavy cloth, nylon, or burlap bag (3 ft long by 1.5 ft diameter) filled with foam rubber or other soft, compressible material. The bag is quickly placed over the head, shoulders, and thorax, and the otter is held by pressing down on the bag. This is best accomplished while the otter is in a net or squeeze box with the animal on its back and the hind legs held by a second handler (Williams et al., 1990; Williams and Sawyer, 1995). Debilitated or very lethargic animals may be held for treatments and for washing and drying by placing the forelegs through a figure-eight loop of thick rope or flexible canine “pull-toy” grasped from behind the head and neck.
FIGURE 2 Drawing of squeeze box used for sea otter restraint. (From Williams, T.M., and Davis, R.W. (Eds.), Emergency Care and Rehabilitation of Oiled Sea Otters, University of Alaska Press, Fairbanks, 1995, 40. With permission.)
0839_frame_C44 Page 973 Tuesday, May 22, 2001 2:09 PM
Sea Otters
973
Chemical agents and dosages reported as used in sea otters are listed in Chapter 31 (Pharmaceuticals). The most commonly used agents are reversible narcotic sedatives and neuroleptanalgesics, such as fentanyl or oxymorphone, combined with diazepam and azaperone or acepromazine (Williams and Kocher, 1978; Williams and Sawyer, 1995; Sawyer and Williams, 1996). Intramuscular (IM) injections of these drugs may be combined in one syringe and administered by blow dart, jab pole, or with squeeze restraint. Fentanyl, when combined with diazepam and administered IM to wild, healthy otters in field studies, required a higher dose (0.33 to 0.22 mg/kg) than that reported for captive or rehabilitation protocols. These higher doses were associated with profound decreases in oxygen saturation of the blood (Mazet, pers. comm.). Diazepam, even in very small dosages (0.07 to 0.11 mg/kg), was effective in controlling convulsions and tremor associated with the use of fentanyl in these otters. Reversal with naltrexone at a dose equaling 1.5 to 2 times the total fentanyl administered was preferred (Monson et al., 2000). Naloxone (1 mg/0.1 mg of fentanyl dose) has been used for narcotic reversal but has occasionally been associated with complications in the field and in rehabilitation efforts due to renarcotization (the reversal agent is metabolized before the narcotic is eliminated) and subsequent risk of drowning. The dose of naloxone is often divided giving half intravenously for immediate arousal and half IM to provide longer effect. The use of other agents has been reported in sea otters with variable results. Oxymorphone (0.3 to 0.55 mg/kg IM) may be given alone or in combination with diazepam (0.5 mg/kg IM). This provides good sedation after 15 to 20 min of quiet rest and may be followed with inhalation anesthesia. This sedation may be reversed with naloxone (0.03 mg/kg or 0.4 mg/45 to 60 kg animal) (Brown and Huff, pers. comm.). Medetomidine has been reported safe and effective in river otters in combination with various other agents (Spelman, 1997) and has been used in combination with butorphanol in captive sea otters with atipamizole reversal (Murray, pers. comm.). Dissociative anesthetics have been associated with fatal complications (Williams and Kocher, 1978) and are not recommended. Isoflurane gas anesthesia is effective and generally safe in otters of all ages, but should not be used on heavily oiled animals because of apparent aggravation of lung damage from volatile hydrocarbon vapors (Williams and Sawyer, 1995). Mask induction and endotracheal intubation follow standard mammalian techniques. Premedication with atropine has not been reported in sea otters. Peripheral vasodilatation associated with the use of isoflurane may potentiate hypotensive effects of shock and hypothermia (Tuomi and Williams, 1995). Temperature monitoring is essential during both physical and chemical restraint of sea otters (see below) and adjustments should made before critical limits occur.
Clinical Examination Physical examination of wild or untrained sea otters can be challenging. Initial evaluation should be conducted from a distance, preferably while the animal is at rest. Standard considerations include an estimate of age based on body size and coat color and character. Note should be made of respiratory rate and character, attitude, mobility and posture, and presence of wounds, discharges, or obvious injuries. Heart rate may be determined by observing the apex beat moving under the ribs on the ventral chest. Normal respiration at rest is 17 to 20 breaths/ min and heart rate is 144 to 159 beats/min (Williams, 1990). The fur should be evaluated for loss of waterproofing (see Loss of Coat Condition, below) and, if present, the location and size of the abnormal pelage recorded. Weight should be recorded during any handling or examination and record should be made of whether the coat is wet or dry. The guard hair may hold significant amounts of water when an otter is first removed from a pool, and considerable error is introduced if weights are not taken consistently.
0839_frame_C44 Page 974 Tuesday, May 22, 2001 2:09 PM
974
CRC Handbook of Marine Mammal Medicine
With the otter physically restrained, body temperature may be roughly estimated by feeling the rear feet. Body warmth can normally be felt through the shorter fur on the toes and interdigital web. Cold or unusually warm feet signal hypothermia or hyperthermia, respectively. If possible, body temperatures should be taken using a flexible thermometer inserted several inches into the rectum (Electrotherm Digital Thermometer®, Veterinary Specialty Products, Boca Raton, FL). Normal body temperature for adult sea otters is 37.5 to 38.1°C (99.5 to 100.6°F) (Williams, 1990). Short-term restraint with a stuff bag and net or squeeze box is usually adequate for administration of IM or subcutaneous injections; flipper or microchip tagging; palpation of the abdomen and pelvic limbs; sampling of hair, feces, milk, or urine and for routine blood sampling from the femoral or popliteal veins (see Chapter 19, Clinical Pathology, for venipuncture sites). Urine can easily be collected from a full bladder by cystocentesis using a 1.5- or 2-in., 21- or 22-gauge needle placed about one third of the distance anteriorly from the brim of the pubis toward the umbilicus. Care should be taken with adult female otters to ensure that the bladder is palpated and differentiated from an enlarged uterus. Radiographs and ultrasound, prolonged treatments, painful procedures, and thorough examination of the chest, forelegs, and head must be performed under sedation (above). Ocular injuries are frequently seen as corneal ulcers or lacerations progressing to scars or phthisis bulbi. Dental wear, caries, and fractured teeth are common in middle-aged and older animals and may lead to malnutrition and fatal infections. Auscultation of the chest may reveal crepitus associated with pulmonary interstitial emphysema, or a heart murmur secondary to septic valvular lesions.
Medical Abnormalities Hypoglycemia Hypoglycemia, hypothermia, and dehydration are common findings in debilitated sea otters. Presumptive therapy can be initiated by the administration of warmed (37 to 39°C, 98.6 to 102°F) lactate-free, balanced electrolyte and glucose solution (2.5% dextrose in normal saline) given subcutaneously, intraperitoneally, or via the jugular vein at a dose of 20 ml/kg (Williams et al., 1995a). Prolonged intravenous access may be obtained by placement of an intraosseous catheter in the proximal femur (Black and Williams, 1993). Hypoglycemia is rapidly confirmed by use of over-the-counter blood glucose monitoring kits sold for home use by people with diabetes. Blood sugar of less than 60 mg/dl may cause profound depression and seizures, and should be immediately treated by intravenous administration of 10 to 20% dextrose (10 to 20 ml/ kg to effect). Oral dosing of 50% dextrose (1 mg/kg) can be given via stomach tube in debilitated but conscious otters, or dextrose may be added to chipped ice balls and offered for the otter to chew (“sno-cones”). Food should be provided as soon as possible if the otter will eat or a high calorie slurry of pureed seafood and human enteral supplements may be given by stomach tube (Williams et al., 1995a).
Hyperthermia The high metabolic rate of sea otters predisposes them to rapidly developing hyperthermia (body temperature >38°C or 101°F) whenever a healthy animal is held out of water. This effect may be worsened by struggling, drying of the hair coat, or warm ambient temperatures. The risk is also exacerbated during anesthesia and anesthetic recovery. Provision must be made to maintain or adjust body temperature. Cooling may be achieved by wetting the hair coat, using ice on the flippers, trunk, and neck, and by the use of fans and air conditioners.
0839_frame_C44 Page 975 Tuesday, May 22, 2001 2:09 PM
Sea Otters
975
Additional fluid intake may be accomplished orally by offering otters ice chunks or chipped ice sno-cones to eat.
Hypothermia Debilitated animals, especially those with inadequate grooming or nutrition, or sedated animals may become hypothermic. Hypothermia is best prevented in debilitated sea otters by ensuring adequate caloric intake and by drying of the fur. Animals with body temperature less than 35°C (95°F) are classified as hypothermic and require immediate attention. Blood sugar should be assayed to determine if hypoglycemia is a contributing factor and supplemental calories provided, if necessary, by administration of warmed (37 to 39°C, or 98.6 to 102°F) subcutaneous, intravenous, intraosseous, or intraperitoneal fluids, or by oral intubation (if the animal is sufficiently conscious). Passive rewarming at a rate of 0.5°C (1°F) per hour is achieved by placing the animal in a warm room (20°C, or 68°F) in a dry cage and stimulating grooming with towels or slightly warm air from hair dryers or fans set on “cool” or “low.” Active or rapid rewarming of severely hypothermic animals (temperature less than 35°C, or 95°F) with warm immersion water baths can be dangerous and may lead to lactic acidosis, paradoxical temperature after-drop, or hypokalemic cardiac arrythmias (Williams et al., 1995a). Complications of severe hypothermia may occur in humans following rewarming and include pneumonia, gastric erosions, intravascular erosions, and acute tubular necrosis (Bowen and Bellamy, 1988). Acute and chronic renal lipidosis, and hepatic necrosis and gastric erosions and hemorrhage were observed at post-mortem examination of otters dying after treatment for severe oiling and hypothermia during the rehabilitation effort following the Exxon Valdez oil spill (Lipscomb et al., 1993).
Loss of Coat Condition Loughlin (1977) described normal grooming behaviors in wild sea otters and divided them into four stages, each apparently designed to clean, align, and trap air into the pelage and distribute sebaceous secretions over the skin and hair. Grooming activities account for 5 to 16% of the daily activity of a wild otter (Estes et al., 1986), and captive otters may spend 25% of each day in grooming (Antonelis et al., 1981). Familiarity with normal grooming behaviors and the appearance of normal (Figure 3a) and abnormal (Figure 3b) pelage is essential to good sea otter husbandry. Grooming typically takes place in the water, especially after eating. Otters vigorously rub their fur with forepaws and nose, roll repeatedly forward or laterally (“log rolling”), and appear actually to blow air into the coat. Hauling out to dry and sleep appears to be more common in northern latitudes where ambient water temperatures tend to be much lower, especially in winter. If a sea otter’s fur becomes fouled by food, fecal debris, oil, or any other hydrophilic material, waterproofing is lost. This may occur if the otter is too debilitated by other conditions to groom, if the otter does not have access to adequate amounts of clean water, or if the fur is contaminated by oil or other agents in the water or on the haul-out area. If the otter is unable to groom soiled areas successfully, the animal becomes wet to the skin in the fouled area. The loss of air insulation causes body heat to be lost at an increased rate, especially when the otter is in water feeding or grooming. Costa and Kooyman (1982) and Davis et al. (1988) demonstrated that fouling of only 18 to 20% of the coat of a sea otter with crude oil resulted in a 40 to 200% increase in metabolic rate to compensate for the loss of insulation, and the increased activity expended in attempts to clean the area. Otters that have lost coat condition may be unable to remain in water or to forage; they rapidly deplete body stores and become hypoglycemic and hypothermic, and this can lead
0839_frame_C44 Page 976 Tuesday, May 22, 2001 2:09 PM
976
CRC Handbook of Marine Mammal Medicine
A
B
FIGURE 3 (A) Normal appearance of (wet) pelage in a sea otter. Note the striated appearance of normal, waterrepellent fur (From Williams, T.M., and Davis, R.W. (Eds.), Emergency Care and Rehabilitation of Oiled Sea Otters, University of Alaska Press, Fairbanks, 1995, 97. With permission.) (B) Abnormal, oiled pelage (rear-end of the body) in a sea otter. Note the matted or slick appearance of the fur that is saturated with oil.
to circulatory collapse, shock, and death unless supportive care is instituted (Williams et al., 1995a).
Oil Exposure Oil spills represent one of the greatest challenges for continued survival of the sea otter, because of their inshore distribution. Sea otters in California are especially at risk, because of their limited habitat. Efforts have been made to translocate sea otters into new habitat in California, but these efforts have been hampered by the strong tendency for translocated animals to return to their established home ranges. The grounding of the tanker vessel Exxon Valdez in Prince William Sound, Alaska, is estimated to have killed up to 5000 sea otters (Garrott et al., 1993; DeGange et al., 1994; Garshelis, 1997), with the documented recovery of nearly 1000 carcasses (Ballachey et al., 1994). Oil-contaminated sea otters were treated in rehabilitation facilities between April and August 1989 (Williams and Davis, 1990). Dawn® (Procter & Gamble, Cincinnati) dishwashing detergent diluted 1:16 in water (temperature 28 to 32°C, or 82 to 90°F) was used to wash oil from the fur while otters were given intravenous, subcutaneous, or intraperitoneal fluids,
0839_frame_C44 Page 977 Tuesday, May 22, 2001 2:09 PM
Sea Otters
977
antibiotics, corticosteroids, vitamins, and oral activated charcoal. Otters were thoroughly rinsed using temperature-controlled fresh water and hand-held high-pressure showerheads and then dried with towels and pet hair dryers set at room temperature. Most otters were anesthetized or sedated for the wash-and-dry procedures although moribund or severely lethargic animals could be restrained manually. Body temperature was monitored throughout the procedure, and animals were cooled or warmed as needed by adjusting wash and rinse water temperature. Histopathological lesions frequently observed in sea otters exposed to Prudhoe Bay crude oil in 1989 included gastric erosion and hemorrhage, centrilobular hepatic necrosis, periportal and diffuse hepatic lipidosis, and renal tubular lipidosis (Lipscomb et al., 1993). The authors theorized that hepatic and renal lipidosis may have been caused by an oil-associated increase in energy demand occurring with decreased food intake, resulting in mobilization of stored fat. Centrilobular hepatic necrosis could have been a result of hepatic ischemia such as occurs with anemia and shock. Because gastric erosion and hemorrhage are common findings in captive sea otters, and one of six uncontaminated sea otters that died at the rehabilitation centers also showed these lesions, it was suggested that these were a result of stress. A significant number of otters exposed to the volatile fractions of the crude oil during the early days of the spill developed pulmonary interstitial emphysema and exhibited varying degrees of dyspnea and increased mortality. Although the exact mechanism remains unknown, Lipscomb et al. (1993) suggested that sea otters may be anatomically predisposed to interstitial emphysema by the presence of welldeveloped interlobular septa and that exposure to fresh crude oil resulted in remarkably high incidence of the lesion. Recommendations for treatment of oil-exposed sea otters have been published by Geraci and Williams (1990) and Williams and Davis (1995) and include early triage, supportive care for correction of shock, nutritional deficits, and hypothermia, and eventual washing with Dawn detergent. Long-term holding is necessary to allow otters to be supported medically and fed while they groom in copious amounts of clean salt water to restore the water-repellent air layer in their fur. Preemptive capture techniques to move otters from potentially contaminated habitat to safe captive care facilities is theoretically preferable, but logistically difficult. Although sea otter populations in many areas in Prince William Sound have increased, other sites have exhibited decreased survival rates in the years following the spill. Evidence suggests that the effects continued until at least 1998 (Monson et al., 2000). Most recently, continued exposure to petroleum hydrocarbons has been indicated by elevation of cytochrome P-4501A levels in sea otters living in the oiled areas of Prince William Sound compared with levels in sea otters in the un-oiled areas (Ballachey et al., 2000). The long-term significance of oil exposure is still under study (see Chapter 22, Toxicology), but Mazet et al. (1995) reported significantly reduced reproductive success in mink that had ingested crude oil. Vertical transmission of hydrocarbons may occur through the milk of contaminated females to their offspring (Tuomi and Williams, 1995), resulting in decreased survival of the young.
Abnormalities of Clinical Chemistry Hematological and serum chemistry values are similar to those found in other marine mammal species (see Chapter 19, Clinical Pathology) including relatively high hematocrit (HCT), hemoglobin, and blood urea nitrogen compared with domestic dogs and cats. These variations appear related to adaptations to diving and to a high-protein diet. Williams and Pulley (1983) and Williams et al. (1992) have published results of blood values from wild-caught sea otters. They reported that pups had significantly lower hematocrits, red blood cell counts, and hemoglobin levels when compared with adults, and that adult females had higher hemoglobin, white blood cell counts, and neutrophil counts compared with adult males.
0839_frame_C44 Page 978 Tuesday, May 22, 2001 2:09 PM
978
CRC Handbook of Marine Mammal Medicine
Analysis of clinical laboratory findings in oiled sea otters treated in rehabilitation centers following the Exxon Valdez oil spill revealed that lymphopenia and leukopenia were common, probably from stress and the severe inflammation associated with diarrhea, gastrointestinal hemorrhage, and secondary gastroenteritis. Hyperkalemia and hypoalbuminemia were also noted and may have been related to the diarrhea and gastrointestinal hemorrhage. Azotemia was common and may have been related to decreased renal perfusion from shock and dehydration. Hypoglycemia occurred apparently as a result of depletion of hepatic energy stores, and was probably the cause of many terminal seizures. Elevated hepatocellular transaminases may have occurred from leakage due to primary hepatotoxicity, or to hepatic lipidosis causing increased cell permeability and leakage into the bloodstream. Anemia was common, possibly related to gastrointestinal loss associated with erosion and hemorrhage or to direct red blood cell damage by hydrocarbon toxicity (Rebar et al., 1995).
Gastroenteritis Hemorrhagic gastroenteritis is common in sea otters and occurs rapidly after physical or physiological stress (Stullken and Kirkpatrick, 1955; Kenyon, 1969; Lipscomb et al., 1993). Affected animals will lose appetite, develop profuse foul diarrhea rapidly progressing to melena, and may die within 24 hours. Treatment may include intravenous or subcutaneous fluids, broad-spectrum antibiotics, motility modifiers, and corticosteriods. Prophylactic and therapeutic use of antacids, histamine (H2) blockers (cimetadine or ranitadine), protectants (sucralfate), and mood-modifying drugs (diazepam) may decrease morbidity and mortality but reduction of stress and maintainence of normal food intake are vital (Williams et al., 1995a) (see Chapter 31, Pharmaceuticals).
Parasites Infestation with nasal mites (Halarachne miroungae) has been documented (Kenyon et al., 1965), and may predispose to sinus or turbinate infections. Gastric ulceration and gastric and small bowel perforation and peritonitis can result from migration of larval forms of anisakid nematodes acquired when sea otters feed on infected fish (Rausch, 1953; Tuomi and Burek, 1999) (see Chapter 18, Parasitic Diseases). The cestode Diplogonoporus tetrapterus is associated with eating fish and occurs with moderate prevalence (12%) in sea otters in Prince William Sound, Alaska, but is absent from sea otters in California. Varying degrees of enteritis were reported associated with the presence of intestinal trematodes (Microphallus pirum) (Rausch, 1953). Parasite-related peritonitis occurs in southern sea otters infested with immature acanthocephalids of Profilicollis altmani (previously known as Polymorphus). Marine birds are the definitive host for this parasite and sea otters appear to become infested by ingesting intermediate stages found in sand-dwelling crabs. Mortality due Profilicollis infestation may be an important factor in the slow recovery of the southern sea otter population (Dailey and Mayer, 1999). By contrast, the sea otter is considered the definitive host for Corynosoma spp., which cause little morbidity unless present in very large numbers. The gall bladder fluke Orthosplanchnus fraterculus is common is some geographic areas, but heavy infestations do not appear to cause obstruction or debility despite fibrosis and scarring in the gall bladder (Rausch, 1953). Protozoal encephalitides associated with Toxoplasma gondii (Thomas and Cole, 1996) or with Sarcocystis neurona–like organisms (Chechowitz et al., 1999; Rosonke et al., 1999) have recently been reported in sea otters in California and Oregon (see Chapter 18, Parasitic Diseases). Serological testing may be helpful in diagnosing these conditions ante-mortem.
0839_frame_C44 Page 979 Tuesday, May 22, 2001 2:09 PM
Sea Otters
979
Miscellaneous Conditions Capture myopathy is a potentially fatal complication of lactic acid accumulation in muscle, frequently reported in other wildlife as a result of rigorous physical activity that may occur in sea otters (Williams and Van Blaricom, 1989). For otters with suspected myopathy or for prophylaxis in capture or stress situations, treatment with supplemental vitamin E (400 IU/day orally) and selenium (0.1 mg/kg once IM) has been recommended (Williams et al., 1995a). A herpes-like virus was associated with a high incidence of oral ulcerations in sea otters in oil spill rehabilitation centers in Alaska in 1989. Similar virus-associated ulcers were subsequently demonstrated to occur naturally in the sea otter population of Prince William Sound, with no apparent debility noted in affected otters (Harris et al., 1990). Serum antibodies to canine distemper virus were reported in two Kurilian sea otters harvested in 1990, but neither animal showed any clinical signs of viral disease (Birkun and Krivokhizhyn, 1991). Bacterial infections (see Chapter 16, Bacterial Diseases) may occur as a result of fight or predation wounds, damaged or infected teeth or gums, and from parasite migration or perforation. Injection site abscesses were seen in some animals during oil spill rehabilitation (Wilson et al., 1990) and following intramuscular injection of leuprolide acetate (Calle et al., 1997). Septic arthritis was reported by Wilson et al. (1990), apparently as a result of bite wounds to an otter’s hock. Fatal valvular endocarditis and thromboembolism have also been observed (Joseph et al., 1990; Tuomi, unpubl. obs.). Coccidiodomycosis has been reported as a cause of mortality in sea otters in California (Ames et al., 1983), where it was felt that exposure occurred through windborne spores in dust particles from endemic inshore sites. Fractures may occur secondary to falls from rocky haul-out areas, collisions with boats, gunshot wounds, natural predation, or interspecific aggression (see Chapter 23, Noninfectious Diseases). Healed fractures of the baculum are sometimes noted incidentally on post-mortem examination (Morejohn et al., 1975). The aquatic lifestyle of sea otters may allow many fractures to heal naturally as long as the animal can feed and groom adequately. A partially paralyzed female otter that presented to the oil spill rehabilitation center in Seward, Alaska, with a misaligned compression fracture of the lumbar vertebrae made a full recovery after the animal was fed and allowed to rest undisturbed in a large pool (Wilson et al., 1990). A variety of neoplastic conditions have been reported in sea otters (see Chapter 23, Noninfectious Diseases). Most are found at post-mortem and may be incidental, but heavy tumor load would predispose an otter to weight loss, poor grooming, hypoglycemia, starvation, or stress-induced gastroenteritis. Ante-mortem diagnosis should be attempted by standard techniques of palpation, radiography, ultrasonography, and biopsy. Treatment is theoretically possible in captive animals in selected cases using surgical excision, radiation, or intralesional chemotherapy.
Surgery Although there are few published reports of surgical procedures in sea otters other than implantation of radio transmitters, standard small-animal techniques for soft-tissue and orthopedic surgery can generally be employed. Shaving of the fur should be avoided as hair loss can create loss of insulation and compromise a debilitated animal. As an alternative, a water-soluble, disinfectant gel may be prepared by mixing equal parts of a sterile lubricating jelly with povidone–iodine solution and working the mixture into a small area of the coat. The hair can then be carefully parted to expose sufficient skin for surgical incisions or intravenous access.
0839_frame_C44 Page 980 Tuesday, May 22, 2001 2:09 PM
980
CRC Handbook of Marine Mammal Medicine
Uterine torsion (Rennie and Woodhouse, 1988; Tuomi and Williams, 1995), dystocia (Morejohn et al., 1975; Huff, pers. comm.), and intestinal volvulus or torsion (Williams et al., 1983) may potentially be corrected surgically, if diagnosed before organ damage is too severe. The surgical approach is a ventral midline incision similar to the technique used for implantation of abdominal radio transmitters in wild otters (Ralls et al., 1989).
Dentistry Dental caries, fractures, and abscesses are common in wild sea otters and have been associated with death due to starvation in older otters. Juvenile sea otters also experience higher mortality during immediate postweaning, when deciduous premolar and molars are being replaced by adult dentition. Periodontal disease has been reported in captive otters. Extraction of damaged teeth eliminates discomfort and reduces infection, but dental loss may reduce foraging ability. Young et al. (1999) reported on the use of long-acting zinc chlorhexidate gel, offered as ice cubes, for the long-term control and prevention of gingivitis and periodontal infections in captive northern sea otters.
Preventive Medicine Routine vaccination of captive sea otters for domestic animal diseases or rabies has not been reported in the United States. Schaftenaar (pers. comm.) has utilized commercially available canine distemper subunit vaccine, inactivated canine parvovirus vaccine, and leptospirosis bacterin in two sea otters housed at the Rotterdam Zoo in the Netherlands. Although other marine mammal and mustelid species have been shown to be susceptible to morbillivirus (Duignan, 1999), leptospirosis (Gulland et al., 1996), clostridial enteritis (Kollias, 1999), and rabies (Rupprecht, 1995), and may benefit from vaccination, these diseases have not been reported in sea otters and the efficacy of vaccination is unknown. Prophylactic use of ivermectin products has been used for prevention of heartworm (Dirofilaria immitis) infestation in endemic areas and appears to be safe and efficacious at canine dose rates.
Acknowledgments This chapter would not have been completed without the tremendous effort and contributions of Daniel M. Mulcahy, to whom it is dedicated. The author is sincerely grateful to Frances Gulland and Leslie Dierauf for entrusting her with the task of assembling this information, and to Jonna Mazet and Michael Murray for their reviews and comments.
References Ames, J.A., and Morejohn, G.V., 1980, Evidence of white shark, Carcharodon carcharias, attacks on sea otters, Enhydra lutris, Cal. Fish Game, 66: 196–209. Ames, J.A., Hardy, R.A., Wendell, F.E., and Geibel, J., 1983, Sea otter mortality in California, Report of the Marine Resources Division, California Department of Fish and Game, Monterey, 49 pp. Ames, J.A., Geibel, J., Wendell, F.E., and Pattison, C.A., 1996, White shark-inflicted wounds of sea otters in California, 1968–1992, in Great White Sharks: The Biology of Carcharodon carcharius, Pyle, P. (Ed.), Academic Press, New York, 309–316. Antonelis, G.A., Leatherwood, S., Cornell, L.H., and Antrim, J.G., 1981, Activity cycle and food selection of captive sea otters, Murrelet, 62: 6–9. Antrim, J.E., and Cornell, L.H., 1980, Reproduction of the sea otter, Int. Zoo Yearb., 20: 76–80.
0839_frame_C44 Page 981 Tuesday, May 22, 2001 2:09 PM
Sea Otters
981
Ballachey, B.E., Bodkin, J.L., and DeGange, A.R., 1994, An overview of sea otter studies, in Marine Mammals and the Exxon Valdez, Loughlin, T.R. (Ed.), Academic Press, San Diego, CA, 47–59. Ballachey, B.E., Stegeman, J., Snyder, P.W., Blundell, G.M., Bodkin, J.L., Dean, T.A., Duffy, L., Esler, D., Golet, G., Jewett, S. et al., in press, Final report to Exxon Valdez oil spill trustee council: Restoration project 99025, U.S. Geological Survey, Anchorage, AK. Barabash-Nikiforov, I.I., Marakov, S.V., and Nikolaev, A.M., 1947, The Kalan or Sea Otter, Academy of Sciences of the U.S.S.R., Moscow, 16–20. Birkun, A.A., Jr., and Krivokhizhyn, S.V., 1991, Pathomorphological and parasitological findings in sea otters from the Kuril and Commander Islands, presented at Third Joint U.S.S.R.–U.S. Sea Otter Conference, Petropavlovsk-Kamchatsky, September 9–15. Black, M., and Williams, T.D., 1993, Intraosseous infusion in the sea otter, in Proceedings of the International Association for Aquatic Animal Medicine, 24:12. Bodkin, J.L., Mulcahy, D., and Lensink, C.J., 1993, Age-specific reproduction in female sea otters (Enhydra lutris) from south-central Alaska: Analysis of reproductive tracts, Can. J. Zool., 71: 1811–1815. Bodkin, J.L., Ames, J.A., Jameson, R.J., Johnson, A.M., and Matson, G.M., 1997, Estimating age of sea otters with cementum layers in the first premolar, J. Wildl. Manage., 61: 967–973. Bodkin, J.L., Burdin, A.M., and Ryazanov, D.A., 2000, Age- and sex-specific mortality and population structure in sea otters, Mar. Mammal Sci., 16: 201–219. Bolin, R.L., 1938, Reappearance of the southern sea otter along the California coast, J. Mammal., 19: 301–303. Bowen, T.E., and Bellamy, R.F., 1988, Emergency War Surgery, U.S. Government Printing Office, Washington, D.C. Brennan, E.J. (Ed.), 1996, Conservation and management of the southern sea otter, Endangered Species Update, 13: 1–91. Brosseau, C., Johnson, M.L., Johnson, A.M., and Kenyon, K.W., 1975, Breeding the sea otter at Tacoma Aquarium, Int. Zoo Yearb., 15: 144–147. Bryant, H.C., 1915, Sea otters near Point Sur, Calif. Dep. Fish Game Bull., 1: 134–135. Calkins, D.G., and Lent, P.C., 1975, Territoriality and mating behavior in Prince William Sound sea otters, J. Mammal., 56: 528–529. Calle, P.P., Stetter, M.E., Raphael, B.L., Cook, R.A., McClave, C., Basinger, J.A., Walters, H., and Walsh, K., 1997, Use of depot leuprolide acetate to control undesirable male associated behaviors in the California sea lion (Zalophus californianus) and California sea otter (Enhydra lutris), in Proceedings of the 28th Annual International Association for Aquatic Animal Medicine, 28: 6–7. Chechowitz, M.A., Lowenstine, L.J., Gardner, I., Barr, B.C., Conrad, P.A., Gulland, F.M., and Jessup, D., 1999, Protozoal encephalitis in California sea otters and harbor seals: An update, in Proceedings of the 30th Annual International Association for Aquatic Animal Medicine, 30: 5. Costa, D.P., 1982, Energy, nitrogen, and electrolyte flux and sea-water drinking in the sea otter, Physiol. Zool., 55: 35–44. Costa, D.P., and Kooyman, G.L., 1982, Oxygen consumption, thermoregulation, and the effects of fur oiling and washing on the sea otter, Enhydra lutris, Can. J. Zool., 60: 2761–2767. Crissey, S.D., 1998, Handling Fish Fed to Fish-Eating Animals: A Manual of Standard Operating (Enhydra lutris) Procedures, U.S. Department of Agriculture, Agricultural Research Service, National Agricultural Library, Hyattsville, MD. Cronin, M.A., Bodkin, J., Ballachey, B., Estes, J., and Patton, J.C., 1996, Mitochondrial-DNA variation among subspecies and populations of sea otters (Enhydra lutris), J. Mammal., 77: 546–557. Dailey, M., and Mayer, K., 1999, Parasitic helminth (Acanthocephalan) infection as a cause of mortality in the California sea otter (Enhydra lutris), in Proceedings of the 30th Annual International Association for Aquatic Animal Medicine, 30: 126–127. Davis, J., and Lidicker, W.Z., Jr., 1975, The taxonomic status of the southern sea otter, in Proceedings of the California Academy of Sciences, Fourth Series, 40: 429–437.
0839_frame_C44 Page 982 Tuesday, May 22, 2001 2:09 PM
982
CRC Handbook of Marine Mammal Medicine
Davis, R.W., Williams, T.M., Thomas, J.A., Kastelein, R.A., and Cornell, L.H., 1988, The effects of oil contamination and cleaning on sea otters (Enhydra lutris). II. Metabolism, thermoregulation, and behavior, Can. J. Zool., 66: 2780–2790. DeGange, A.R., Doroff, A.M., and Monson, D.H., 1994, Experimental recovery of sea otter carcasses at Kodiak Island, Alaska, following the Exxon Valdez oil spill, Mar. Mammal Sci., 10: 492–496. Denison, D.M., and Kooyman, G.L., 1973, The structure and function of the small airways in pinniped and sea otter lungs, Respir. Physiol., 17: 1–10. Drabek, C.M., and Kooyman, G.L., 1984, Histological development of the terminal airways in pinniped and sea otter lungs, Can. J. Zool., 62: 92–96. Duignan, P.D., 1999, Morbillivirus infections of marine mammals, in Zoo and Wild Animal Medicine: Current Therapy, 4th ed., Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 497–501. Estes, J.A., 1980, Enhydra lutris, Mammal Species, 133: 1–8. Estes, J.A., 1990, Growth and equilibrium in sea otter populations, J. Anim. Ecol., 59: 385–401. Estes, J.A., Underwood, K., and Karmann, M., 1986, Activity time budgets of sea otters in California, J. Wildl. Manage., 50: 626–636. Estes, J.A., Doak, D.F., Bodkin, J.R., Jameson, R.J., Monson, D., Watt, J., and Tniker, M.T., 1996, Comparative demography of sea otter populations, in Endangered Species Update Special Issue: Conservation and Management of the Southern Sea Otter, Vol. 13, December, University of Michigan, Ann Arbor, 11–13. Estes, J.A., Tinker, M., Williams, T.M., and Doak, D.F., 1998, Killer whale predation on sea otters linking oceanic and nearshore ecosystems, Science, 282: 473–476. Fausett, J., 1976, Assimilation Efficiency of Captive Sea Otters, Enhydra lutris, M.A. thesis, California State University, Long Beach, 39 pp. Fisher, E.M., 1940, Early life of a sea otter pup, J. Mammal., 21: 132–137. Foott, J.O., 1971, Nose scars in female sea otters, J. Mammal., 51: 621–622. Gallo-Reynoso, J.-P., and Rathbun, G.B., 1997, Status of sea otters (Enhydra lutris) in Mexico, Mar. Mammal Sci., 13: 332–340. Garrott, R.A., Eberhardt, L.E., and Burn, D.M., 1993, Mortality of sea otters in Prince William Sound following the Exxon Valdez oil spill, Mar. Mammal Sci., 9: 343–359. Garshelis, D.L., 1984, Age estimation of living sea otters, J. Wildl. Manage., 48: 456–463. Garshelis, D.L., 1997, Sea otter mortality estimated from carcasses collected after the Exxon Valdez oil spill, Conserv. Biol., 11: 905–916. Garshelis, D.L., Garshelis, J.A., and Kimker, A.T., 1986, Sea otter time budgets and prey relationships in Alaska, J. Wildl. Manage., 50: 637–647. Gentry, R.L., and Peterson, R.S., 1967, Underwater vision of the sea otter, Nature, 216: 435–436. Geraci, J.R., and Williams, T.D., 1990, Physiological and toxic effects of oil on sea otters, in Sea Mammals and Oil: Confronting the Risks, Geraci, J.R., and St. Aubin, D.J. (Eds.), Academic Press, San Diego, CA, 211–221. Greenwalt, L., 1979, Determination that the southern sea otter is a threatened species, Fed. Reg., 42: 2965–2968. Gulland, F.M.D., Koski, M., Lowenstine, L.J., Colagross, 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. Hall, K.R.L., 1965, Tool-using behavior of the Californian sea otter, Med. Biol. Illus., 15: 216–217. Hall, K.R.L., and Schaller, G.B., 1964, Tool-using behavior of the California sea otter, J. Mammal., 45: 287–298. Harris, R.K., Moeller, R.B., Lipscomb, T.P., Haebler, R.J., Tuomi, P.A., McCormick, C.A., DeGange, A.R., Mulcahey, D., Williams, T.D., and Pletcher, J.M., 1990, Identification of a herpes-like virus in sea otters during rehabilitation after the T/V Exxon Valdez oil spill, in Sea Otter Symposium: Proceedings of a Symposium to Evaluate the Response Effort on Behalf of Sea Otters after the T/V Exxon Valdez oil spill into Prince William Sound, Anchorage, AK, April 17–19, Bayha, K., and Kormendy, J. (Eds.), U.S. Fish and Wildlife Service Biological Report, 90 (12): 366–368.
0839_frame_C44 Page 983 Tuesday, May 22, 2001 2:09 PM
Sea Otters
983
Hildebrand, M., 1954, Incisor tooth wear in the sea otter, J. Mammal., 35: 595. Jameson, R.J., 1983, Evidence of birth of a sea otter on land in central California, Calif. Fish Game, 69: 122–123. Jameson, R.J., 1999, Status of wild sea otters in California, Oregon and Washington, Sea Otter Conservation Workshop Report, February 20, Seattle Aquarium, Seattle, WA, 9 pp. Jameson, R.J., and Johnson, A.M., 1993, Reproductive characteristics of female sea otters, Mar. Mammal Sci., 9: 156–167. Jameson, R.L., and Bodkin, J.L., 1986, An incidence of twinning in the sea otter (Enhydra lutris), Mar. Mammal Sci., 2: 304–309. Johnson, A.M., 1982, Status of Alaska sea otter populations and developing conflicts with fisheries, in Transactions of the 47th North American Wildlife and Natural Resources Conference, 293–299. Joseph, B.E., Spraker, T.R., and Migaki, G., 1990, Valvular endocarditis in a northern sea otter (Enhydra lutris), J. Zoo Wildl. Med., 21: 88–91. Kenyon, K.W., 1969, The sea otter in the eastern Pacific Ocean, North American Fauna, U.S. Fish and Wildlife Service, Washington, D.C., 69 pp. Kenyon, K.W., Yunker, C.E., and Newell, I.M., 1965, Nasal mites (Halarachnidae) in the sea otter, J. Parasitol., 51: 960. Kollias, G.V., 1999, Health assessment, medical management, and prerelease conditioning of translocated North American river otters, in Zoo and Wild Animal Medicine: Current Therapy, 4th ed., Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 443–448. Kooyman, G.L., 1973, Respiratory adaptations in marine mammals, Am. Zool., 13: 457–468. Lenfant, C., Johansen, K., and Torrance, J.D., 1970, Gas transport and oxygen storage capacity in some pinnipeds and the sea otter, Respir. Physiol., 9: 277–286. Lensink, C.J., 1960, Status and distribution of sea otters in Alaska, J. Mammal., 41: 172–182. Lensink, C.J., 1962, The History and Status of Sea Otters in Alaska, Ph.D. dissertation, Purdue University, West Lafayette, IN, 188 pp. Levy, B., and Sivak, J.G., 1980, Mechanisms of accommodation in the bird eye, J. Comp. Physiol., 137: 267–272. Ling, J.K., 1970, Pelage and molting in wild mammals with special reference to aquatic forms, Q. Rev. Biol., 45: 16–54. Lipscomb,T.P., Harris, R.K., Moeller, R.B., Pletcher, J.M., Haebler, R.J., and Ballachey, B.E., 1993, Histopathologic lesions in sea otters exposed to crude oil, Vet. Pathol., 30: 1–11. Loughlin, T.R., 1977, Activity Patterns, Habitat Partitioning, and Grooming Behavior of the Sea Otter, Enhydra lutris, in California, Ph.D. dissertation, University of California, Los Angeles, 110 pp. Loughlin, T.R., Ames, J.A., and Vandevere, J.E., 1981, Annual reproduction, dependancy period, and apparent gestation period in two California sea otters, Enhydra lutris, Fish. Bull., 79: 347–348. Lyons, K.J., 1991, Behavior patterns of individual sea otters: Relatedness and resource sharing, presented at 9th Biennial Conference on the Biology of Marine Mammals, Chicago, IL, 43. Mazet, J.A., Gardner, I.A., and Lowenstine, L.J., 1995, Reproductive effects of petroleum product exposure on American mink Mustela vison as a laboratory model for sea otters Enhydra lutris, in Proceedings of the 1995 Joint Conference, American Association of Zoo Veterinarians, Wildlife Disease Association, and American Association of Wildlife Veterinarians, East Lansing, MI, August 12–17, 41–42. Monson, D.H., and DeGange, A.R., 1995, Reproduction, preweaning survival, and survival of adult sea otters at Kodiak Island, Alaska, Can. J. Zool., 73: 1161–1169. Monson, D.H., McCormick, C., and Ballachey, B.E., in press, Chemical restraint of northern sea otters: Results of past field studies, J. Zoo Wildl. Med. Morejohn, G.V., Ames, J.A., and Lewis, D.B., 1975, Post mortem studies of sea otters, Enhydra lutris, Marine Resources Technical Report, California Department of Fish and Game, 30: 1–84. Morrison, P., Rosenmann, M., and Estes, J.A., 1974, Metabolism and thermoregulation in the sea otter, Physiol. Zool., 47: 218–229.
0839_frame_C44 Page 984 Tuesday, May 22, 2001 2:09 PM
984
CRC Handbook of Marine Mammal Medicine
Murphy, C.J., Bellhorn, R.W., Williams, T., Burns, M.S., Schaffel, F., and Howland, H.C., 1990, Refractive state, ocular anatomy, and accommodative range of the sea otter (Enhydra lutris), Vision Res., 30: 23–32. Newby, T.C., 1975, A sea otter (Enhydra lutris) food dive record, Murrelet, 56: 19. Nightingale, J.W., 1981, Elements of a successful breeding program with captive sea otters, in Proceedings of the American Association of Zoo Veterinarians, Seattle, WA, 84–86. Ogden, A., 1941, The California Sea Otter Trade, 1784–1848, University of California Press, Berkeley, 251 pp. Packard, J.M., and Ribic, C.A., 1982, Classification of the behavior of sea otters (Enhydra lutris), Can. J. Zool., 60: 1362–1373. Payne, S.F., and Jameson, R.J., 1984, Early behavioral development of the sea otter, Enhydra lutris, J. Mammal., 65: 527–531. Ralls, K., Ballou, J., and Brownell, R.L., Jr., 1983, Genetic diversity in California sea otters: Theoretical considerations and management implications, Biol. Conserv., 25: 209–232. Ralls, K., Siniff, D.B., Williams, T.D., and Kuechle, V.B., 1989, An intraperitoneal radio transmitter for sea otters, Mar. Mammal Sci., 5: 376–381. Rausch, R., 1953, Studies on the helminth fauna of Alaska, XIII, disease in the sea otter, with special reference to helminth parasites, Ecology, 34: 584–604. Rebar, A.H., Lipscomb, T.P., Harris, R.K., and Ballachey, B.E., 1995, Clinical and clinical laboratory correlates in sea otters dying unexpectedly in rehabilitation centers following the Exxon Valdez oil spill, Vet. Pathol., 32: 346–350. Rennie III, C.J., and Woodhouse, C.D., 1988, Scoliosis and uterine torsion in a pregnant sea otter (Enhydra lutris) from California, J. Wildl. Dis., 24: 582–584. Riedman, M.L., and Estes, J., 1988, A review of the history, distribution and foraging ecology of sea otters, in The Community Ecology of Sea Otters, Van Blaricom, G.R., and Estes, J.A. (Eds.), SpringerVerlag, Berlin, 4–21. Riedman, M.L., and Estes, J., 1990, The sea otter (Enhydra lutris): Behavior, ecology, and natural history, U.S. Fish and Wildlife Service, Biological Report, 90: 126. Roest, A.I., 1973, Subspecies of the sea otter, Enhydra lutris, Contr. Sci., 252: 1–17. Roest, A.I., 1976, Systematics and the status of sea otters, Enhydra lutris, Bull. S. Calif. Acad. Sci., 75: 267–270. Rosonke, B.J., Brown, S.R., Tornquist, S.J., Snyder, S.P., Garner, M.M., and Blythe, L.L., 1999, Encephalomyelitis associated with a Sarcocystis neurona-like organism in a sea otter, J. Am. Vet. Med. Assoc., 215: 1839–1842. Rotterman, L.M., and Simon-Jackson, T., 1988, Sea otter, in Selected Marine Mammals of Alaska: Species Accounts with Research and Management Recommendations, Lentfer, J.W. (Ed.), Marine Mammal Commission, Washington, D.C., 237–275. Rupprecht, C.E., 1995, Rabies: Global problem, zoonotic threat, and preventative management, in Zoo and Wild Animal Medicine: Current Therapy, 4th ed., Fowler, M.E., and Miller, R.E. (Eds.), W.B. Saunders, Philadelphia, 136–146. Sanchez, M.S., 1992, Differentiation and Variability of Mitochondrial DNA in Three Sea Otter, Enhydra lutris, Populations, Ph.D. dissertation, University of California, Santa Cruz, 101 pp. Sandegren, F.E., Chu, E.W., and Vandevere, J.E., 1973, Maternal behavior in the California sea otter, J. Mammal., 54: 668–679. Sawyer, D.C., and Williams, T.D., 1996, Chemical restraint and anesthesia of sea otters affected by the oil spill in Prince William Sound, Alaska, J. Am. Vet. Med. Assoc., 208: 1831–1834. Scheffer, V.B., 1951, Measurements of sea otters from western Alaska, J. Mammal., 32: 10–14. Scheffer, V.B., 1964, Estimating abundance of pelage fibres on fur seal skin, Proc. Zool. Soc. London, 143: 37–41. Schneider, K.B., 1972, Reproduction in the Female Sea Otter, Project Progress Report, Federal Aid in Wildlife Restoration Project W-17-4, Alaska Department of Fish and Game, 36 pp.
0839_frame_C44 Page 985 Tuesday, May 22, 2001 2:09 PM
Sea Otters
985
Schneider, K.B., 1973, Reproduction in the Female Sea Otter, Project Progress Report, Federal Aid in Wildlife Restoration Project W-17-4 and W-17-5, Alaska Department of Fish and Game, 13 pp. Schneider, K.B., 1978, Sex and Age Segregation of Sea Otters, Federal Aid in Wildlife Restoration Projects W-17-4, W-17-5, Project Progress Report, Alaska Department of Fish and Game, 45 pp. Schneider, K.B., and Faro, J.B., 1975, Effects of sea ice on sea otters (Enhydra lutris), J. Mammal., 56: 91–101. Schusterman, R.J., and Barrett, B., 1973, Amphibious nature of visual acuity in the Asian “clawless” otter, Nature, 244: 518–519. Sherrod, S.K., Estes, J.A., and White, C.M., 1975, Depredation of sea otter pups by bald eagles at Amchitka Island, Alaska, J. Mammal., 56: 701–703. Simenstad, C.A., Estes, J.A., and Kenyon, K.W., 1978, Aleuts, sea otters, and alternate stable-state communities, Science, 200: 403–411. Sinha, A.A., and Conaway, C.H., 1968, The ovary of the sea otter, Anat. Rec., 160: 795–806. Sinha, A.A., Conaway, C.H., and Kenyon, K.W., 1966, Reproduction in the female sea otter, J. Wildl. Manage., 30: 121–130. Siniff, D.B., and Ralls, K., 1991, Reproduction, survival and tag loss in California sea otters, Mar. Mammal Sci., 7: 211–229. Sivak, J.G., 1980, Accommodation in vertebrates: A contemporary survey, Curr. Top. Eye Res., 3: 281–330. Spelman, L.H., Jochem, W.J., Sumner, P.W., Redmond, D.P., and Stoskopf, M.K., 1997, Postanesthetic monitoring of core body temperature using telemetry in North American river otters (Lutra canadensis), J. Zoo Wildl. Med., 28: 413–417. Staedler, M.M., and Riedman, M.L., 1989, A case of adoption in the California sea otter, Mar. Mammal Sci., 5: 391–394 Staedler, M., and Riedman, M., 1993, Fatal mating injuries in female sea otters (Enhydra lutris nereis), Mammalia, 57: 135–139. Stoskopf, M.K., Spelman, L.H., Sumner, P.W., Redmond, D.P., Jochem, W.J., and Levine, J.F., 1997, The impact of water temperature on core body temperature of North American river otters (Lutra canadensis) during simulated oil spill recovery washing protocols, J. Zoo Wildl. Med., 28: 407–412. Stullken, D.E., and Kirkpatrick, C.M., 1955, Physiological investigation of captivity mortality in the sea otter (Enhydra lutris), in Transactions of the Twentieth North American Wildlife Conference, 476–494. Sweeney, J.C., 1965, Common diseases of pinnipeds, J. Am. Vet. Med. Assoc., 147: 1090. Tarasoff, F.J., 1974, Anatomical adaptations in the river otter, sea otter, and harp seal with reference to thermal regulation, in Functional Anatomy of Marine Mammals 2, Harrison, R.J. (Ed.), Academic Press, New York, 111–141. Tarasoff, F.J., and Kooyman, G.L., 1973a, Observations on the anatomy of the respiratory system of the river otter, sea otter, and harp seal. I. The topography, weight, and measurements of the lungs, Can. J. Zool., 51: 163–170. Tarasoff, F.J., and Kooyman, G.L., 1973b, Observations on the anatomy of the respiratory system of the river otter, sea otter, and harp seal. II, The trachea and bronchial tree, Can. J. Zool., 51: 171–177. Taylor, W.P., 1914, The problem of aquatic adaptation in the Carnivora, as illustrated in the osteology and evolution of the sea-otter, University of California, Geology, 7: 465–495. Thomas, N.J., and Cole, R.A., 1996, The risk of disease and threats to the wild population, in Special Issue: Conservation and Management of the Southern Sea Otter, Endangered Species Update, University of Michigan, Ann Arbor, 13: 23–27. Tuomi, P.A., 1990, Husbandry at the Valdez Otter Rehabilitation Center, in Sea Otter Symposium: Proceedings of a Symposium to Evaluate the Response Effort on Behalf of Sea Otters after the T/V Exxon Valdez Oil Spill into Prince William Sound, Anchorage, AK, April 17–19, Bayha, K., and Kormendy, J. (Eds.), U.S. Fish and Wildlife Service Biological Report, 90: 274–284. Tuomi, P., and Burek, K., 1999, Septic peritonitis in an adult northern sea otter (Enhydra lutris) secondary to perforation of gastric parasitic ulcer, in Proceedings of the 30th Annual International Association for Aquatic Animal Medicine, 30: 63–64.
0839_frame_C44 Page 986 Tuesday, May 22, 2001 2:09 PM
986
CRC Handbook of Marine Mammal Medicine
Tuomi, P.A., and Williams, T.M., 1995, Rehabilitation of pregnant sea otters and females with newborn pups, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving FurBearing Marine Mammals, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 121–132. Tuomi, P.A., Donoghue, S., and Otten-Stanger, J.M., 1995, Husbandry and nutrition, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-Bearing Marine Mammals, Williams, T.M., and Davis, R.W. (Eds.) University of Alaska Press, Fairbanks, 103–119. U.S. Department of Agriculture, 1995, Animal and Plant Health Inspection Service Rules and Regulations, Marine Mammals: Humane Handling, Care, Treatment and Transportation, Subchapter AAnimal Welfare, Code of Federal Regulations, 9: 84–101. Van Blaricom, G.R., 1988a, Effects of foraging by sea otters on mussel-dominated intertidal communities, in The Community Ecology of Sea Otters, Van Blaricom, G.R., and Estes, J.A. (Eds.), Springer-Verlag, Berlin, 48–91. Van Blaricom, G.R., 1988b, Concentrations of fecal coliform bacteria associated with housing of wild sea otters at the Monterey Bay Aquarium, summer and fall 1987, unpublished report, U.S. Fish and Wildlife Service, Santa Cruz, CA. Wendell, F.E., Ames, J.A., and Hardy, R.A., 1984, Pup dependency period and length of reproductive cycle: Estimates from observations of tagged sea otters, Enhydra lutris, in California, Calif. Fish Game, 70: 89–100. Wild, P.W., and Ames, J.A., 1974, A report on the sea otter, Enhydra lutris L., in California, California Department of Fish and Game, Marine Research Technical Report, 20: 93. Williams, T.D., 1990, Sea otter biology and medicine, in Handbook of Marine Mammal Medicine: Health, Disease and Rehabilitation, Dierauf, L.A. (Ed.), CRC Press, Boca Raton, FL, 625–648. Williams, T.D., and Kocher, F.H., 1978, Comparison of anesthetic agents in the sea otter, J. Am. Vet. Med. Assoc., 173: 1127–1130. Williams, T.D., and Pulley, L.T., 1983, Hematology and blood chemistry in the sea otter (Enhydra lutris), J. Wildl. Dis., 19: 44–47. Williams, T.D., and Sawyer, D.C., 1995, Physical and chemical restraint, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-Bearing Marine Mammals, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 39–43. Williams, T.D., and Van Blaricom, G.D., 1989, Rates of capture myopathy in translocated sea otters, with implications for management of sea otter rescue following oil spills, in Proceedings from the 8th Biennial Conference on the Biology of Marine Mammals, Pacific Grove, CA, 72. Williams, T.D., Mattison, J.A., and Ames, J.A., 1980, Twinning in a California sea otter, J. Mammal., 61: 575–576. Williams, T.D., Hoefler, L., and Pinard, W., 1983, Pneumoperitoneum associated with intestinal volvulus in a sea otter, J. Am. Vet. Med. Assoc., 183: 1288–1289. Williams, T.D., Baylis, D.M., Downey, S.H., and Clark, R.O., 1990, A physical restraint device for sea otters, J. Zoo Wildl. Med., 21: 105–107. Williams, T.D., Allen, D.D., Groff, J.M., and Glass, R.L., 1992, An analysis of California sea otter Enhydra lutris pelage and integument, Mar. Mammal Sci., 8: 1–18. Williams, T.M., and Davis, R.W. (Eds.), 1995, Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-Bearing Marine Mammals, University of Alaska Press, Fairbanks. Williams, T.M., Davis, R.W., McBain, J.F., Tuomi, P.A., Wilson, C.R., and Donoghue, S., 1995a, Diagnosing and treating common clinical disorders of oiled sea otters, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-Bearing Marine Mammals, Williams, T.M., and R.W. Davis, (Eds.), University of Alaska Press, Fairbanks, 59–94. Williams, T.M., O’Connor, D.J., and Nielsen, S.W., 1995b, The effects of oil on sea otters: Histopathology, toxicology, and clinical history, in Emergency Care and Rehabilitation of Oiled Sea Otters: A Guide for Oil Spills Involving Fur-Bearing Marine Mammals, Williams, T.M., and Davis, R.W. (Eds.), University of Alaska Press, Fairbanks, 3–22.
0839_frame_C44 Page 987 Tuesday, May 22, 2001 2:09 PM
Sea Otters
987
Wilson, D.E., Bogan, M.A., Brownell, R.L., Jr., Burdin, A.M., and Maminov, M.K., 1991, Geographic variation in sea otters Enhydra lutris, J. Mammal., 72: 22–36. Wilson, R.K., Tuomi, P.A., Schroeder, J.P., and Williams, T.D., 1990, Clinical treatment and rehabilitation of oiled sea otters, in Sea Otter Rehabilitation Program: 1989 Exxon Valdez Oil Spill Report to Exxon Company, USA, International Wildlife Research, Williams, T.M., and Davis, R.W. (Eds.), 101–117. Young, S.J.F., Huff, D.G., and Anthony, J.M.G., 1999, A safe and cost effective oral care regime to control gingivitis and periodontal disease in the northern sea otter (Enhydra lutris lutris), in Proceedings of the 30th Annual International Association for Aquatic Animal Medicine, 30: 67.
0839_frame_C44 Page 988 Tuesday, May 22, 2001 2:09 PM
0839_frame_C45 Page 989 Tuesday, May 22, 2001 2:10 PM
45 Polar Bears Michael Brent Briggs
Introduction The polar bear (Ursus maritimus) is unique among the Family Ursidae, because it is adapted to a highly specialized, semiaquatic lifestyle. Even though a bear, it is classified as a marine mammal because of the intimate link between the animal and the sea. Polar bears may spend as much as 50% of their lives on ice floes and have been known to swim as far as 90 km (60 miles) from floe to floe. The polar bear is the largest terrestrial predator and is classified as a threatened species (Reynolds and Rommel, 1999) (see Chapter 33, Legislation, Table 1). A number of polar bears are maintained in zoological collections; thus, a basic understanding of their biology and natural history is needed to provide adequate care. This chapter presents current information on the biology, husbandry, and physiology of this unique animal. With sound veterinary medical protocols and clinical experience, these animals may be long-lived and hearty members of any zoological collection.
Natural History and Physiology The polar bear is a member of the Order Carnivora, Family Ursidae, genus Ursus, and species maritimus. It shares a common evolutionary ancestor, U. etruscus, with the brown bear (U. arctos) (Stirling, 1988). Polar bears vary in color from white as youngsters to yellowish brown as they age. The range of the polar bear extends throughout the circumpolar Arctic, including Canada, Greenland, Denmark, Norway, Russia, and the United States. Polar bears tend to live at the edge of the ice pack, moving south as winter approaches, following ice formation, and then north again when the ice begins to melt in the spring. Other populations move ashore during summer. Male polar bears tend to live near the coast during the winter, whereas females may move inland with their cubs and for denning. The bears have several unique adaptations for life on the polar sea ice. These include short tails and small ears to help reduce heat loss, thick fat layers of up to 11.5 cm (∼3 in.), and completely furred bodies (aside from nose and foot pads), to protect them from temperatures that can drop to −45°C (−49°F) during the severe cold of winter. Polar bear hair is of two types: (1) short, white, dense fur close to the body; and (2) long, transparent, hollow guard hairs that help keep the fur from matting while the animal is in the water. The bear’s fat also aids in buoyancy for swimming. The skin of polar bears is black, which tends to absorb more radiant heat. Unlike other bears, the polar bear has a small, streamlined head and an elongated body that makes it more streamlined for swimming.
0-8493-0839-9/01/$0.00+$1.50 © 2001 by CRC Press LLC
989
0839_frame_C45 Page 990 Tuesday, May 22, 2001 2:10 PM
990
CRC Handbook of Marine Mammal Medicine
There is much variation in the sizes reported for polar bears, but all agree that there is extreme sexual dimorphism, with the adult male being as much as 250% larger than the female. The average adult male weighs 350 to 600 kg (770 to 1320 lb), with a length of 2.5 to 3.0 m (8 to 10 ft). The average female weighs 150 to 300 kg (330 to 660 lb) with a length of up to 2.5 m (∼8 ft). In extreme cases, male bears have been recorded to weigh as much as 800 kg (1760 lb) in the wild, but this is rare (Stirling, 1988). Accurate field weights are difficult to obtain, and morphometry is somewhat inaccurate in the polar bear and other large mammals (Cattet et al., 1997a). In captivity, these weights may vary greatly. The average rectal temperature of captive polar bears is 37.5°C (99.6°F), with a range of 37.2 to 38.4°C (99 to 101°F). Free-ranging polar bears, at about 36.5°C, have a slightly lower body temperature than captive bears. The normal resting pulse rate is 60 to 90 beats/min, and the respiratory rate is 15 to 30 respirations/min. The body temperature, pulse rate, and respiratory rates vary with certain factors, including activity level, state of excitement, ambient temperature, and age (Lee et al., 1981; Wallach and Boever, 1983; Matthews, 1993). Polar bears are better adapted to retain heat rather than to dissipate it. When ambient temperatures increase, the bears are known to lie spread-legged on ice to help conduct heat away from their bodies. They may also simply stop walking, and sit or lie down. When on land, they dig small depressions that enable them to lie next to the cool earth (Oritsland, 1970; Best, 1982). Free-ranging bears are known to travel great distances. Once thought to be random travelers, they are actually mostly territorial and consist of distinct subpopulations with ranges determined by weather, ice, and food conditions (Stirling, 1988). The size of the range varies, and appears to be directly related to the amount of food available within the range. The average 2 2 size of a feeding range is approximately 500 km ; however, ranges as large as 5000 km have been recorded. Young bears may travel 1000 km (∼645 miles) from their dam before establishing their own range. In food-dense regions, ranges tend to overlap. Polar bears do not hibernate, but they can go into a dormancy state called denning. During this period the bears’ heart rate slows, but their body temperatures do not decrease as they would during hibernation (Stirling, 1988). Although polar bears do not hibernate, the bears have been shown to contain hibernation induction trigger (HIT) in their blood at levels consistent with levels in black bears that do hibernate (Bruce et al., 1990). Polar bears are metabolically able to sustain long periods without eating. In the late summer and fall, when food supply is scarce, they have been known to survive more than a month with virtually no food. Some people refer to this with the misnomer “walking hibernation” as these polar bears have urea and creatinine levels similar to those of hibernating black bears (Stirling, 1988). The female in a den with cubs, and in times of food shortage, can control urea recycling, and thus can survive extended periods of fasting (Stirling, 1974; Ramsay et al., 1991). The life span of a polar bear in captivity is approximately 25 years, but several captive bears have been reported to live well into their 30s. Life span in the wild is shorter, but accurate figures are unknown (Latinen, 1987).
Nutrition Free-ranging polar bears are seasonal hunters, with ringed seals (Phoca hispida) their primary food source. These seals live mostly on solid ice or land, but can be found on ice floes. Other prey includes bearded seals (Erignathus barbatus), other smaller mammals, and beach carrion. Polar bears hunt most often during the months of November through June, when seals are abundant and the ice is good for hunting. They use a variety of methods for obtaining their prey. The most common method is that the bear waits by a breathing hole in the ice; when
0839_frame_C45 Page 991 Tuesday, May 22, 2001 2:10 PM
Polar Bears
991
the seal comes up to breathe, the bear grabs it. Another method is that the bear patiently crawls toward a seal sleeping near a crack in the ice. If the seal is roused, the bear stops its movement until the seal resettles. When close enough, the bear springs to the attack (Ovsyanikov, 1996). Although the mainstay of the polar bear diet is generally ringed seals, polar bears have also been observed killing and eating belugas (Delphinapterus leucas) in the Bering and Chukchi Seas (Lowry et al., 1987), walrus (Odobenus rosmarus) calves (Ovsyanikov, 1996), and little auk (Alle alle) (Stempniewicz, 1993). Most adult polar bears only eat the skin and blubber of their prey, leaving the muscle and organs (Nelson, 1983). The metabolism of blubber conserves water, since it produces less urea than a high-protein meat diet. Eating blubber decreases the need for an external source of water, which can be difficult to attain in deep winter; this diet also helps with thermoregulation by decreasing the need to ingest snow for hydration. Polar bears’ cholesterol levels are lower in animals eating a strict seal blubber diet than in fasting bears. This difference is likely due to the protective quality of the omega-3 fatty acids contained in seal blubber. Young, growing animals and lactating females that have higher protein requirements generally eat whole carcasses. On average, an adult polar bear will kill one 65 kg (∼140 lb) ringed seal every 5 or 6 days during peak hunting season (Stirling, 1974). The vitamin A content of polar bear liver is higher than in any other mammal, and varies seasonally. In midsummer, the vitamin A content is highest at 22,000 to 29,000 IU/g liver, whereas in winter levels are 13,000 to 18,000 IU/g liver. These ranges mirror that of the quantity in the seals being consumed (Lewis and Lentfer, 1967; Leighton et al., 1988). Vitamin A in the liver is stored primarily in the ester form (98%). The esters are retinyl palmitate (37.3%), retinyl oleate (20.9%), stearate (12.8%), and linoleate (7.7%) (Ball et al., 1986). In sharp contrast to the seasonal variation of diet in free-ranging bears, captive bears are commonly fed a consistent daily ration of commercial omnivore pellets or dog food. At least 60% of the diet, by weight, should consist of dry food. These diets are generally supplemented with a variety of other food items, both for nutritional diversity and behavioral enrichment. As much as 40% of the diet may include fruits, vegetables, and fish. If animals are being fed fish, thiamin should be supplemented at a rate of 25 mg/kg of food eaten and vitamin D supplemented at 1000 IU/kg feed. Both freshwater (trout) and marine (herring, capelin) fish have been fed with no apparent problems. The total diet is based on a variety of factors, including age, season, weight, and activity level (see Chapter 36, Nutrition). At this juncture, a diet based on kcal/kg of body weight is not often used for feeding these animals but will likely be developed in the near future. The Brookfield Zoo (Illinois) polar bear diet currently consists of a commercially prepared dry diet, together with vegetables, fruit, fish, and prepared meat diets (Slifka, pers. comm.).
Nutrition of Juveniles, Early Pregnant, and Lactating Females The nutritional requirements of young, pregnant, and lactating female polar bears should resemble that of any other species (see Chapter 36, Nutrition). Literature is sparse, but logic should prevail. Since it is known from literature on the free-ranging population that pregnant and nursing females and young bears will eat entire carcasses (cubs will eat carcasses once the female has removed the blubber), thus significantly increasing their protein intake over the strict blubber diet, one can deduce that growing and lactating bears have higher protein requirements (Stirling, 1988). The female bear’s milk composition changes over the 18 to 24 months it nurses the cub. The fat content is highest upon emerging from the den (35.8%) and then decreases to weaning the following autumn (20.6%) for land bears, whereas there is no change in the composition of the sea ice bears’ milk (Derocher et al., 1993). During these periods,
0839_frame_C45 Page 992 Tuesday, May 22, 2001 2:10 PM
992
CRC Handbook of Marine Mammal Medicine
young captive animals must have the calories of their diets, as well as the percentage of protein on a dry matter (DM) basis, increased.
Infants Infant bears are difficult to hand-rear; methods used for polar bears are described in Chapter 37 (Hand-Rearing).
Geriatrics Captive polar bears can experience increased longevity compared with their free-ranging counterparts. Thus, it is important to be conscious of the dietary needs of geriatric animals in the zoo environment. To guard against obesity, decreases in activity levels should correspond to decreases in the calories offered to those animals. Many older animals lose the ability to excrete excess nitrogenous waste properly, so a reduction of the protein concentration in the diet may be necessary. In geriatric bears with worn teeth, digestibility of the diet may require improvement to avoid the need to offer increased nutrients to the animals. Many older bears develop arthritis, and this too can lead to decreased activity levels.
Reproduction Polar bears are considered mature between 4 and 8 years old. In the wild, they congregate during the breeding season (which lasts from late March to May) and during the fall (although with some regional variation). Polar bears are polygamous. The fertilized ova do not implant until fall. At that time, the female will build a den, which contains two chambers, in the snow. Denning follows a time of heavy feeding. Since the female undergoes delayed implantation and the ova does not implant until September or October, the gestation period is between 195 and 265 days (Boyd et al., 1999). Bears give birth in November/December and remain in the den until March or April (Stirling, 1988). The polar bear has a zonary endotheliochorial placenta. Dystocia is unlikely due to the size of the neonate and the low birth weight (600 to 700 g) (1 to 1.5 lb) compared with the size of the female; dystocia has not been recorded in the literature. As many as four altricial cubs can be born, with their eyes closed; twins are most common. It takes as long as a month for the cubs to open their eyes. In captive situations, the female will often become anorexic prior to denning. When the animals emerge from the den, the young weigh around 10 kg (22 lb) each. The mother will take care of the cubs for up to 28 months. The length of care is dependent upon weather conditions and the age of the female. The bears living farther north tend to be with their young longer than those that live farther south (Ramsay and Stirling, 1988).
Endocrinology Reproductive Hormones Endotheliochorial placentation in association with corpus luteal (CL) growth leads to an increase of circulating progesterones during pregnancy. It has been hypothesized that serum progesterone concentrations rise rapidly at conception in the spring (to ∼5 ng/ml); furthermore, this concentration of progesterone is maintained throughout the preimplantation period. With implantation in autumn, there is an additional twofold to threefold increase in serum progesterone as the CL undergoes greater vascularization with the implantation event (Lono, 1972). Nonpregnant females appear to have consistently low serum progesterone levels of 0.1 to
0839_frame_C45 Page 993 Tuesday, May 22, 2001 2:10 PM
Polar Bears
993
0.2 ng/mL. With this information, one can predict pregnancy in the polar bear with some degree of accuracy. Females begin their estrous cycling at 3 to 4 years of age and may, in the wild, produce cubs throughout their entire life span. Reproductive success in captivity has not been duplicated to this degree. Progesterone levels are lowest in the very young and very old animals, but still are adequate to maintain pregnancy. Of wild pregnant bears in the Hudson Bay region, only about 67% have viable cubs 1 year after the diagnosis of pregnancy. This figure could be due to a variety of factors, such as neonatal mortality, failure to implant, and abortion (Derocher et al., 1992). Testosterone levels in male bears show seasonal variation. These hormone levels are highest in the spring, just as in black bears. There is a strong correlation with photoperiod, but not with metabolic state. There is a corresponding increase in testicular size with the increase in testosterone, as well as a direct correlation with increasing age (Palmer et al., 1988).
Thyroid Hormones The plasma concentrations of thyroid hormones, triiodo-1-thyronine (T3) and 1-thyroxine (T4), vary with gender and age (see Chapter 10, Endocrinology). The adult males tend to have lower levels of both T3 and T4 than adult females. The adult male values average 0.68 ng/ml (T3) and 32.0 ng/ml (T4), whereas the females are at 1.23 and 45.0 ng/ml, respectively. In juveniles, the values are reversed, with the males having higher values than the females (Leatherland and Ronald, 1981).
Housing There are minimum requirements dictated by the U.S. Department of Agriculture for housing and maintaining polar bears (USDA, 1984). The minimum requirements include provision of a pool of water, a dry resting and/or social area, and a den (see Chapter 33, Legislation). The current specific requirements for pool size are a minimum horizontal dimension (MHD) 2 2 of not less than 2.44 m (8.0 ft), a surface area of not less than 8.93 m (96.0 ft ), and a minimum depth of 1.52 m (5 ft). The regulations do not specify a specific shape, but minimum dimensions must be maintained, and normal horizontal and vertical space must exist to allow for normal posture and movement. These dimensions are for one or two bears; for each additional bear, 2 2 the surface area must be increased by at least 3.72 m (40 ft ). Any area of the pool that does not meet the 5-ft-depth requirement cannot be used to provide the additional area. 2 2 The dry resting and social activity area must be at least 37.16 m (400 ft ). Each additional 2 2 bear is required to have an additional 3.72 m (40 ft ). There must also be enough shade in this area to provide coverage for all bears in the enclosure. The den is essential for pregnant females and must be at least 1.63 × 1.63 m (6 × 6 ft) and 1.52 m (5 ft) in height. Each female must be provided with its own den. The den area is a secluded area that will minimize any noise and visualization of humans and other bears, especially the male who may be housed nearby. There is no need to make a den that is too large, as large dens are not effective at providing the proper environment for successful cub rearing (Wemmer, 1974). Natural dens often have two chambers, but these do not appear to be necessary, as long as the den provides the female with security and quiet. The female should be checked throughout the denning period. Electronic surveillance equipment assists in maintaining a quiet environment, while allowing for frequent checks of the female and the cubs. The chamber should be bedded with clean, dry straw before the female dens, but should not be disturbed once the animal accepts the den.
0839_frame_C45 Page 994 Tuesday, May 22, 2001 2:10 PM
994
CRC Handbook of Marine Mammal Medicine
These are human-derived minimum standards for captivity, and are not necessarily considered adequate for a captive animal’s comfort and space, particularly considering the immense spaces that polar bears utilize in the wild.
Behavior Polar bears are solitary, migratory animals that generally have little association with their own species, except for times of extremely plentiful food supplies, during mating season, or when a female is taking care of cubs. Cannibalism has been observed in polar bears (Furr and Stenhaus, 1985). It is therefore appropriate to house females with cubs alone for up to 2.5 years, or animals without cubs entirely alone. Polar bears are prone to stereotypic behaviors, such as pacing in captivity, which has been documented on a number of occasions. From 55 to 100% of captive polar bears exhibit some sort of stereotypic behavior, of which pacing is the most common. This pacing may be related to the fact the animals walk long distances in their free-ranging existence. Behavioral enrichment programs have helped to decrease this stereotypy (Ames, 1993). Fluoxetine (Prozac, Eli Lilly & Co., Indianapolis, IN) at a dose of 1 mg/kg, orally, once a day (SID), may reduce or eliminate some stereotypic behavior without affecting normal behavior such as eating, sleeping, other motor activities, and interactions with other animals and keepers. This drug is a second-generation antidepressant, and its use should be monitored carefully as there is variation in its effects between patients (Poulsen et al., 1996; Teskey et al., 1996). Introductions of new mates can be difficult because of aggression between the two animals. The introduction of two individuals can be accomplished in a variety of ways using “creep doors” or those where the animals can see, hear, and smell one another for period of time before actually being placed in the same enclosure. When introduced, escape routes need to be made available for the smaller bear. Other methods of introduction involve the use of oral or injectable tranquilizers, such as diazepam (Valium, Abbott Laboratories, Chicago, IL), midazolam HCl (Versed, Roche Pharmaceuticals, Puerto Rico), or acepromazine maleate (Acepromazine, Boehringer Ingelheim, Burlington, Ontario, Canada). The dosing and route of administration are variable and depend on the nature of the animals and many other factors, such as time of year, age, previous association with other animals, and, most importantly, the sexual receptivity of the female to a male (Wemmer et al., 1976). There are many anecdotal reports of trauma inflicted by mother polar bears on their young, possibly inducing fractures (Cook et al., 1994; Briggs, unpubl. data). However, there have been observations of free-ranging female bears adopting cubs from other females (Derocher and Wiig, 1999). Thus, it is believed that if proper facilities are made for the denning mother, the likelihood of problems is greatly reduced. It is important to maintain proper standards to minimize any disturbance, not only during the denning time, but also during the perinatal period.
Physical Examination The physical examination is performed on the polar bear as on any other carnivore. Except in the case of very young, hand-reared animals, the examination must be performed while the animal is anesthetized (see below). An examination should be performed on a predetermined schedule, or when dictated by a clinical disease. The examination schedule should be modified to fit the facility housing the bear and the frequency of disease under observation. An examination should always be performed before shipment of an animal and during the quarantine period of a new arrival.
0839_frame_C45 Page 995 Tuesday, May 22, 2001 2:10 PM
Polar Bears
995
The examination should include the collection of basic biological parameters, and also focus on any specific complaints or maladies noted prior to the examination. Once immobilized, the animal should be secured so that movements will not threaten the veterinarians and technicians involved in the procedure. This can easily be done by the use of a catchpole and mechanical restraint, such as ropes, around the feet or legs. Once secured and the team is assured that the animal is sufficiently anesthetized to manipulate it, vital signs (body temperature, pulse and respiration rates) are taken. If available, an electrocardiogram (ECG) and pulse oximeter can monitor heart function and oxygen saturation of the blood (see Chapter 29, Anesthesia). Basic examinations should include auscultation of the heart and lungs, palpation of the abdomen and joints, and visualization of the general body and hair condition. It is a good idea to physically palpate the entire body, as the thick hair can easily conceal small masses, cuts, and abrasions. The use of an otoscope to examine the ears and an ophthalmoscope to examine the eyes is also recommended. Visualization of the genitalia is important, and palpation of testes or mammary glands is recommended to check for the presence of masses. Each time a bear is immobilized, a complete oral examination should be performed. Visualization of the teeth is important. The teeth should be checked for any fractures, evidence of caries, or periodontal disease. Any dental problems must be addressed at this time. Blood should be collected and a complete blood count (CBC) and serum profile should be performed (see Chapter 19, Clinical Pathology). Fecal samples or rectal cultures, as well as urine for urinalysis, are collected during the examination. Limbs may be radiographed with a portable x-ray machine, but any thoracic or abdominal radiographs will require the use of a large, nonmobile, high-mA machine, such as are used in human hospitals and advanced large-animal facilities (see Chapter 25, Radiography). Generally, these machines are fixed and in a radiology room; thus, radiology will generally require transport of the animal after immobilization and is a consideration that needs to be addressed prior to anesthesia. Portable ultrasound machines are very helpful in the diagnosis of certain conditions and are easily brought to the site of immobilization (see Chapter 26, Ultrasonography).
Venipuncture Blood collection in the polar bear is a relatively simple task on a healthy bear. There are always exceptions to the rule, such as a very sick animal with low blood pressure, an obese animal, or a very young bear without anesthesia. The primary sites for blood collection are the jugular veins and the femoral veins. The two jugular veins are located in the neck and travel from the angle of the jaw to the thoracic inlet down the ventrolateral aspect of the neck. The venipuncture site should be clipped and aseptically prepared by cleaning with an appropriate, fast-acting, broad-spectrum antiseptic, such as tincture of alcohol or alcohol mixtures. The vessels are distended by occluding them with pressure applied to the area over them, just anterior to the clavicle. Three methods can be used: a syringe and needle, a Vacutainer (Becton Dickinson, Franklin Lakes, NJ) setup, or an extension set with or without a Vacutainer setup. The femoral vein traverses the medial aspect of both upper hind limbs. It can be palpated in the tissue between the extensor and the flexor muscles of the hind limb. It is best accessed by having the animal in lateral recumbency, and having an assistant raise the opposite leg to allow for access to the limb being bled. The sublingual vein is useful for administering emergency medication. The saphenous vein can be used for either blood collection or the administration of intravenous fluids. The cephalic vein on the dorsal aspect of the forelimb is also accessible.
0839_frame_C45 Page 996 Tuesday, May 22, 2001 2:10 PM
996
CRC Handbook of Marine Mammal Medicine
Mechanical or Manual Restraint Polar bears can easily be managed and trained to enter a chute or squeeze system with the use of positive reinforcement (Ament, pers. comm.). Once in the area, the animal can be visualized and, if necessary, injected either by a jab-stick or with a blowgun. The material for the chute must be of a gauge and construction that prevents the bear from destroying or damaging it, and prevents the animal from hurting itself by biting it. It is very easy for a bear to fracture a canine tooth in the mesh if the gauge and dimensions are inappropriate. Because of the limitations of the chute or a squeeze cage, anesthesia is the most practical form of restraint for polar bears (Wallach and Boever, 1983; Fowler, 1986).
Anesthesia The use of anesthesia is the most efficient and safe way to examine an adult polar bear. If a painful procedure is to be performed, such as major or minor surgery, dental work, or obstetrics, the animal should be at a deep enough level of anesthesia to induce complete analgesia. There are many drug combinations and delivery systems available to immobilize a polar bear successfully. The choice should be that which is safe, effective, and adequate for the procedure. It should also be a combination that is comfortable for the person performing the anesthesia. Whatever the choice of anesthetic and mode of delivery, it should be reviewed with the medical team so all are aware of the expected effects and potential side effects of the drug combination and procedure. Choose the anesthetic based on the procedure length, amount of manipulation required, and the history of the individual animal. As with any medical procedure on a large carnivore, it must be well planned, and equipment and personnel must be assembled well ahead of time. Prior planning will aid in the efficiency of the procedure as well as allow for a greater comfort level of all involved. Worldwide, there are at least five manufacturers of projectile/dart systems for administration of pharmaceuticals in polar bears (see box below). Healthy polar bears have a thick layer of fat over most of their body, with particularly thick areas over the rump and the thighs. It is therefore most desirable to dart the bear in the neck, shoulder, or triceps muscle, where the fat is thinner. If darting in the rump, use a very long needle. A minimum of 7.5 cm (2.5 in.) is suggested. Suggestions have been made that one should use a
PROJECTILE SUPPLIERS Cap-hur Equipment 421 Tidwell Rd. Powder Springs, GA 30127
Tel-Inject USA, Inc. 9316 Soledad Canyon Road Saugus, CA 91350
Dan-Inject SA Private Bag X402 Skukuza 1350 South Africa
Zoolu Arms of Omaha 10315 Wright Street Omaha, NE 68124
Pneu-Dart, Inc. P.O. Box 1415 Williamsport, PA 17703
0839_frame_C45 Page 997 Tuesday, May 22, 2001 2:10 PM
Polar Bears
997
rifle-powered projectile dart, but in this author’s experience, even the very lightweight blow darts (Tel-Inject, Saugas, CA) are effective, if placed properly in the animal. If the anticipated effect is not achieved after darting, to ensure against overdosing, it is important to allow enough time to pass before an additional dose is delivered. If the dart is poorly placed in thick adipose tissue and the bear is overweight, the anesthetic will not be absorbed as quickly, and two things occur: (1) the animal does not become anesthetized quickly enough or adequately enough for the intended procedure to be conducted, and (2) recovery can be prolonged, since the drug may be slowly released from fat and only eventually metabolized. During induction, the bear should be kept in subdued light, if possible, and both visual and auditory input minimized. This decrease in arousal stimuli will lead to a quicker, smoother induction with less need for anesthetic supplementation during the procedure. During anesthesia, after applying a protective topical ophthalmic ointment, the animal should have its eyes covered, and noise should be kept to a minimum. The compulsion of people to pet and stroke the bear must be kept to a minimum, as this only stimulates the animal. Another consideration of consequence is how to move the bear, whether this is necessary for the procedure or if this is a primary goal of the immobilization. Movement of bears in a net and sling greatly increases the mean arterial blood pressure and may cause hypoxia. Simple disregard for body posture and compression of the thorax and airway may severely compromise the patency of available ventilation. This disregard for posture and possible compression has caused the death of several bears, so caution should be taken when moving polar bears by net and sling (Cattet et al., 1999). Monitoring the bear during anesthesia is essential. Equipment required will depend on the length and nature of the procedure (see Chapter 29, Anesthesia). At a minimum, body temperature, heart rate, and respiratory rate must be monitored. Other monitoring equipment often used includes a pulse oximeter and an ECG. If the animal is being switched to an inhalant anesthetic, the ability to monitor both inspired and expired oxygen and CO2, as well the anesthetic itself is desirable. If the procedure is to be prolonged, an intravenous catheter should be established and a slow, continuous fluid drip started. There are several areas that can support a catheter, but the cephalic vein provides easy access and is easy to maintain. Other physical parameters that must be monitored include the deep pain reflex, palpebral reflex, capillary refill time, jaw tone, and the panniculus reflex. There is a plethora of drug combinations used to immobilize polar bears, and they are summarized in Chapter 29, Anesthesia, and Chapter 31, Pharmaceuticals. One must use good judgment when selecting the agent and dosage to be used. This selection is dependent upon a number of factors including age, mode of delivery, side effects, procedure, need to reverse the anesthetic, analgesia, and the clinician’s familiarity with the drug. Another consideration for the immobilization of the animal is whether it is captive or free-ranging. Many of the studies with large numbers of individuals have been conducted on free-ranging animals. The variation between drug effects on free-ranging and captive animals is notable. There are differences in lean muscle mass, metabolic condition, diet, fat stores, and relative state of anxiety between a well-habituated, handinjected, captive bear compared with a free-ranging bear that may have been darted with a powder charge rifle from a helicopter. All of these differences influence drug effects.
Ketamine Ketamine (Ketamine, Fort Dodge Animal Health, Fort Dodge, IA) is a dissociative, nonbarbiturate anesthetic. Induction usually occurs within 10 min. It is relatively short acting (15 to 30 min), thus should only be used in short procedures. It causes muscle rigidity and may induce tonic– clonic convulsions, a typical effect of dissociative anesthetics, such as ketamine. It is painful upon injection because of the low pH of the solution. The animal’s eyelids will remain open
0839_frame_C45 Page 998 Tuesday, May 22, 2001 2:10 PM
998
CRC Handbook of Marine Mammal Medicine
throughout the procedure, so care must be taken to avoid damage to the eyes by applying ophthalmic ointment; the palpebral reflexes persist. The wide range of dosage of ketamine is one of the reasons it is difficult to use. Suggested doses range from 2 to 50 mg/kg administered via deep intramuscular (IM) injection, with smaller animals generally using a higher mg/kg dose than larger animals. It has been used alone in polar bears, but because of potentially large volumes, chances of convulsions, and short downtimes, it is used alone only on rare occasions (Sedgwick and Robinson, 1973). If an animal develops convulsions, 10 to 20 mg of diazepam should be given intravenously (if it safe to do so); otherwise, at the least, the drug should be administered deep IM. Ketamine may be used successfully as a supplement to any of the other anesthetics described below. Administration is best accomplished by using an established catheter and administering a bolus of 100 mg intravenously at regular intervals for an adult bear. In general, giving a bolus every 10 to 15 min will work well, but this schedule must be modulated, depending on biological variations in the animal, the animal’s response to the drug, and the procedure being performed.
Ketamine / Xylazine A ketamine/xylazine (Xylazine–100 Injectable, Butler Co., Columbus, OH), combination of drugs has replaced the previously used phencyclidine/promazine mixture and is used in a wide variety of carnivores. Xylazine is a non-narcotic that affects on the central nervous system by acting on the α-2-adrenergic receptors. It has been shown to enhance the effects of dissociatives such as ketamine and narcotics. With its use, one can effectively reverse the effects of the anesthesia. The advantages of this combination over that of ketamine alone are the reduced volume needed to immobilize the animal and the reversibility of the xylazine (α-2-agonist portion). The ketamine/xylazine combination is given in a 1:1 ratio using 6.7 mg/kg of each drug. Cubs require a dose of only 2.3 mg/kg. Induction can take 10 to 15 min, and arousal is possible in the early stages. It is best to wait long enough to be completely sure the animal will not react (xylazine typically requires approximately 20 min for full effect when administered IM). This combination has a wide safety margin and, with the highly concentrated products (200 mg/ml), a much smaller dart can be used. There is a fairly long recovery period with this drug combination when an antagonist is not used; however, there is at least one antagonist readily available (Lee et al., 1981; Schweinsburg et al., 1982). In other reports, a dosage of 4.5 to 7.0 ml/135 kg body weight of a combination of 200 mg ketamine:40 mg xylazine also produced excellent results. This mixture has been dubbed Capture-All 5 (Jessup and Clark, 1991). The antagonist yohimbine hydrochloride (Antagonil, Wildlife Laboratories, Inc., Fort Collins, CO) is used to reverse the effects of xylazine (Ramsay et al., 1985). Upon its administration, the animal usually has only a small amount of ketamine in its system and will abruptly wake. Yohimbine hydrochloride, given at 0.1 mg/kg intravenously, provides a safe and effective reversal of the anethesia. Polar bears recover in approximately 5 to 10 min, with some animals walking in this time frame (Ramsay et al., 1985). Atipamezole hydrochloride (Antiseden, Pfizer Animal Health, Exton, PA) is a more specific α-2-antagonist and is a better choice for xylazine or medetomidine (Domitor, Pfizer Animal Health) reversal. Its use is replacing yohimbine in many situations, because of this increased specificity (Phillips, pers. comm.).
Tiletamine HCl and Zolazepam HCl The combination of tiletamine and zolazepam in a 1:1 mixture is a commonly used anesthetic in the veterinary community (Telazol, Fort Dodge Animal Health, Fort Dodge, IA). This is a combination of a potent cyclohexamine (tiletamine) and diazpinone (zolazepam). Both
0839_frame_C45 Page 999 Wednesday, May 23, 2001 11:09 AM
Polar Bears
999
tiletamine and ketamine are in the same class of drug and therefore exhibit similar properties (Haigh et al., 1985). This combination of drugs is generally dosed at between 2 and 4 mg/kg (Jessup and Clark, 1991); yet, it is safe and effective as high as 5.0 mg/kg (Haigh et al., 1985). Sternal recumbency occurs within 5 min if the drug is administered IM. Maximum effect is achieved after about 20 min. Although variable, this drug combination generally offers between 1 and 2 hours of working time (Stirling et al., 1989).
Telazol /Medetomidine The combination of Telazol with the α-2-agonist medetomidine results in excellent anesthesia and seldom requires additional dosing even with procedures over 1 hour. The combination quickly immobilizes the bear with a small volume and allows for manipulation of the animal with relative safety. This α-2-agonist is more likely to result in markedly decreased respiration or apnea with peripheral vascular constriction; thus external pulse oximetry is unreliable. The bear may have some respiratory depression and hypoxemia at first, but this does not appear to be a major problem. The animals are quickly reversed with the use of atipamezole (Cattet et al., 1997b; 1999).
Etorphine Etorphine (M99, Kruger-Med Pharmaceuticals, Johannesburg, South Africa) is a potent Schedule II narcotic. It is a morphine derivative and has been successfully used in polar bears at a dosage of 0.020 to 0.050 mg/kg (Schweinsburg et al., 1982). Induction is rapid, and one sees a decrease in respiratory rate and, at times, apnea. If apnea occurs, the procedure should be aborted and the anesthesia reversed with the antagonist, diprenorphine (M50-50, Kruger-Med Pharmaceuticals). Diprenorphine is administered intravenously at a dosage of twice that of the etorphine. The reversal occurs within 5 min. It may also be administered IM with reversal occurring between 20 and 45 min. Renarcatization has been reported, but is not a common occurrence. If it should occur, the reversal agent needs to be administered again.
Carfentanil Carfentanil (Wildnil, Wildlife Pharmaceuticals, Inc., Fort Collins, CO), a derivative of fentanyl, is considered up to 20,000 times more potent than morphine. Its actions are similar to etorphine, and it is a respiratory depressant. This drug is generally given IM using remotedarting equipment at 0.02 mg/kg (Haigh et al., 1983), but it also may be used orally at a dose of 5.7 to 9.0 mg/kg (Ramsay et al., 1994). The induction time is very short, and there is a propensity for renarcatization. The preferred antagonist is naltrexone. It should be dosed at 100 mg/mg of carfentanil used. The dose should be split with 50% given IM and 50% intravenously. This dose will help decrease the incidence of renarcitization (Haigh et al., 1983).
Fentanyl Citrate Fentanyl citrate (Innovar, RX Veterinary Products, Porterville, CA) is not used on a regular basis, but has been used historically to immobilize polar bears. It is a narcotic and dosages of 0.34 to 0.68 mg/kg work well, with a quick induction and no respiratory depression (Paternaude, 1979). Naloxone is the reversal drug of choice.
Inhalation Agents There are a variety of ways to use inhalant anesthetics, but it is recommended to use one of the above-listed drugs to gain access to the animal, followed by intubation, and placing the
0839_frame_C45 Page 1000 Tuesday, May 22, 2001 2:10 PM
1000
CRC Handbook of Marine Mammal Medicine
animal on a rebreathing gas anesthesia machine. In general, any of the currently used veterinary anesthetics work well, with isoflurane the most commonly used. When intubating the bear, it is important to use a strong bite block for safety of both the person intubating the animal and the endotracheal tube. The use of a recirculating system with a cuffed endotracheal tube allows for excellent control of the level of anesthesia and for any aided respirations if required, and it decreases the amount of escaping anesthetic.
Systemic Diseases The polar bear is likely susceptible to most diseases contracted by other carnivores and specifically other Ursidae. There has been speculation that bears are resistant to many of the viruses that infect other carnivores (Fowler, 1986). Despite this speculation, polar bears do become infected with viruses, bacteria, parasites, protozoa, and fungi; they also experience developmental problems, nutritional diseases, neoplasia, and traumatic lesions. Diagnosis and treatment of the various problems may involve a complete physical examination, as described above, or a rule-out list related to physical signs followed by appropriate treatment.
Developmental /Anomalous Diseases There are cases of female pseudohermaphroditic polar bears at Svalbard. These may be caused by excessive androgen secretion from a tumor in the particular female, or as a result of endocrine disruption due to environmental pollutants (Wiig et al., 1998). Hypospadias was reported in a 1.5-year-old male, but was surgically corrected without incident (Stamper et al., 1999). The first case of agenesis of the radius in a nondomestic species was reported in a polar bear cub delivered by Cesarean section. The cause is unknown (Lanthier et al., 1998). The occurrence of supernumerary mammae has also been reported (Derocher, 1990). Gastric dilatation and volvulus has been reported as the cause of death in free-ranging polar bears and in several other species of captive bears (Amstrup and Nielson, 1989). Acute renal failure was documented by Crawshaw (1980).
Nutritional Diseases Multiple disease problems related to nutrition have been reported in polar bears. Some of these nutritional problems may arise from the unique diet of polar bears in the wild and deviations from those diets in captive situations. Rickets has been reported in two hand-reared cubs, but the condition resolved after dietary changes (Kenny et al., 1999). It has also been shown that hypovitaminosis A results in dermatitis in polar bears. This dermatitis appears to be treatable with vitamin A supplementation, either as an increase in the overall content of the ration or by supplementation with a vitamin A–rich additive, such as cod liver oil (Kock et al., 1985). Surveys have revealed that zoos that feed bears diets with over 20,000 IU of vitamin A/kg of feed have bears with good to excellent hair coats (Foster, 1981). Although wild polar bears have high levels of vitamin A in their livers (Leighton et al., 1988), any supplementation must be considered carefully to avoid toxemia. Calcium deficiencies have also been reported in polar bears fed on a meat-only diet, but this is not commonly seen in a modern zoo setting (Wallach, 1970).
Neoplasia There have been a limited number of reports of neoplasia in polar bears; this may be partially because the neoplasia is found as ancillary components at necropsy and not the cause of death. Biliary adenocarcinomas have been described in the polar bear (Miller et al., 1985), as well as
0839_frame_C45 Page 1001 Tuesday, May 22, 2001 2:10 PM
Polar Bears
1001
hepatocellular carcinomas (Miller and Boever, 1984). Signs of these diseases are associated with dysfunctional livers and include ataxia, icterus, acites, and lethargy. These are most commonly seen in bears older than 17 years.
Infectious Diseases Viral Disease
Viral disease in the polar bear has been limited to two major viruses: rabies and morbillivirus. However, one should not be complacent about investigating potential viral disease. A complete diagnostic workup will help determine the potential role of underlying viral disease in other maladies and will help identify potential viral etiologies for ill bears. Rabies has been reported in a polar bear, but it is not likely a concern for the overall wild population, nor is it a large concern in the captive community (Taylor et al., 1991). Signs are highly variable. In the one described case of rabies, the animal was found with posterior paralysis. Diagnosis was made using mouse inoculation and immunoperoxidase stain on the spinal cord and Gasserian ganglion. In areas that have a high prevalence of rabies in wild and feral animals, it may be prudent to vaccinate bears with a killed product. Morbillivirus titers have been found in many free-ranging polar bears. Because of the dieoffs of other marine mammals from morbillivirus epidemics (see Chapter 15, Viral Diseases, and Chapter 2, Emerging Diseases), there was much concern about the potential for spread to polar bears. Despite the high prevalence, however, it has been shown that the polar bear virus is most similar to the canine distemper virus and not that of the other marine species. It is likely to be indigenous in the population and not a real health threat (Garner, 1996; Garner et al., 2000). Although canine adenovirus type 1 has been shown to cause central nervous system disease in the black bear (Ursus americanis), there is little evidence to show it is a disease problem in polar bears (Collins et al., 1984). It is only mentioned here to indicate that vaccination of polar bears against this virus is not warranted. Bacterial Disease
Leptospirosis is a disease in polar bears characterized by nonspecific signs of weakness, diarrhea, icterus, and possibly muscle fasciculations (Nall, 1975). Diagnosis is established on serological evidence and clinical signs. It is also possible to culture the organism from urine. In one case, a young bear was successfully treated with chloramphenicol. Since the organism is carried by rodents, it is important to keep rodent numbers to a minimum in and around bear enclosures. In most institutions, it is impossible to keep rodents out of bear enclosures, but it is important to attempt to do so. To help assure the health of polar bears, a vaccination program should be implemented to help decrease the chance of contracting the disease in areas where there is a high prevalence of both the disease and the vector. One case of pleuritis and peritonitis caused by Pasteurella multocida has been reported (Keahey, 1968). Vaccination is not advised because of the rarity of occurrence. It is prudent to be vigilant for any of the other myriad potential bacterial infections that could cause disease in polar bears. Omphalophlebitis and necrotic enteritides with mixed bacteria were observed in polar bear cubs that died in zoological parks (Griner, 1983). Mycotic Disease
Blastomycosis in the polar bear was described as a diffuse pulmonary disease with a pleural effusion. Diagnosis was established with cytology and positive cultures. The animal was successfully treated with itraconazole, 4.5 mg/kg/day divided into two oral doses for 90 days (Morris, pers. comm.). In general, the signs of blastomycosis are increasing lethargy and
0839_frame_C45 Page 1002 Tuesday, May 22, 2001 2:10 PM
1002
CRC Handbook of Marine Mammal Medicine
anorexia with weight loss. Abnormalities in bloodwork are inconsistent, and quality thoracic radiographs are difficult to obtain. Diagnosis is made by culture, cytology, and the use of an AGID serological test (Clyde et al., 1996). There has been a report of candidiasis in the oral cavity and stomach of a cub being treated with erythromycin. The causative agent was Candida albicans, which can be treated with a number of agents, including nystatin, amphotericin B, flucytosine, or one of the imidazoles (Finn, 1969; Kirk, 1986) (see Chapter 17, Mycotic Diseases). Parasitic Disease
Parasitic disease in free-ranging marine mammals is considered to be one of the major causes of disease and death, although few species of parasites have been documented in polar bears (see Chapter 18, Parasitic Diseases). In captive situations, a preventative program consisting of routine fecal analysis and diagnosis of parasitism, combined with a regular deworming program, is essential for the reduction and possible elimination of internal and external parasites. No single system will be effective for all situations, and the program will need to be developed specifically for the individual institution. The drug dosages for anthelmintics in polar bears can be found in Chapter 31 (Pharmaceuticals, Table 5). The two ascarids of polar bears are Baylisascaris transfuga and B. multipapillata (Wallach and Boever, 1983; Fowler, 1986). Once established, it is very difficult to rid an enclosure completely of these organisms, because the ova may be viable in the environment for as long as 2 years. Clinical signs in polar bears infected with Baylisascaris spp. are variable, depending on the intensity of infection, but can include loose stool to diarrhea, rough hair coat, and, in extreme cases, severe weight loss and intestinal obstruction, leading to death. Diagnosis may be established by fecal float or by direct observation. Trichinella spiralis is common in polar bears (McColl, 1982; Wallach and Boever, 1983; Fowler, 1986; Pozio et al., 1990; Sleeman et al., 1994; YepezMulia et al., 1996). It is generally considered an incidental finding and does not cause overt disease in the animals. When signs do occur, they generally are muscular pain and eosinophilia. At times there can be central nervous system involvement. Cestodes have been reported but rarely cause disease (see Chapter 18, Parasitic Diseases). Although external parasites are ubiquitous, they seldom pose a real problem for polar bears, other than dermatitis, which is described below. Polar bears have been reported with infestations of mites and ticks (see below).
Skin Disease Captive polar bears have a variety of skin diseases. Most of these are described as dermatitis, with or without alopecia. There are a number of etiologies and a few are examined below. Polar bears are affected by a variety of species of mites, including Audycoptes, Sarcoptes, and Demodex (Fowler et al., 1979). Dermatitis is generally more severe in winter and is very pruritic. Diagnosis can be accomplished by a deep skin scraping at the edge of the lesion, and subsequent microscopic evaluation of the scraping. There are multiple therapies available, and many of the products used for horses, dogs, and cats are effective on bears. Treatment generally includes immobilizing the bear and treating it topically with one of the commercially available products. Dermatophilus congolensis has been diagnosed as a cause of severe, pruritic dermatitis (Smith and Cordes, 1972; Newman et al., 1975). It can be a severe disease and generally starts on the dorsum and spreads laterally. The coat becomes yellowed and oily in appearance. This skin disease is diagnosed by clinical signs, microscopic examination of the lesions, and by biopsy.
0839_frame_C45 Page 1003 Tuesday, May 22, 2001 2:10 PM
Polar Bears
1003
Treatment is generally a combination therapy of cleaning the animal with a disinfectant soap and placing it on penicillin or doxycycline. Other skin problems are as follows. Hypovitaminosis A has been shown to cause alopecia and general poor coat condition; this was discussed in the nutrition section and will not be addressed again here. Inhalant allergic dermatitis (atopy) has been reported in the polar bear (Harper et al., 1988). It was successfully treated with steroids and by hyposensitization. The common occurrence of green bears in captivity is due to an algal growth within the hollow hair shafts of the guard hairs (Lewin and Robinson, 1979). An ulcerative dermatitis linked to thyroid disease resolved after treatment with l-thyroxine (Hoff, 1980).
Dental Disease Although diagnosis and treatment of dental disease is fairly straightforward, it is important to know that most of the endodontic procedures performed on small carnivores and primates can be done on these massive creatures (Shagram, 1983). Apicoectomy and endodontic repair have been reported (Jensen et al., 1986), as well as root canals and extraction of canines (Forier et al., 1975; Dufor et al., 1995).
Trauma As one may expect from large animals who seldom cross paths with one another, polar bear meetings and interactions can be traumatic. Just as they can harm one another, they also, at times, will injure themselves on bars of enclosures or transport crates. Many of the polar bear’s dental problems are traumatically induced by inanimate objects. There are several reports of polar bears sustaining severe fractures from cage mates or even their mothers (Van Foreest et al., 1987; Kohm, 1991; Cook et al., 1994). These incidences can be treated as other traumatic cases, but the size and strength of the animal must be considered when choosing treatment.
Toxins There are relatively few reports of toxicities in polar bears, but they do exist. The polar bear is likely susceptible to any toxin that would affect other carnivores. There have been reports of excess heavy metals in polar bears (Lentfer and Galster, 1987) as well as ethylene-glycol (antifreeze) poisoning in free-ranging bears (Amstrup et al., 1989). There are dramatic metabolic changes in polar bears exposed to crude oil or petroleum distillates on their fur (as would occur in maritime spills). They cannot thermoregulate properly, and thus their metabolic rates significantly increase (Hurst et al., 1991).
Zoonoses There are few disease entities that can be transmitted from the polar bear to humans. Of the reports available, the probability of a human contracting trichinosis is the most likely (Nozais et al., 1996). Leptospirosis is also communicable, but exposure is usually limited with even rudimentary sanitary precautions by keepers. Baylisascaris larval migrans is also of potential concern with infections of Baylisascaris transfuga and B. multipapillata. Rabies is also zoonotic, but the likelihood of surviving an attack by a rabid polar bear seems slim.
0839_frame_C45 Page 1004 Tuesday, May 22, 2001 2:10 PM
1004
CRC Handbook of Marine Mammal Medicine
Acknowledgments The author thanks Lyndsay Phillips, Cheryl Messinger, and Robert Wilson for their editorial contributions to this chapter.
References Ames, A., 1993, The Behaviour of Captive Polar Bears, University Federation of Animal Welfare, South Mimms, Hertfordshire, U.K., 67 pp. Amstrup, S.C., and Nielson, C.A., 1989, Acute gastric dilation and volvulus in a free-living polar bear, J. Wildl. Dis., 25: 601–604. Amstrup, S.C., Gardner, C., Myers, K.C., and Oehme, F.W., 1989, Ethylene-glycol (antifreeze) poisoning in a free-ranging polar bear, Vet. Hum. Toxicol., 31: 317–319. Ball, M.D., Furr, J.S., and Olson, J.A., 1986, Acyl coenzyme A: Retinol acyltransferase activity and the vitamin A content of polar bear (Ursus maritimus) liver, Comp. Biochem. Physiol. B, 84: 513–517. Best, R.C., 1982, Thermoregulation in resting and active polar bears, J. Comp. Physiol., 146: 63–73. Boyd, A.L., Lockyer, C., and Marsh, H.D., 1999, Reproduction in marine mammals, in Biology of Marine Mammals, Reynolds, J.E., and Rommel, S.A. (Eds.), Smithsonian Institution Press, Washington, D.C., 218–286. Bruce, D.S., Darling, K.K., Seeland, K.J., Oeltgen, P.R., Nilekani, S.P., and Amstrup, S.C., 1990, Is the polar bear (Ursus maritimus) a hibernator? Continued studies on opioids and hibernation, Pharmacol. Biochem. Behav., 35: 705–711. Cattet, M.R.L., Atkinson, S.N., Polischuk, S.C., and Ramsay, M.A., 1997a, Predicting body mass in polar bears: Is morphometry really useful? J. Wildl. Manage., 61: 1083–1090. Cattet, M.R.L., Caulkett, N.A., Polischuk, S.C., and Ramsay, M.A., 1997b, Reversible immobilization of free-ranging polar bears with medetomidine-zolazepam-tiletamine and atipamezole, J. Wildl. Dis., 33: 611–617. Cattet, M.R.L., Caulkett, N.A., Streib, K.A., Torske, K.E., and Ramsay, M.A., 1999, Cardiopulmonary response of anesthetized polar bears to suspension by net and sling, J. Wildl. Dis., 35: 548–556. Clyde, V.L., Ramsay, E.C., and Munson, L., 1996, A review of blastomycosis in large zoo carnivores, in Proceedings of the American Association of Zoo Veterinarians, 554. Collins, J.E., Leslie, P., Johnson, D., Helson, D., Peden, W., Boswell, R., and Draayer, H., 1984, Epizootic of adenovirus infection in American black bears, J. Am. Vet. Med. Assoc., 185: 1430. Cook, R., Thacher, C., Calle, P., Raphael, B., Kapatkin, A., and Stetter, M., 1994, Multiple hindlimb fracture repair in an adolescent polar bear (Ursus maritimus), in Proceedings of the International Association for Aquatic Animal Medicine, 25: 154–155. Crawshaw, G.J., 1980, Acute renal failure in a polar bear, in Proceedings of the American Association of Zoo Veterinarians, 135. Derocher, A.E., 1990, Supernumerary mammae and nipples in the polar bear (Ursus maritimus), J. Mammal., 71: 236–237. Derocher, A.E., and Wiig, O., 1999, Observation of adoption in polar bears, Arctic, 52: 413–415. Derocher, A.E., Stirling, I., and Andriashek, D., 1992, Pregnancy rates and serum progesterone levels of polar bears in western Hudson Bay, Can. J. Zool., 70: 561–566. Derocher, A.E., Andriashek, D., and Arnauld, J.P.Y., 1993, Aspects of milk composition and lactaction in polar bears, Can. J. Zool., 71: 561–567. Dufor, P., Ballingand, M., Pypendop, B., Loneux, P., and Bonnevie D., 1995, Extraction of a mandibular canine of a polar bear, Ann. Med. Vet., 139: 205–207. Finn, P.J., 1969, Pyocephalus and gastritis in a polar bear, J. Am. Vet. Med. Assoc., 155: 1086. Forier, R.C., Miller, T., and Suigert, J., 1975, Root canal therapy on two polar bears, in Proceedings of the American Association of Zoo Veterinarians.
0839_frame_C45 Page 1005 Tuesday, May 22, 2001 2:10 PM
Polar Bears
1005
Foster, J.W., 1981, Dermatitis in polar bears-a nutritional approach to therapy, in Proceedings of the American Association of Zoo Veterinarians, 58–60. Fowler, M.E., 1978, Carnivores, in Restraint and Handling of Wild and Domestic Animals, Iowa State University Press, Ames, 19. Fowler, M.E., 1986, Ursidae, in Zoo and Wild Animal Medicine, W.B. Saunders, Philadelphia, 811 pp. Fowler, M.E., Lavoiperre, M., and Schultz, T., 1979, Audycoptic mange in bears, in Proceedings of the American Association of Zoo Veterinarians, 104. Furr, N.J., and Stenhaus, G.B., 1985, An observation of possible cannibalism by polar bears, Can. J. Zool., 63: 1516–1517. Garner, G., 1996, Serological evidence of morbillivirus infection in polar bears (Ursus maritimus) from Alaska and Russia, Vet. Rec., 138: 615–618. Garner, G., Evermann, J.F., Saliki, F.T., Follmann, E.H., and McKeirnan, A.J., 2000, Morbillivirus ecology in polar bears (Ursus maritimus), Polar Biol., 23: 474–478. Griner, L.A., 1983, Pathology of Zoo Animals, Zoological Society of San Diego, San Diego, CA, 423 pp. Haigh, J.C., Lee, L.J., and Schweinsburg, R.E., 1983, Immobilization of polar bears with carfentanil, J. Wildl. Dis., 19: 140. Haigh, J.C., Stirling, I., and Broughton, E., 1985, Immobilization of polar bears with a mixture of tiletamine hydrochloride and zolazepam hydrochloride, J. Wildl. Dis., 21: 43–47. Harper, J., White S., Stewart, L., and Pelto, J., 1988, Inhalant allergic dermatitis in a polar bear, in Proceedings of the American Association of Zoo Veterinarians, 97–98. Hoff, S., 1980, Skin disease in two polar bears at the Stanley Park Zoo, in Proceedings of the American Association of Zoo Veterinarians, 107. Hurst, R.J., Watts, P.D., and Oritsland, N.A., 1991, Metabolic compensation in oil-exposed polar bears, J. Thermal Biol., 16: 53–56. Jensen, J., Dorn, A., and Morris, P.J., 1986, Apicoectomy and endodontic repair in a polar bear, in Proceedings of the American Association of Zoo Veterinarians. Jessup, D.A., and Clark, W.E., 1980, Wildlife Restraint Manual, California Department of Fish and Game, Stockton, CA, 192 pp. Keahey, K.K., 1968, Incidence and classification of exotic animal disease, in Proceedings of the American Association of Zoo Veterinarians. Kenny, D.E., Irlbech, N.A., and Eller, J.L., 1999, Rickets in two hand reared polar bear (Ursus maritimus) cubs, J. Zoo Wild Anim. Med., 30: 132–140. Kirk, R.W. (Ed.), 1986, Current Veterinary Therapy IX, Small Animal Practice, W.B. Saunders, Philadelphia, 1346 pp. Kock, R.A., Thomsett, L.R., and Henderson, G.M., 1985, Alopecia in polar bears (Thalarctos maritimus): A report of two cases, in Proceedings of International Symposium on Wild Animals, Rostok, East Germany, 27: 63. Kohm, A., 1991, Problematic hand-rearing of a polar bear as a result of a fractured pelvic symphysis and an infection of the upper throat region, Berl. Münch. Tierärztl. Wochenschr., 104: 10–12. Lanthier, C., Dupuis, J., and Pare, J., 1998, Agenesis of a radius in a polar bear cub (Ursus maritimus), J. Zoo Wild Anim. Med., 29: 65–67. Latinen, K., 1987, Longevity and fertility of the polar bear (Ursus maritimus phipps) in captivity, Zool. Gart., 57: 197. Leatherland, J.F., and Ronald, K., 1981, Plasma concentrations of thyroid hormones in a captive and feral polar bear, Comp. Biochem. Physiol. A, 70: 575. Lee, J., Schweinsburg, R., Kernan, F., and Haigh, J., 1981, Immobilization of polar bears (Ursus maritimus phipps) with ketamine hydrochloride and xylazine hydrochloride, J.Wildl. Dis., 17: 331. Leighton, F.A., Cattet, M., Norstrom, R., and Trudeau, S., 1988, A cellular basis for high levels of vitamin A in livers of polar bears (Ursus maritimus): The Ito cell, Can. J. Zool., 66: 480–487. Lentfer, J.W., and Galster, W.A., 1987, Mercury in polar bears from Alaska, J. Wildl. Dis., 23: 338–341. Lewin, R.A., and Robinson, P.T., 1979, The greening of polar bears in zoos, Nature, 278: 445–447.
0839_frame_C45 Page 1006 Tuesday, May 22, 2001 2:10 PM
1006
CRC Handbook of Marine Mammal Medicine
Lewis, R.W., and Lentfer, J.W., 1967, Vitamin A content of polar bear liver: Range and variability, Comp. Biochem. Physiol. B, 22: 923–926. Lono, O., 1972, Polar bears fetuses found in Svalbard, Nor. Polarinst. Arbok, 149: 294. Lowry, L.F., Burns, J.J., and Nelson, R.R., 1987, Polar bear, Ursus maritimus, predation on belugas, Delphinapterus leucas, in the Bering and Chukchi Seas, Can. Field Nat., 101: 141–146. Matthews, D., 1993, Polar Bear, Chronicles Books, San Francisco, CA, 110 pp. McColl, K.A., 1982, Trichinosis in a polar bear, Thalarctos maritimus, from the Royal Melbourne Zoo, Aust. Vet. J., 59: 61–64. Miller, R.E., and Boever, W.J., 1984, Case reports—A hepatocellular carcinoma and biliary adenocarcinoma in two polar bears (Thalarctos maritimus), in Proceedings of the American Association of Zoo Veterinarians, 112–114. Miller, R.E., Boever, W.J., Thornburg, L.P., and Curtis-Velasco, M., 1985, Hepatic neoplasia in two polar bears, J. Am. Vet. Med. Assoc., 187: 1256–1258. Nall, J.D., 1975, Leptospirosis outbreak in the Birmingham, Alabama Zoo, in Proceedings of the American Association of Zoo Veterinarians, 162. Nelson, R.A., 1983, Feeding strategies and metabolic adjustments of the polar bear, in Proceedings, 3rd Annual Dr. Scholl Conference on the Nutrition of Captive Wild Animals, 3: 93–96. Newman, M.S., Cook, R.W., Appelhof, W.K., and Kitchen, H., 1975, Dermatophilus congolensis in polar bears, J. Am. Vet. Med. Assoc., 167: 561. Nozais, J.P., Mannevy, B., and Danis, M., 1996, Two cases of trichinosis from polar bear (Thalarctos maritimus) meat, Med. Mal. Infect., 26: 732–733. Oritsland, N.A., 1970, Temperature regulation of the polar bear (Thalarctos maritimus), J. Comp. Biochem. Physiol. A, 37: 225–233. Ovsyanikov, N., 1996, Polar Bears, Voyageur Press, Stillwater, MN, 144 pp. Palmer, S.S., Nelson, M.A., Ramsay, M.A., Stirling, I., and Bahr, J.M., 1988, Annual changes in serum sex steroids in male and female black (Ursus americanus) and polar (Ursus maritimus) bears, Biol. Reprod., 38: 1044–1050. Paternaude, R.P., 1979, Evaluation of fentanyl citrate, etorphine hydrochloride and naloxone hydrochloride in captive polar bears, J. Am. Vet. Med. Assoc., 175: 1006. Poulsen, E.M.B., Honeyman, B., Valentine, P.A., and Teskey, G.C., 1996, Use of fluoxetine for the treatment of stereotypical pacing behavior in a captive polar bear, J. Am. Vet. Med. Assoc., 209: 1470. Pozio, K.V., Mortelmans, D.J., and Demeurichy, W., 1990, Characterization of trichinella isolate from polar bear, Ann. Soc. Belge Med. Trop., 70: 131–135. Ramsay, E.C., Sleeman, J.M., Clyde, V.L., and Gieser, D., 1994, Immobilization of bears using orally administered carfentanil citrate, in Proceedings of the American Association of Zoo Veterinarians, 214. Ramsay, M.A., and Stirling, I., 1988, Reproductive biology and ecology of female polar bears, J. Zool., 214: 601–634. Ramsay, M.A., Stirling, I., Knutsen, L.O., and Boughton, D., 1985, Use of yohimbine hydrochloride to reverse immobilization of polar bears by ketamine hydrochloride and xylazine hydrochloride, J. Wildl. Dis., 21: 396–400. Ramsay, M.A., Nelson, R.A., and Stirling, I., 1991, Seasonal changes in the ratio of serum urea to creatinine in feeding and fasting polar bears, Can. J. Zool., 69: 298–302. Reynolds, J.E., and Rommel, S.A. (Eds.), 1999, Biology of Marine Mammals, Smithsonian Institution Press, Washington, D.C., 578 pp. Schweinsburg, R.E., Lee, L.J., and Haigh, J.C., 1982, Capturing and handling polar bears in the Canadian Arctic, in Chemical Immobilization of North American Wildlife, Nielsen, L., Haigh, J.C., and Fowler, M.E. (Eds.), Wisconsin Humane Society, Milwaukee, 267–289. Seal, U.S., Swain, W.R., and Erickson, A.W., 1967, Hematology of the Ursidae, Comp. Biochem. Physiol., B, 22: 451. Sedgwick, C.J., and Robinson, P.T., 1973, Immobilizaton of a polar bear (Thelarctos maritimus) with ketamine HCl, J. Zoo Wild Anim. Med., 4: 27.
0839_frame_C45 Page 1007 Tuesday, May 22, 2001 2:10 PM
Polar Bears
1007
Shagram, E.V., 1983, Step by step endodontic techniques in exotic animal dentistry, in Proceedings of the American Association of Zoo Veterinarians. Sleeman, J.M., Ramsay, E.C., Faulkner, C.T., Patton, S., and Mason, G., 1994, Trichinosis in a polar bear (Ursus maritimus), in Proceedings of the American Association of Zoo Veterinarians, 352–353. Smith, C.F., and Cordes, D.O., 1972, Dermatitis caused by Dermatophilus congolensis in polar bears, Br. Vet. J., 128: 366. Stamper, M.A., Norton, T., Spodnick, G., Marti, J., and Loomis, M., 1999, Hypospadias in a polar bear (Ursus maritimus), J. Zoo Wild Anim. Med., 30: 141–144. Stempniewicz, L., 1993, The polar bear Ursus maritimus feeding in a seabird colony in Frans Josef Land, Polar Bear Res., 12: 33–36. Stirling, I., 1974, Midsummer observations of the behavior of wild polar bears, Can. J. Zool., 52: 1191–1198. Stirling, I., 1988, Polar Bears, University of Michigan Press, Ann Arbor, 232 pp. Stirling, I., Spencer, C., and Andriashek, D., 1989, Immobilization of polar bears; Ursus maritimus with telazol in the Canadian Arctic, J. Wildl. Dis., 25: 159–168. Taylor, M., Elkin, B., Maier, N., and Bradley, M., 1991, Observation of a polar bear with rabies, J. Wildl. Dis., 27: 337–339. Teskey, G.C., Valentine, P.A., Paulsen, M.B., Honeyman, V., and Cooper, R.M., 1996, Treatment of stereotypic behavior in the polar bear (Ursus maritimus), in Proceedings of the American Association of Zoo Veterinarians, 334. USDA (U.S. Department of Agriculture), 1984, Animal and Plant Health Inspection Service rules and regulations, marine mammals: Humane handling, care, treatment and transportation, in Fed. Regis. 49, 26672. Van Foreest, A.W., Barneveld, A., Dik, K.J., Lagerwey, E., Merkens, H.W., and Nemeth, F., 1987, Arthrodesis of a luxated stifle joint in a polar bear, in Proceedings of the American Association of Zoo Veterinarians, 399. Wallach, J.D., 1970, Nutritional Disease of Exotic Animals, J. Am. Vet. Med. Assoc., 157: 583–599. Wallach, J.D., and Boever, W.J., 1983, Ursidae, in Disease of Exotic Animals: Medical and Surgical Management, W.B. Saunders, Philadelphia, 1159 pp. Wemmer, C., 1974, Design for polar bear maternity dens, Int. Zoo Yearb., 14: 222. Wemmer, C., Von Ebers, M., and Scow, K., 1976, An analysis of chuffing vocalizations in the polar bear, J. Zool., 180: 425–439. Wiig, O., Derocher, A.E., Cronin, M.M., and Skaare, J.U., 1998, Female pseudohermaphrodite polar bears at Svalbard, J. Wildl. Dis., 34: 792–796. Yepez Mulia, L., Arriaga, D., Pena, M.A., Gual, F., and OrtegaPierres, G., 1996, Serologic survey of trichinellosis in wild mammals kept in a Mexico City zoo, Vet. Parasitol., 67: 237–246.
0839_frame_C45 Page 1008 Tuesday, May 22, 2001 2:10 PM
Appendix Page 1009 Wednesday, May 23, 2001 11:12 AM
Appendices
Appendix Page 1010 Wednesday, May 23, 2001 11:12 AM
Appendix Page 1011 Wednesday, May 23, 2001 11:12 AM
Appendix A
Conversions
TABLE 1 Blood Values To Convert
Item Albumin Bilirubin, total BUN Calcium Cholesterol Cortisol Creatinine Globulin Glucose Insulin Iron Lactate Lead Magnesium Phosphorus Progesterone Protein Thyroxine Uric acid
To Convert
From
To
Multiply by
From
To
Multiply by
g/dl mg/dl mg/dl mg/dl mg/dl µg/dl mg/dl g/dl mg/dl µU/ml µg/dl mg/dl µg/dl mg/dl mg/dl ng/dl g/dl µg/dl mg/dl
g/l µmol/l mmol/l mmol/l mmol/l nmol/l µmol/l g/l mmol/l pmol/l µmol/l mmol/l µmol/l mmol/l mmol/l nmol/l g/l nmol/l mmol/l
10.0 17.1 0.714 0.25 0.02586 27.59 88.4 10.0 0.05551 7.175 0.1791 0.111 0.04826 0.4114 0.3229 0.032 10.0 12.87 0.059
g/l µmol/l mmol/l mmol/l mmol/l nmol/l µmol/l g/l mmol/l pmol/l µmol/l mmol/l µmol/l mmol/l mmol/l nmol/l g/l nmol/l mmol/l
g/dl mg/dl mg/dl mg/dl mg/dl µg/dl mg/dl g/dl mg/dl µU/ml µg/dl mg/dl µg/dl mg/dl mg/dl ng/dl g/dl µg/dl mg/dl
0.1 0.059 1.4 4.0 38.7 0.0362 0.0113 0.1 18.0 0.1296 5.58 9.009 20.72 2.43 3.097 31.25 0.1 0.0777 16.95
Item Blood volume Plasma volume 500 ml blood PCV 1 mEq/l
Equal to approx. 8% body weight approx. 4–5% body weight approx. 1 lb Hematocrit (HCT) 1 mmol/l
1011
Appendix Page 1012 Wednesday, May 23, 2001 11:12 AM
1012
Handbook of Marine Mammal Medicine
TABLE 2 Weights and Measures To Convert
Category Weight
Linear
Liquid
Energy
To Convert
From
To
Multiply by
pound (lb) ounce (oz) gram (g) g g mg µg ng mile yard (yd) foot (ft) inch (in.) mm gallon (gal) quart (qt) qt fl.oz. fl.oz. fl.oz. cup (c) fl.oz. milliliter (ml) microliter (µl) nanoliter (nl) picoliter (pl) femtoliter (fl) tablespoon (tbsp) teaspoon (tsp) ml drop calorie (cal) calorie (cal) kcal kcal
kilogram (kg) lb oz grain (gr) mg µg ng picogram (pg) km m cm cm cm liter (l) gal liter (l) qt l cup (c) pint (pt) ml l ml µl nl pl ml
0.454 0.0625 0.0352 15.43 1000 1000 1000 1000 1.6 0.917 30.48 2.54 0.1 3.785 0.25 0.946 0.0313 0.0296 0.125 0.5 30.0 0.001 0.001 0.001 0.001 0.001 15.0
ml cubic cm (cc) ml joules (J) kcal kJ MJ
From
To
Multiply by
kg lb oz gr mg µg ng pg km m cm cm cm l gal l qt l c pt ml l ml µl nl pl ml
lb oz g g g mg µg ng mile yd ft in mm gal qt qt fl.oz. fl.oz. fl.oz. c fl.oz. ml µl nl pl fl tbsp
2.2 16.0 28.4 0.0648 0.001 0.001 0.001 0.001 0.625 1.09 0.0328 0.394 10.0 0.264 4.0 1.057 32.0 33.8 8.0 2.0 0.033 1000 1000 1000 1000 1000 0.067
5.0
ml
tsp
1.0 0.05 4.18 0.001 4.18 0.00418
cc ml J kcal kJ MI
ml drop cal cal kcal kcal
0.2 1.0 20 0.239 1000 0.239 239.2
Appendix Page 1013 Wednesday, May 23, 2001 11:12 AM
1013
Conversions
TABLE 3 Fahrenheit vs. Centigrade Conversion Chart °F
°C
95.0 95.2 95.4 95.6 95.8 96.0 96.2 96.4 96.6 96.8 97.0 97.2 97.4 97.6 97.8 98.0 98.2 98.4 98.6
35.0 35.1 35.2 35.3 35.4 35.6 35.7 35.8 35.9 36.0 36.1 36.2 36.3 36.4 36.6 36.7 36.8 36.9 37.0
Temperature formulas: °C = 5/9 (°F – 32) °F = (ºC × 9/5) + 32
°F 98.8 99.0 99.2 99.4 99.6 99.8 100.0 100.2 100.4 100.6 100.8 101.0 101.2 101.4 101.6 101.8 102.0 102.2 102.4
°C 37.1 37.2 37.3 37.4 37.6 37.7 37.8 37.9 38.0 38.1 38.2 38.3 38.4 38.6 38.7 38.8 38.9 39.0 39.1
°F 102.6 102.8 103.0 103.2 103.4 103.6 103.8 104.0 104.2 104.4 104.6 104.8 105.0 105.2 105.4 105.6 105.8 106.0 106.2
°C 39.2 39.3 39.4 39.6 39.7 39.8 39.9 40.0 40.1 40.2 40.3 40.4 40.6 40.7 40.8 40.9 41.0 41.1 41.2
Appendix Page 1014 Wednesday, May 23, 2001 11:12 AM
Appendix Page 1015 Wednesday, May 23, 2001 11:12 AM
Appendix B Abbreviations TABLE 1 Pharmacology Abbreviation
Latin Derivation
Meaning
ad lib BID c caps EOD gr H, h, hr IC ID IH IM IP IT IU IV k units mEq min NPO o oo OD OH OID OS pil PO or/os prn q or qq q4h qd or qqd qh or qqh QID qs Rx s or sig s SC, SQ or SubQ SID stat tab TID x 4d
ad libitum bis in die cum capsula
freely twice a day with capsule every other day grain hour intracardiac intradermal inhalation intramuscular intraperitoneal intratracheal international units intravenous thousand units milliequivalent minute nothing by mouth every other day ophthalmic ointment right eye every hour once a day left eye pill by mouth when desirable or necessary every every 4 hours every day every hour 4 times a day a sufficient amount take write (on label) without subcutaneous once a day immediately tablet three times a day give for 4 days
granum hora
non per os omnis ocular dexter omni hora ocular sinister pilula per os pro ne nata quaque quaque 4 horae quaque die quaque hora quarter in die quantum sufficit recipe signa sine singular in die statim tabella ter in die
1015
Appendix Page 1016 Wednesday, May 23, 2001 11:12 AM
1:32 1:1200 ++ − +++ − +++ +++ +++ +++ Neutral
1:200 1:750 ++ − +++ − ++ ++ − + Alkaline
Bacteriocidal Virucidal Lipophilic-virucidal Sporocidal Fungicidal Effective in presence of organic material Effective in presence of soap Effective in hard water Most effective in pH range
Key: – = least active or effective; +++ = most effective; ? = unknown.
Use
Dilution (disinf. water)
Lysol Staphene disinfectant Disinfectant
Phenolics
Roccal Zephiran disinfectant Disinfectant
Quaternary Ammonias
Brand names
.
Disinfectant Class
+++ +++ Neutral
External antiseptic Undiluted (3%) 4:20 ++ − +++ − +++ +++
pHisoHex
Bisphenols
++ ++ Neutral
+++ +++ ++ + +++ ++
External antiseptic Undiluted Undiluted
Betadine Metadine
Iodophores
++ ++ Acid
+++ +++ Alkaline
+++ +++ +++ ++ +++ +++
1:50
1:32 ++ +++ +++ − +++ +
Disinfectant
Cidex
Glutaraldehydes
Disinfectant
Chlorox
Sodium Hypochlorites
+ ? ?
External antiseptic Undiluted (70%) Undiluted ++ − ? − ++ +++
Isopropyl Ethyl
Alcohols
− ++ Alkaline
++ − +++ − ++ ++
1:28
Disinfectant
Nolvasan
Chlorhexidines
Characteristics of Common Disinfectants
Appendix C
Appendix Page 1017 Wednesday, May 23, 2001 11:12 AM
1017
Appendix Page 1018 Wednesday, May 23, 2001 11:12 AM
0839_frame_IDX Page 1019 Friday, May 25, 2001 9:37 AM
Index A AAAS (American Association for the Advancement of Science), 122 Abdomen, see Gastrointestinal system Abdominal imaging, 601 Acanthocephalan parasites, see also Helminths in cetaceans, 362 in pinnipeds, 370, 916 removal and fixation, 358 in sea otters, 23 Acanthocheilonema, 371, 372, 916, 919 Acanthosis in pinnipeds, 913, 915 in polar bears, 531–532 Acepromazine maleate, 994 Acetylcysteine, 714 Acetylpromazine, 709, 714 Acetylsalicylic acid, 719 Acoustics capabilities in cetaceans, 6–7 navigation ability loss, 86 pathology, 459–460 transmitters, 862–863 ACTH (adrenocorticotropic hormone), 169 Active tags for tagging and tracking, 852 Acute-phase proteins, 241 Acute-phase response (immune system), 238 Adaptive immune response, 238–239 Adenovirus described, 291–292 in pinnipeds, 916 Administrative Procedures Act, 759 ADMR (average daily metabolic rate), 792–794 Adrenal glands aldosterone, 181–182 catecholamine function and physiology, 178 cortisol levels, 178–181 cysts and nodules, 535 described, 177 gross anatomy, 148 microscopic anatomy, 150 Adrenocorticotropic hormone (ACTH), 169 Advanced training programs Web sites, 111–112 Aeromonas dermatological diseases caused by, 326 in manatees, 951 in pinnipeds and cetaceans, 312 Age estimation, 454–455
Agencies, regulating, 742–743; see also Federal legislation and regulations AH (aniline hydroxylase), 489 AHH (arylhydrocarbon hydroxylase), 489 Alanine aminotransferase (ALT), 406 Alaska Marine Mammal Tissue Archival Project, 465 Albumins, 414 ALDE (aldrin expoxidase), 489 Aldosterone, 181–182, 260 Aldrin, 484 Aldrin expoxidase (ALDE), 489 Alexandrium spp., 5, 16, 494 Algal blooms, 19, 21, 493, 951 Alimentary canal colonoscopy, 633 colon/rectum findings, cetacean cytology, 445 cytology findings, cetaceans, 445 fecal and urinary energy losses, 809–810 feces sample collection and handling, 166, 439 leukocytes evaluation in colon/rectum, 445 Alkaline phosphatase (ALP) hepatobiliary system disorders and, 409–410 inflammatory disease indication, 902–903 Alopecia in pinnipeds, 913, 915 in polar bears, 531–532, 1003 Altrenogest contraception use, 217, 222 drug dosage, cetaceans, 709 Aluminum hydroxide, 714 Amazonian manatee (Trichechus inunguis), see Manatee Amazon river dolphin (Inia geoffrensis) noninfectious diseases, 532 water requirements, 799–800 American Association for the Advancement of Science (AAAS), 122 American Public Health Association, 785 American Veterinary Medical Foundation (AVMF), 122 Amikacin bacterial disease treatment using, 324, 325 drug dosage cetaceans, 709 pinnipeds, 714 sea otters, 719 sirenians, 718 usefulness of, 699 Aminoglycosides adverse effects, 707 for septicemia treatment, 312 Aminopentamidine sulfate, 719
1019
0839_frame_IDX Page 1020 Friday, May 25, 2001 9:37 AM
1020
Aminophylline, 714, 719 Aminopropazine fumarate, 719 Aminopyrine N-demethylase (APDM), 489 Amoxicillin cetaceans drug dosage, 709 pinnipeds drug dosage, 714 sea otters drug dosage, 719 Amphotericin B drug dosage, cetaceans, 709 mycotic infection use, 349, 350 Ampicillin cetaceans drug dosage, 709 pinnipeds drug dosage, 714 polar bears drug dosage, 721 sirenians drug dosage, 718 Amylase, 405 Amyloidosis, 534 Analgesics, 699 Ancylostomatidae (hookworms), 369, 919 Anemia categorizing, 399–400 classification by RBC indices, 400–401 erythrocytes evaluation, 399–401 in pinnipeds, 919 Anesthesia anesthetic protocol, 655–656 cetaceans emergencies, 662 immobilizing agents used, 658–659 induction, 657, 660 inhalation anesthesia, 660 intubation, 660 monitoring, 660–661 support, 661–662 manatees, 950 monitoring techniques, 656–657 odobenids emergencies, 681 immobilizing agents used, 678–679 induction, 677, 680 intubation and inhalation anesthesia, 680 monitoring, 680 support, 680–681 otariids emergencies, 669–670 immobilizing agents used, 663–665 induction, 662, 666 inhalation anesthesia, 667 intubation, 666–667 monitoring, 668 support, 668–669 phocids emergencies, 676–677 immobilizing agents used, 671–673 induction, 670, 674 inhalation anesthesia, 675 intubation, 674–675 monitoring, 675
CRC Handbook of Marine Mammal Medicine
support, 675–676 polar bears carfentanil, 999 delivery and monitoring, 996–997 etorphine, 999 fentanyl citrate, 999 inhalation agents, 999–1000 ketamine, 997–998 ketamine/xylazine, 998 telazol/medetomidine, 999 tiletamine HCL, 998–999 zolazepam HCL, 998–999 sea otters emergencies, 684 immobilizing agents used, 682 induction, 681, 683 inhalation anesthesia, 683 intubation, 683 monitoring and support, 683 sirenians, 681 support, 657 ursids, 684 walruses, 930 Angiography, 563 Aniline hydroxylase (AH), 489 Animal and Plant Health Inspection Service (APHIS) pinniped husbandry requirements, 907–908 responsibilities of, 743, 759 transport regulations, 881 water quality oversight, 779, 783 Animal Welfare Act (AWA) responsibilities of, 743, 755–757, 759–760 water quality standards setting, 779 Anisakidae (large roundworms), 361–362, 369 Anklets for pinniped tagging and tracking, 859 Ankylosing spondylosis, 532 Anoplura sp. (lice), 372 Anorexia due to drug interactions, 707 importance as sign of illness, 896 therapy for, 694 ANP (atrial natriuretic peptide), 185 Antacids adverse effects, 705–706 Antarctic fur seal (Arctocephalus tropicalis) bacterial diseases in, 314, 320 parasites of, 368 Antennas in tracking systems, 852–853 Anterior pituitary gland, 194 Anthelmintics, 359 Anthracosis, 533 Anthropogenic effects, see also Boat-related maimings; Toxicology causes of disease, 458, 460 noise impact, 7 OC chemical contamination, 4–5 pathogens origins, 5 trauma treatment, 530–531
0839_frame_IDX Page 1021 Friday, May 25, 2001 9:37 AM
1021
Index
Antibiotics intensive care use, 698–699 nematode treatment in pinnipeds, 371 resistance to, 309 Antifungals, 708 Antiparasitic drugs, 708 Aonyx cinerea (Asian short clawed otter), 325 APDM (aminopyrine N-demethylase), 489 APHIS (Animal and Plant Health Inspection Service) responsibilities of, 743, 759 transport regulations, 881 water quality oversight, 779, 783 Apicomplexans in cetaceans, 359–360 in manatees, 372 in pinnipeds, 367–368 in sea otters, 373 Apnea after anesthesia administration odobenids, 680 phocids, 676 during anesthesia, 661 Apophysomyces elegans cytology findings in respiratory tract, 443 lesions, diagnostic methods, and treatments, 345–346 Appetite stimulants, 694 Archival tags, 853 Arctocephalus spp., see Fur seal Argentine stranding networks contacts and programs, 46–47 Arginine vasopressin (AVP), 183–184 Arsenic, 482 Artificial insemination of cetaceans, 219–221 Artificial markings as tags, 851 Artificial milk formulas, see Hand-rearing and artificial milk formulas Arylhydrocarbon hydroxylase (AHH), 489 Ascorbic acid drug dosage cetaceans, 710 polar bears, 721 sea otters, 719 sirenians, 718 nutritional deficiency and, 816 Asexual fungi, see Mycotic diseases Asian elephant (Elephas maximus), 211 Asian short clawed otter (Aonyx cinerea), 325 Aspartate aminotransferase (AST), 407 Aspergillus fumigatus clinical diagnosis, 349 cytology findings in respiratory tract, 443 lesions, diagnostic methods, and treatments, 341–342 mechanisms of pathogenesis, 339 Atipamezole, 670 Atlantic walrus (Odobenus rosmarus), 314
Atlantic white-beaked dolphin (Lagenorhynchus albirostris) bacterial diseases in, 317 viral diseases in morbillivirus, 296 rhabdoviruses, 303 Atlantic white-sided dolphin (Lagenorhynchus acutus) bacterial diseases in, 314 diseases in, 19 hematology and biochemistry values, 392 mass strandings investigation, 86–87 parasites of, 366 reproductive cycle, 205–206 reproductive maturity, 204 reproductive parameters, 213–214 seasonality of fertility, 216–217 sexual maturity, 215 viral diseases in papillomavirus, 290 poxvirus, 286 Atrial natriuretic peptide (ANP), 185 Atropine drug dosage cetaceans, 710 sea otters, 719 for phocids, 670 for sea otters, 683 Attachment methods for tags, 866 Australian fur seal, 534 Australian sea lion (Neophoca cinerea), 320 Australian stranding networks contacts and programs, 47–48 Average daily metabolic rate (ADMR), 792–794 AVMF (American Veterinary Medical Foundation), 122 AVP (arginine vasopressin), 183–184 AWA (Animal Welfare Act) responsibilities of, 743, 755–757, 759–760 water quality standards setting, 779 Azithromycin adverse effects, 707 drug dosage, 710 Azoles adverse effects, 708 mycotic infection use, 349, 351 Azospermia, 218
B Bacterial diseases and infections brucellosis in cetaceans, 313–314 diagnosis, 313 epidemiology, 312–313 pathophysiology, 313 in pinnipeds, 314 current investigations into, 309–310 dermatological, 326–327
0839_frame_IDX Page 1022 Friday, May 25, 2001 9:37 AM
1022
Erysipelothrix in cetaceans, 316–318 diagnosis, 316 in pinnipeds, 318–319 gastrointestinal, 327–328 leptospirosis, 320–321 microbial sampling techniques, 310–311 mycobacterial disease, 319–320 Nocardia in cetaceans, 322, 324–325 clinical signs, 321–322 in pinnipeds, 325 species infected, 323 pasteurellosis, 315–316 in polar bears, 1001 public health significance Brucella, 771 Clostridium, 770 Coxiella burnetii, 772 Edwardsiella, 770 Erysipelothrix, 771 Leptospira, 770 mixed infections, 772 Mycoboacterium, 771–772 Streptococcus, 770–771 Vibrio, 769–770 respiratory, 325–326 in sea otters, 979 septicemia, 312 urogenital, 327 vibriosis, 314–315 Bacula, 201 Baikal seal (Phoca sibirica) osmoregulatory hormones, 184 viral diseases in, 296 Balaena mysticetus, see Bowhead whale Balaenoptera acutorostrata, see Minke whale borealis, see Sei whale edeni (Bryde’s whale), 532 musculus, see Blue whale physalus, see Fin whale Baleen whales, see also Cetaceans; Whales age estimation, 455–456 mercury concentration in liver, 478 parasites of, 366 PCB toxicity and, 486 sexual dimorphisms, 157 trematode treatment, 365 Ballistics use for euthanasia, 734–736 Band neutrophils, 444 Barbiturates, 732 Basal metabolic rate (BMR), 792–794 Basophils, 402 Baylisascaris spp., 374, 1002, 1003 Beaked whale, 534 Bearded seal (Erignathus barbatus) parasites of, 368
CRC Handbook of Marine Mammal Medicine
viral diseases in, 293 Belgium stranding networks contacts and programs, 48–49 Beluga (Delphinapterus leucas) aldosterone levels, 181 bacterial diseases in Erysipelothrix, 317 Nocardia, 322 clinical pathology, 391 contaminants’ effects on organ systems, 474, 475 cortisol concentrations, 181 cortisol levels, 178 hepatobiliary system disorders markers, 409 leukocytes evaluation, 402 lymphoid and hematopoietic system, 150 neoplasm studies, 7 non-infectious diseases endocrine system, 535 nervous system, 537 organochlorine reproductive function effects, 491, 492 papillomavirus found in, 19 parasites of, 359 PCB toxicity and, 486 population structure determination, 274 pregnancy, 212 reproductive cycle, 206 reproductive maturity, 205 reproductive parameters, 213–214 seasonality of fertility, 217 sentinel role, 6 sexual maturity, 216 stress response, 256, 260 thyroid gland and, 177 thyroid ratio, 170 viral diseases in herpesviruses, 292 papillomavirus, 289–290 Benzene hexachloride (BHC), 485 Benzo(a)pyrene monooxygenase (BPMO), 489 Benzodiazepines, 657 BHC (benzene hexachloride), 485 Bile acids, 411 Biliary system, see also Hepatobiliary system disorders markers helminth parasites in pinnipeds, 372 sonography clinical applications, 602 Bilirubin, 410–411 Bioenergetic scheme fecal and urinary energy losses, 809–810 gross energy requirements calculation, 810–811 heat increment of feeding, 807–808 ingested energy, 803–804 lactation, 805–807 maintenance energy, 804 molt, 807 production, 804–805 Bioindicators, see Sentinel systems
0839_frame_IDX Page 1023 Friday, May 25, 2001 9:37 AM
Index
Biological filtration methods for pool waters, 781–782 Biomarkers, 6 Biopsies sample collection, pinnipeds, 911 Biotoxins brevetoxin, 493–494 ciguatera, 496 domoic acid, 495 paralytic shellfish poisoning, 494–495 specimen collection, 468 Biphenyl 4-hydroxylase (biph-4OH), 489 Bithionol, 710 Bivalves sentinel role, 4 Blastomyces dermatitidis lesions, diagnostic methods, and treatments, 347 mechanisms of pathogenesis, 338 Blastomycosis in polar bears, 1001–1002 Blepharipoda spp., 23 Blood collection from cetaceans, 90, 385 equipment and processing, 384–385 manatees, 946–947 pinnipeds, 910 polar bears, 995 sample handling, 166 sites, 385–390 gas parameters and anesthesia depth determination, 661 hematology overview, 390 references, 397–399 values per species, 391–396 hematopoietic system, 150 hemoglobin content, see Erythrocytes, evaluation hemostatic parameters blood types, 420 disorders screening, 420–421 prothrombin time, 421–422 transfusions, 692 types, 420 Blowhole, cetacean, 900 Blubber finger, 772–773 Blubber layer cetaceans, 140 organohalogens accumulation in, 483–484 of polar bears, 989 thermal insult protection during diving, 153 Blue whale (Balaenoptera musculus), 290 BMR (basal metabolic rate), 792–794 Boat related maimings of manatees animal rights considerations, 35 database of scarred individuals, 33 laws pertaining to, 36 mortalities, 31–33 multiple cuts evidence, 33–35 trauma treatment, 954–957 Body condition determination by sonography, 618 Bolbosoma, 362
1023
Bona fide research definition, 760 Bone marrow, 158, 423 Bottlenose dolphin (Tursiops truncatus) adrenal glands studies, 178 bacterial diseases in Brucella, 313–314, 771 Clostridium, 770 Erysipelothrix, 317 Nocardia, 322 respiratory, 326 urogenital, 327 blood transfusion, 692 clinical pathology hematology and biochemistry values, 393 infection cases, 424–425 contaminants’ effects on organ systems, 473, 475 corpora albicantia in, 210 cortisol concentrations, 181 cytology, see Cytology, cetacean estrous cycle, 206–208 external features, 139 fasting, 803 gross anatomy illustration, 136–137 hepatobiliary system disorders markers, 406, 409, 410 intubation for anesthesia, 660 lactational suppression of estrus, 209 lactation energy requirements, 805 lead concentrations in, 481 leukocytes evaluation, 401–403 locomotion metabolic expenditure, 797 lymphoid and hematopoietic system, 150 mercury concentration in liver, 478 milk fat content, 807 mycotic infection, 338 non-infectious diseases digestive system, 534 integumentary system, 531 musculoskeletal system, 532 nutrition thiamine deficiency, 813 vitamin deficiencies, 816 organohalogens accumulation in, 488 osmoregulatory hormones, 183 ovarian physiology, 206–208 parasites of, 359, 360 parathyroids, 144 pectoral limb complex, 157 pregnancy, 211, 212 pseudopregnancy, 210 rehydration, 691, 692 renin-antiotensin system, 185 reproductive cycle, 205, 211, 212 reproductive maturity, 204 reproductive parameters, 213–214 seasonality of fertility, 216 serum analytes and enzymes, 405 sexual maturity, 215
0839_frame_IDX Page 1024 Friday, May 25, 2001 9:37 AM
1024
stranding, 69 stress response, 259, 260 thermal imaging applications, 648 thermoneutral zone, 795 TH fluctuations and, 177 thyroid, 143, 170 thyroxine levels, 174 viral diseases in caliciviruses, 300 morbillivirus, 18–19, 296 papillomavirus, 19, 290, 291 poxvirus, 286 water requirements, 800 Bowhead whale (Balaena mysticetus) age estimation, 455–456 bacterial diseases in, 326–327 non-infectious diseases digestive system, 534 integumentary system, 531 parasites of, 360 sentinel role, 6 viral diseases in adenoviruses, 291 caliciviruses, 300 BPMO (benzo(a)pyrene monooxygenase), 489 Bracelets for pinniped tagging and tracking, 859 Bradycardia after anesthesia administration, 669 Brain, see Neurological dysfunction Branding for tracking cetaceans, 862 manatees, 867 pinnipeds, 858 Brandt’s cormorant (Phalacrocorax penicillatus), 495 Braunina cordiformis, 444 Brazilian stranding networks contacts and programs, 49–50 Brevetoxicosis impact on manatees, 23 manatees, 951 Brevetoxin, 493–494 Bromsulphalein, 715 Bronchopneumonia due to non-infectious diseases, 533 in pinnipeds and cetaceans, 325–326 Brown pelican (Pelecanus occidentalis), 495 Brucella spp. cetacean infection case, 424–425 in manatees, 952 in pinnipeds, 22 public health significance, 771 in walruses, 935 Brucellosis in cetaceans, 19, 313–314 current investigations into, 309 diagnosis, 313 epidemiology, 312–313 pathophysiology, 313 in pinnipeds, 22, 314
CRC Handbook of Marine Mammal Medicine
Bryde’s whale (Balaenoptera edeni), 532 Bulk fluid sterilization, 784 BUN (serum urea nitrogen), 411–412 Buoyancy in cetaceans, 897–898 Burmeister’s porpoises (Phocoena spinipinnis) papillomavirus found in, 19, 290, 291 poxvirus found in, 286, 289 Butorphanol, 699 Butter clam (Saxidomus giganteus), 494 Butyltins concentrate in tissues, 481–482
C CA (Corpora albicantia), 210 Cadmium, 480–481 Calcium, 418 deficiencies in polar bears, 1000 gluconate, for nutritional therapy, 694 Calculi, 534, 920 Caliciviruses (San Miguel sea lion virus) described, 300–302 in pinnipeds, 920 public health significance, 768–769 serodiagnostics of, 244 in walruses, 935 California gray whale (Eschrichtius robustus) caliciviruses in, 300 fasting during lactation, 803 hematology and biochemistry values, 393 California sea lion (Zalophus californianus) analgesics use, 699 bacterial diseases in dermatological, 326 Leptospira, 770 leptospirosis, 320 respiratory, 325 urogenital, 327 biotoxins effects on, 495 contaminants’ effects on organ systems, 474 estrous cycle, 195 external features, 138 fasting during lactation, 803 gross anatomy illustration, 130–131 hand-rearing and artificial milk formulas, 839–840 hematology and biochemistry values, 394–395 hyponatremia care, 696 lactation energy requirements, 805, 806 locomotion metabolic expenditure, 796 mortality from algal blooms, 19, 21 non-infectious diseases cardiovascular system, 536 digestive system, 534 genitourinary system, 534 integumentary system, 531 musculoskeletal system, 532, 533 nervous system, 536, 537 nutrition thiamine deficiency, 813
0839_frame_IDX Page 1025 Friday, May 25, 2001 9:37 AM
Index
vitamin deficiencies, 815 organochlorine reproductive function effects, 490 parasites of, 368, 369, 370, 372 rehydration, 691 renin-antiotensin system, 185 reproductive characteristics, 196 sentinel role, 6 SMSV and, 769 thermal imaging applications, 646 thermoneutral zone, 794–795 thyroids, 145 urogenital neoplasia prevalence, 21 viral diseases in adenoviruses, 291, 292 caliciviruses, 300 herpesviruses, 292, 293 poxvirus, 286, 287, 288 California sea otter (Enhydra lutris) non-infectious diseases, 533, 535 parasites as cause of disease in, 23 thermal insult protection, reproductive organs, 201 Callorhinus ursinus, see Northern fur seal Cameras in endoscopy, 629–630 thermal imaging, 645 Canadian stranding networks contacts and programs, 51 Canaries in mines, 3, 4 Candida spp. cytology findings in respiratory tract, 443 epidemiology, 338 lesions, diagnostic methods, and treatments, 342–343 mechanisms of pathogenesis, 338–339 in polar bears, 1002 species affected, 338 Canine distemper virus (CDV), 296, 917 Cannulas, 638–639 Captive animals and water quality, see Water and environment quality Capture myopathy, sea otters, 979 Carbenicillin cetacean drug dosage, 710 polar bear drug dosage, 721 Carbon dioxide use for euthanasia, 734 Carcass condition classification, 451 Carcass condition code, 453 Carcass disposal after euthanasia, 736–737 Cardiovascular system contaminants’ effects on, 472 diagnostic imaging techniques, 571 gross anatomy, 142 noninfectious diseases, 535–536 pinniped diseases, 919 rate and anesthesia depth determination, 661 rate monitoring in stranded cetaceans, 89 sonography clinical applications, 596–597
1025
Careers in marine mammal medicine advanced training programs, 111–112 federal government job listings, 123 fellowships, 112 full-time employment, 97–98 graduate degree programs, 109–110 internships and residencies at aquaria or rehabilitation centers, 108–109 at institutions or agencies, 109 matched internships, 107–108 matched residencies, 108 at zoos, 109 job search method action plan, 102 communication skills, 106–107 important attributes of applicants, 106 interview preparation, 103–106 jobs locations research, 102–103 self-assessment, 99–100 skill categorization, 100–101 part-time employment, 98 personality traits and factors in success, 98–99 recommendations for search, 113 related programs, 110–111 scientific societies and membership organizations, 112–113 Carfentanil for polar bears, 999 Caribbean stranding networks contacts and programs, 52–53 Carprofen, 719 Carson, Rachel, 3 Caspian seal (Phoca caspica), 296 Castor oil, 715 Castration of pinnipeds, 202 Catecholamines function and physiology, 178 stress response components, 256 Cats, see Dogs and cats Caval sphincter, 144 CDV (canine distemper virus) morbillivirus virus, 296 in pinnipeds, 917 Cefazolin, 719 Ceftriaxone cetacean drug dosage, 710 sirenian drug dosage, 718 Cefuroxime cetacean drug dosage, 710 pinniped drug dosage, 715 Cellular immunity, 241–242 Census definition, 37 Central nervous system cerebrospinal fluid collection, 911 contaminants’ effects on, 473 described, 150–151 emergency care, 696 non-infectious diseases, 536–537, 536–537 pinniped diseases, 921–922
0839_frame_IDX Page 1026 Friday, May 25, 2001 9:37 AM
1026
Cephalexin pinniped drug dosage, 715 sea otter drug dosage, 719 sirenian drug dosage, 718 Cephalexin monohydrate, 710 Cephalorhynchus commersoni (Commerson’s dolphins), 391 Cephalorhynchus hectori (Hector’s dolphin), 286 Cephaloridine, 715 Cephalosporins adverse effects, 707 for septicemia treatment, 312 usefulness of, 698 Cephloridine, 710 Cerebrospinal fluid collection, 911 Cestodes, see also Helminths in pinnipeds, 370 in polar bears, 374 removal and fixation, 358 treatment of, 359 Cetacean reproduction abnormalities, 218–219 artificial insemination cautions for, 219 semen collection and storage, 219–221 female asymmetry of ovulation, 210 contraception and control of aggression, 217 corpora albicantia, 210 estrous cycle, 206–209 lactational suppression of estrus, 209–210 ovarian physiology, 206–209 parturition, 212, 215 pregnancy, 211–212 pregnancy diagnosis, 212 pseudopregnancy, 210–211 reproductive cycle, 205–206 reproductive maturity, 204–205 future applications for management, 224–225 insemination techniques, 223–224 male contraception and control of aggression, 218 seasonality of fertility, 216–217 sexual maturity, 215–216 ovulation manipulation and control induction, 221–222 synchronization, 222–223 reproductive emergencies, 697 reproductive parameters, 213–214 reproductive tract, 150 Cetaceans, see also specific species adrenal glands, 150 age estimation, 455–456 aldosterone concentrations, 182 analgesics use, 699 anesthesia emergencies, 662 immobilizing agents used, 658–659
CRC Handbook of Marine Mammal Medicine
induction, 657, 660 inhalation anesthesia, 660 intubation, 660 monitoring, 660–661 support, 661–662 bacterial diseases in Brucella, 313–314 dermatological, 326–327 Erysipelothrix, 316–318 gastrointestinal, 327–328 mycobacterial disease, 319 Nocardia, 322, 324–325 pasteurellosis, 315 respiratory, 325–326 urogenital, 327 Vibrio, 770 vibriosis, 315 bone marrow, 158 cadmium toxicosis, 480–481 circulatory structures, 151, 152 clinical examination blowhole, 900 body weight, 900 buoyancy, 897–898 hands-on examination, 899 history, 896–897 how animal feels, 897 laboratory tests, 900–901 milk samples, 899 radiography use, 900 social behavior, 898–899 stool samples, 899 ultrasonography use, 900 urine collection, 899 visual, 897 clinical pathology blood collection, 385 hematology and biochemistry values, 391–393 inflammatory disease indications, 901–903 pneumonia presentation, 901 references, 397 cortisol concentrations, 179, 181 cytology, see Cytology, cetacean diagnostic imaging techniques for, 566 digestive efficiencies, 810 digestive system, 146, 149 drug dosages, 709–714 emerging and resurging diseases in morbillivirus epizootics, 16, 18–19 table of, 17–18 viral disease, 19 endoscopy clinical applications colonoscopy, 633 gastroscopy, 630–633 respiratory, 633–635 urogenital, 635 euthanasia by ballistics, 734–736 hand-rearing and artificial milk formulas, 829–831
0839_frame_IDX Page 1027 Friday, May 25, 2001 9:37 AM
Index
hearing and sound production capabilities, 6–7 heart and pericardium gross, 142 hepatobiliary system disorders markers, 408, 409 integument microanatomy, 139–140 larynx, 144 leukocytes evaluation, 402 lymphoid and hematopoietic system, 150 mass strandings current investigations into, 86–87 definition, 83 disposition of animals, 92, 94 east coast of U.S. (1987-99), 84–85 evaluation of, 87 graded histopathological findings, 93 management of, 88–92 serum chemistry findings, 91 theories to explain, 83, 85–86 treatment precautions, 94 metabolic requirements estimation, 793 minimally invasive surgical techniques, 640 need for decisions and action, 895–896, 905 nervous system, 151 non-infectious diseases cardiovascular system, 535–536 integumentary system, 531 musculoskeletal system, 532–533 nutrition therapy for, 694 vitamin deficiencies, 816 parasites of apicomplexans, 359–360 ciliates, 359 flagellates, 360 helminths, 361–367 sarcodina, 360 parathyroids, 144 PCB toxicity and, 486 pectoral limb complex, 156–157 pleura and lungs, 143 population estimation procedures, 37 regulating agency, 742 rehydration, 691 reproduction, see Cetacean reproduction respiratory emergencies, 695 respiratory system, 144 ribs, 155 salinity and pH requirements, 781 skeletal muscles use for swimming, 141 space requirements, 780 species status under ESA, 746–748 stress response, 259 tagging and tracking attachment methods, 863, 864, 866 branding, 862 data loggers, 863 impact of devices, 866 implantable tags, 866 passive tags, 862
1027
roto-radio tag, 863–864 telemetry device location and attachment, 862–863 therapeutics medical therapy, 903–905 surgery, 903 thermal imaging applications, 647–649 thermal insult protection during diving, 153 thyroids, 145 thyroxine concentrations, 171 thyroxine levels, 170 transport body support techniques, 882 equipment requirements, 886–887 ground and air, 885–886, 887 improvements in, 884 removal from transport unit, 888 stretcher use, 884–885 temperature control, 882, 884 water requirements, 885 triiodothyronine concentrations, 173 urinary tract, 147 vertebrae, 156 viral diseases in herpesviruses, 295 morbillivirus, 296–298 poxvirus, 286 CFR (Code of Federal Regulations), 742, 760 CG (chorionic gonadotropin), 199 Charcoal, activated, 719 Chediak-Higashi syndrome, 403 Chelonibia manatii, 953 Chevron bones, 156 Chilodonella, 359 Chilomastix, 360 Chloramphenicol adverse effects, 707 drug dosage cetaceans, 710 pinnipeds, 715 polar bears, 721 Chlordane, 484 Chlordiazepoxide HCL, 710 Chloride, 417 Chlorinated norbornane, 485 Chlorine-based oxidation, 784–785 Cholelithiasis, 534 Cholesterol, serum analytes and enzymes, 404–405 Chorionic gonadotropin (CG), 199 Chromosome heteromorphism analysis, 275–276 Ciguatera, 496 Ciliates, 359 Cimetidine adverse effects, 705–706 drug dosage cetaceans, 710 pinnipeds, 715 sea otters, 719
0839_frame_IDX Page 1028 Friday, May 25, 2001 9:37 AM
1028
Ciprofloxacin cetacean drug dosage, 711 pinniped drug dosage, 715 Circulatory system diseases of, 370–372 structures of, 151–152 CITES (Convention on International Trade in Endangered Species of Wild Fauna and Flora), 746, 750, 760, 881 CK, CPK (creatinine phosphokinase), 420 CL, see Corpus luteum Cladophialophora bantiana, 343 Clavulanic acid cetacean drug dosage, 709 pinniped drug dosage, 714 Clindamycin cetacean drug dosage, 711 pinniped drug dosage, 715 Clinical pathology abnormalities and artifact determination, 383–384 blood collection sampling equipment and processing, 384–385 sites, 385–390 bone marrow evaluation, 423 clinical cases bottlenose dolphins, 424–425 harbor seal, 427–428 killer whale, 425–426 manatees, 428–429 Pacific white-sided dolphin, 426–427 sea otters, 429 electrolytes calcium, 418 chloride, 417 magnesium, 419 phosphorus, 419 potassium, 416–417 sodium, 416 total carbon dioxide, 417–418 erythrocytes evaluation anemia, 399–401 indices, 391–393, 396, 399 hematology overview, 390 references, 397–399 values per species, 391–396 hemostatic parameters blood types, 420 disorders screening, 420–421 prothrombin time, 421–422 hepatobiliary system disorders markers γ-glutamyltransferase, 408–409 alanine aminotransferase, 406 alkaline phosphatase, 409–410 aspartate aminotransferase, 407 bile acids, 411 bilirubin, 410–411 glutamate dehydrogenase, 407–408
CRC Handbook of Marine Mammal Medicine
lactate dehydrogenase, 408 sorbitol dehydrogenase, 407–408 inflammation markers erythrocyte sedimentation rate, 422 serum iron, 422–423 kidney-associated serum analytes creatinine, 412 urea nitrogen, 411–412 leukocytes evaluation age and, 402 basophils, 402 disease and, 403 eosinophils, 401–402 heterophils, 401 lymphocytes, 402 monocytes, 402 neutrophils, 401 serum analytes creatinine phosphokinase, 420 uric acid, 420 serum analytes and enzymes amylase, 405 cholesterol, 404–405 glucose, 403–404 lipase, 405 lipids, 404 pancreatic enzymes, 404 triglycerides, 404–405 trypsin-like immunoreactivity, 405 serum proteins albumins, 414 globulins, 414–415 hematocrit, 413 total plasma protein, 413 urinalysis, 423 Cloprostenol, 199 Clostridial diseases, 327–328 Clostridium spp. in cetaceans, 327–328 in manatees, 951 in pinnipeds, 915–916 public health significance, 770 in walruses, 935 Coccidia in manatees, 372 removal and fixation, 358 Coccidioides immitis lesions, diagnostic methods, and treatments, 347 mechanisms of pathogenesis, 338 in pinnipeds, 916, 918 Coccobacilli, 314 Cochleotrema cochleotrema, 953 Code of Federal Regulations (CFR), 742, 760 Cold stress syndrome, 951–952 Coliform bacteria, 783 Colloid goiter, 535 Colon/rectum, see Alimentary canal
0839_frame_IDX Page 1029 Friday, May 25, 2001 9:37 AM
Index
Commerson’s dolphin (Cephalorhynchus commersoni), 391 Common dolphin (Delphinus delphis) bacterial diseases in, 313–314 hematology and biochemistry values, 391 morbillivirus found in, 18–19 non-infectious diseases, 536 thermoneutral zone, 795 tremotode as cause of strandings, 365 viral diseases in morbillivirus, 296 papillomavirus, 290 poxvirus, 286, 289 water requirements, 800 Computed tomography (CT) anatomy, 586 described, 558–559 indications for use, 563 Computer radiography (CR), 557–558 Congressional Record, 742 Connective tissue helminth parasites in pinnipeds, 372 parasitic diseases in cetaceans and, 366–367 Conservation medicine, 98 Contaminants, see also Halogenated organics chemical pollutants speciman collection, 466, 468 by feral and domestic cat populations, 24 oil constituents, 496 contact exposure effects, 496–498 exposure problems for sea otters, 976–977 oral-dosing studies, 498 petroleum constitutents in animals, 498–499 thyroid pathology association, 177 in water, effects on organ systems, 472, 473, 475 Contracaecum corderoi, 22, 369 Contraception and control of aggression cetaceans, 217–218 pinniped reproduction females, 202 males, 202–203 purpose of, 201 Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES), 746, 750, 760, 881 Copper, 482 Copper sulfate, 711 Coronavirus harbor seal infection case, 302 pinniped infection case, 427–428 Corpora albicantia (CA), 210 Corpus luteum (CL) estrous cycle and, 196 pinniped hormonal profiles, 197 in polar bears, 992 reproduction regulation role, 194 sonography imaging, 615 Corticols, see Glucocorticoids
1029
Corticosterone, see Glucocorticoids Corticotropin-releasing factor (CRF), 255 Cortisol levels, 178–181 Corynebacterium in pinnipeds and cetaceans, 312, 327 in walruses, 931 Corynosoma spp. in cetaceans, 362 in pinnipeds, 370 in sea otters, 23, 373, 978 Coxiella burnetii in pinnipeds, 920 public health significance, 772 CR (computer radiography), 557–558 Crabeater seal (Lobodon carcinophagus), 296 Crassicauda, 366 C-reactive protein (CRP), 241 Creatinine, 412 Creatinine phosphokinase (CK, CPK), 420 Crenosomatidae, 370 CREN Web site, 121 CRF (corticotropin-releasing factor), 255 Crittercams, 857 Croatian stranding networks contacts and programs, 52 CRP (C-reactive protein), 241 Cryopreservation of cetacean semen, 220 Cryptococcus neoformans lesions, diagnostic methods, and treatments, 343–344 mechanisms of pathogenesis, 339 in pinnipeds, 918 species affected, 338 Cryptosporidium spp. in pinnipeds, 22 public health significance, 774 CT, see Computed tomography Cutaneous viral papillomatosis, 23 Cyproterone, 719 Cystic follicles in cetaceans, 218 Cystoisospora delphini, 360 Cystophora cristata, see Hooded seal Cytochrome P-450, 489 Cytokines, 239–240 Cytology, cetacean colon/rectum findings, 445 interpretation, 441–442 overview, 437–438 respiratory tract findings, 442–443 sample collection aspirates from masses, 439 fecal, 439 gastric, 438–439 guidelines, 438 respiratory tract, 438 urinary tract, 439 slide preparation, 439–440 specimen examination, 441 stomach findings, 444 urinary tract findings, 445–446
0839_frame_IDX Page 1030 Friday, May 25, 2001 9:37 AM
1030
D Dalhousie University Web site, 121 Dall’s porpoise (Phocoenoides dalli) contaminants’ effects on organ systems, 474, 475 organochlorine reproductive function effects, 491 Data loggers in tracking, 863 Dawn liquid detergent, 719 DDE, 485 DDT (dichlorodiphenyl trichloroethane), 3, 484, 485, 920; see also Organochlorine (OC) chemicals Decarbamoyl saxitoxin, 21 Delphinapterus leucas (beluga), 6 Delphinidae, see Cetaceans Delphinus delphis, see Common dolphin Demodex (mites), 372 Denmark stranding networks contacts and programs, 53 Denning, 990, 992, 994 Dental system diagnostic imaging techniques, 571 non-infectious diseases, 532–533 polar bear diseases, 1003 sea otters, 980 surgical repair, 695 teeth age estimation use, 455–456 anesthesia for extractions, 657 polar bear diseases, 1003 of polar bears, 995 of sea otters, 963 walruses, 928 Dermatological diseases described, 326–327 in polar bears, 1002–1003 Dermatophilus congolensis, 288, 1002–1003 Dermatophyte infections, 338 Deslorelin, 218, 719 Detomidine, 670 Deuteromyces, 344 Dexamethasone adverse effects, 708 drug dosage cetaceans, 711 pinnipeds, 715 polar bears, 721 sea otters, 719 sirenians, 718 nematode treatment in pinnepeds, 370, 371 Dextrose pinniped drug dosage, 715 sea otter drug dosage, 719 Diagnostic imaging, see also Radiology; Ultrasonography application of techniqes data archiving and retrieval, 554–555 physics involved, 555
CRC Handbook of Marine Mammal Medicine
positioning and orientation standardization, 554 post-mortem examination, 555 imaging science, 551–552 limitations to techniques, 552 Diaphragm, 142 Diazepam for nutritional therapy, 694 for otariids, 666, 669, 670 for polar bears, 994 for sea otters, 683 DIC (disseminated intravascular coagulation), 421 Dichlorodiphenyl trichloroethane (DDT), see DDT Dichlorvos cetacean drug dosage, 711 pinniped drug dosage, 715 polar bear drug dosage, 721 Dieldrin, 484, 485 Diet, see Nutrition Digestive system gross anatomy, 145–147 of manatees, 941 microscopic anatomy, 148–149 non-infectious diseases, 533–534 pinniped diseases, 916–917 Dihydrostreptomycin, 711 Dimercaptosuccinic acid, 711 Dinoflagellates, 493 Dinophysis acuta, 494 Diphenhydramine, 719 Diphenoxylate, 719 Diphenylhydantoin, 719 Diphyllibothrium, 370 Diplogonoporus tetrapterus, 978 Dirofilaria, 371, 919 Diskospondylitis, 532 Disophenol, 715 Disorders screening, 420–421 Disseminated intravascular coagulation (DIC), 421 Diuretics, 708 Diving adrenal glands and, 178 circulatory structures and, 151–152 thermal insult protection, reproductive organs, 152–153 DMV (dolphin morbillivirus), 16, 296 DNA fingerprinting, 272 DNA sequencing, 271–272 functions, 273 sample collection and handling, 278 Dogs and cats hepatobiliary system disorders markers, 410 serum analytes and enzymes, 405, 406 water contamination by, 24 Dolphin morbillivirus (DMV), 16, 296 Dolphins, see also Cetaceans; specific species circulatory structures, 151 diagnostic imaging applications MRI, 587–589
0839_frame_IDX Page 1031 Friday, May 25, 2001 9:37 AM
1031
Index
radiographic anatomy, 574–579 radiographic pathology, 579–581 digestive system, 146, 148–149 external features, 139 nutritional therapy, 693 organochlorine effects on, 489 organohalogens accumulation in, 487 pleura and lungs, 143 sentinel role, 4 thermal insult protection during diving, 153 thyroxine levels, 170 Domoic acid described, 495 gross necropsy investigation, 467 morbidity and mortality induced by, 19 in pinnipeds, 921 in sea lions, 7, 495 Doxapram for odobenids, 680 for otariids, 669 Doxycycline cetacean drug dosage, 711 polar bear drug dosage, 721 seal finger treatment, 773 Drug dosages cetaceans, 709–714 pinnipeds, 714–718 polar bear, 721–722 sea otters, 719–721 sirenians, 718–719 Drug interactions aminoglycosides, 707 antacids, 705–706 antifungals, 708 antiparasitic drugs, 708 azithromycin, 707 cephalosporins, 707 chloramphenicol, 707 cimetidine, 705–706 diuretics, 708 florfenicol, 707 fluoroquinolones, 706 rifampin, 707 steroids, 708 sulfonamides, 707 tetracyclines, 706 Drug routes, 704–705 Dry mustard, 715 Dugongs (Dugong dugon) FWS jurisdiction, 742 organohalogens accumulation in, 487 population estimation procedures, 37 species status under ESA, 749 vertebrae, 156 Dusky dolphin (Lagenorhynchus obscurus) herpesviruses found in, 292 morbillivirus found in, 18–19 papillomavirus found in, 19, 290, 291
poxvirus found in, 286, 289 Dyspnea therapy, 695 Dystocia care, 219, 697–698
E E2OH (estradiol-2-hydroxylase), 489 EAAM (European Association for Aquatic Mammals), 905 Echinophthirius horridus (seal louse), 372, 919 Echocardiography, 596 ECOD (ethoxycoumarin-o-deethylase), 489 Ectoparasites in pinnipeds, 372 Edwardsiella spp. in manatees, 951 in pinnipeds and cetaceans, 312 public health significance, 770 Eimeria phocae, 367 Electrolyte regulation, 181 Electrolytes calcium, 418 chloride, 417 magnesium, 419 phosphorus, 419 potassium, 416–417 sodium, 416 total carbon dioxide, 417–418 Electronic information gathering precautions for, 117–118 reference databases evidence-based medicine, 120 federal government listings, 123 fellowships, 122 foundations, 123 general biomedical, 118–119 grants, 122–123 information sites, 121 listservs, 120–121 meetings and proceedings, 123 miscellaneous, 123–125 model Web sites, 119–120 online journals and textbooks, 121–122 veterinary medical sites, 118–119 Elephant seal (Mirounga angustirostris) analgesics use, 699 bacterial diseases in, 318, 320 contaminants’ effects on organ systems, 472 contraception, 203 fasting, 801 hand-rearing and artificial milk formulas, 836–838 heat increment of feeding, 808 hematology and biochemistry values, 394–395 hepatobiliary system disorders markers, 409 integument microanatomy, 140 liver, 148 molt energy requirements, 807 muscle mass, 141
0839_frame_IDX Page 1032 Friday, May 25, 2001 9:37 AM
1032
non-infectious diseases cardiovascular system, 536 genitourinary system, 534 integumentary system, 531 osmoregulatory hormones, 183–184 parasites of, 370, 371, 372 pineal gland size, 167 rehydration, 691 sexual dimorphisms, 157 thyroid gland, 145, 176 viral diseases in, 300 water requirements, 800 ELISA (enzyme-linked immunosorbent assay), 245 El Niño, 16 Emaciation of pups, 533 therapy for, 693–694 Embryonic diapause, 195, 198 Emerging and resurging diseases in cetaceans morbillivirus epizootics, 16, 18–19 table of, 17–18 in manatees, 22–23 modes of disease emergence, 15–16 in pinnipeds bacterial, 22 domoic acid-induced, 19, 21 table of, 20–21 urogenital neoplasia examination, 21–22 in polar bears, 24 sample processing, 25 in sea otters, 23–24 significance of, 24–25 Emerita spp. (crabs), 23 Employment in marine mammal medicine, see Careers in marine mammal medicine Encephalitis etiology unclear, 537 in manatees, 952 in pinnipeds, 368, 921, 922 Encephalomyelitis, 429 Endangered Species Act (ESA) about, 742, 743–744, 760 CITES implementation, 746, 750 consultations, 745 enforcement, 745–746 listing procedure, 744 manatee protected by, 41 permits, 745 species protection provision, 744–745 species status, 746–749 Endemic fungi infections, 338 Endocrine disruptor hypothesis, 477 Endocrine system contaminants’ effects on, 474–475 halogenated organics effects on, 490–491 non-infectious diseases, 535 pinniped diseases, 920
CRC Handbook of Marine Mammal Medicine
polar bears, 992–993 Endocrinology adrenal glands aldosterone, 181–182 catecholamine function and physiology, 178 cortisol levels, 178–181 described, 177 endocrine pancreas, 185–186 hypothalamus-pituitary, 169 osmoregulatory hormones atrial natriuretic peptide, 185 renin-antiotensin system, 185 vasopressin, 183–184 pineal gland, 167–168 sample collection and handling, 166–167 stress response components catecholamines, 256 glucocorticoids, 256, 259–260 mineralocorticoids, 260 other hormones, 261 stress indicators, 257–258 thyroid hormones, 260–261 thyroid gland circulating concentrations of thyroxine, 171–172 concentrations of fT4 175–176 fluctuations in TH levels, 176–177 hormone storage, 170 protein binding, 170 reverse T3, 174, 176 thyroxine levels, 170, 174 triiodothyronine concentrations, 173–174 Endoscopy clinical applications, cetaceans abdominal imaging, 562 colonoscopy, 633 gastroscopy, 630–633 respiratory imaging, 633–635 urogenital imaging, 635 clinical applications, other marine mammals, 635–636 equipment accessories and instruments, 627–629 cameras, 629–630 flexible endoscopes, 624–626 light sources, 626–627 rigid telescopes, 626 video monitors/recorders, 630 indications, 622–623 limitations, 623–624 minimally invasive surgical techniques access, 637–638 cannulas, 638–639 in cetaceans, 640 closure, 636–640 insufflation, 636–637 in other marine mammals, 640 trocars, 638–639 overview, 621–622
0839_frame_IDX Page 1033 Friday, May 25, 2001 9:37 AM
1033
Index
Endosulfan, 485 Endrin, 484 Energy requirements, see Bioenergetic scheme Enflurane, 734 Enhancement permits, 760 Enhydra lutris, see California sea otter Enrofloxacin cetacean drug dosage, 711 pinniped drug dosage, 715 sea otter drug dosage, 719 Entamoeba, 360 Enteritides, necrotic, 1001 Enterocolitis, 428–429 Environmental contamination, see Water and environment quality Environmental diseases in manatees brevetoxicosis, 951 cold stress syndrome, 951–952 Enzyme-linked immunosorbent assay (ELISA), 245 Eosinophils, 401–402 EPA (U.S. Environmental Protection Agency), 785 Epinephrine cetacean drug dosage, 711 dive response and, 178 for otariids, 670 Epizootics, 16, 491–493 Epstein-Barr virus, 21 Equipment blood collection, 384–385 endoscopy accessories and instruments, 627–629 cameras, 629–630 flexible endoscopes, 624–626 light sources, 626–627 rigid telescopes, 626 video monitors/recorders, 630 sonography, 594–595 tagging and tracking, see Tagging and tracking Erignathus barbatus (bearded seal) parasites of, 368 viral disease in, 293 EROD (ethyoxyresorufin-o deethylase), 489 Erysipelothrix cetacean infection case study, 425–426 in cetaceans, 316–318 diagnosis, 316 in pinnipeds, 318–319 public health significance, 771 septicemia and, 312 Erythrocytes cytology interpretation, 442 evaluation anemia, 399–401 indices, 391–393, 396, 399 in respiratory tract, 443 sedimentation rate immunodiagnostics and, 240–241 inflammatory disease indication, 422, 902
in stomach, 444 in urine, 446 Erythromycin, 715 Erythropoietin, 711 ESA, see Endangered Species Act Escherichia coli, 327 Eschrichtius robustus (California gray whale) caliciviruses in, 300 fasting during lactation, 803 hematology and biochemistry values, 393 ESR (erythrocyte sedimentation rate) immunodiagnostics and, 240–241 inflammatory disease indication, 422, 902 Estradiol-2-hydroxylase (E2OH), 489 Estrogen photoperiod changes in pinnipeds, 198 reproduction regulation role, 194, 197 Estrous cycle false killer whales, 208–209 follicular maturation and, 197 in harbor seals, 149 in pinnipeds, 196–197 Ethoxycoumarin-o-deethylase (ECOD), 489 Ethyoxyresorufin-o-deethylase (EROD), 489 Etorphine euthanasia use, 732–733 for polar bears, 999 Eumetopias jubatus, see Steller sea lion European Association for Aquatic Mammals (EAAM), 905 European harbor seal, 293, 294, 296; see also Harbor seal Euthanasia carcass disposal, 736–737 criteria for stranded animals, 729–730 display and collection animals, 730 humaneness criteria for methods, 730–731 inhalants, 734 injectable agents barbiturates, 732 etorphine, 732–733 paralytics, 733 route of administration, 731–732 T-61, 733 physical methods ballistics, 734–736 explosives, 736 as stranding response, 94 verification of death, 736 Evidence-based medicine Web site, 120 Explosives use for euthanasia, 736 Exxon Valdez oil spill, 69, 264, 496–497 Eyes, see Ophthalmology
F Factor XII (Hageman factor) activity, 421 False killer whale (Pseudorca crassidens) bacterial diseases in, 322
0839_frame_IDX Page 1034 Friday, May 25, 2001 10:00 AM
1034
cytology, see Cytology, cetacean estrous cycle, 208–209 hematology and biochemistry values, 392 non-infectious diseases, 534 ovarian physiology, 208–209 parasites of, 359 pod cohesion factor in strandings, 86 pregnancy diagnosis, 212 pseudopregnancy, 210 reproductive cycle, 206 reproductive maturity, 205 reproductive parameters, 213–214 seasonality of fertility, 217 serum analytes and enzymes, 405 Fasting and starvation, 801–803 FDA (Food and Drug Administration), 118 Feces sample collection and handling, 166 Fecundity rates for bottlenose dolphin, 205 Federal legislation and regulations Animal Welfare Act (AWA), see Animal Welfare Act definitions and abbreviations, 759–762 discussion, 741–742 Endangered Species Act (ESA) about, 743–744 CITES implementation, 746, 750 consultations, 745 enforcement, 745–746 listing procedure, 744 permits, 745 species protection provision, 744–745 species status, 746–749 Fur Seal Act (FSA), 758 Lacey Act (LA), 758 Marine Mammal Protection Act (MMPA) incidental take exemptions and permits, 750, 752–753 moratorium on taking, 750 reauthorizations of, 753 species included, 750 strandings and health, 753–754 permits, frequently asked questions (FAQs) for observations, 765 public display, 764–765 research and enhancement, 763–764, 765 stranding networks, 762–763 regulating agencies, 742–743 Federal Register, 742 Fellowships Web sites, 112 Fenbendazole drug dosage cetaceans, 711 pinnipeds, 715 polar bears, 721 sirenians, 718 for parasite treatment, 359 in cetaceans, 362, 366 in pinnipeds, 369, 370, 371 Fentanyl, 683
CRC Handbook of Marine Mammal Medicine
Fentanyl citrate, 999 Feral and domestic cat contamination, 24 Fibrin, 443 Fibrous nodules, 536 Filariidae, 370 Filaroididae, 370 Filtration methods for pool waters biological, 781–782 flocculation, 782 foam fractionation, 783 mechanical, 782 Finless porpoise (Neophocoena phocoenoides), 325–326 Fin whale (Balaenoptera physalus) age estimation, 455–456 bacterial diseases in, 314 reproduction energy requirements, 805 viral diseases in, 300 Fish and Wildlife Service (FWS), see U.S. Fish and Wildlife Service Fishing line dangers, 957–958 Flagellates in cetaceans, 360 in pinnipeds, 368 Fletcher Factor activity (plasma prekallikrein), 421 Flexible endoscopes, 624–626 Flipper tags for pinnipeds, 858 Flocculation filtration for pool waters, 782 Florfenicol adverse effects, 707 cetacean drug dosage, 711 Florida manatee (Trichechus manatus latirostris), see also Manatee boat related maimings animal rights considerations, 35 database of scarred individuals, 33 laws pertaining to, 36 multiple cuts evidence, 33–35 boat related mortalities, 31–33 carcass counts, 39 external features, 138 gross anatomy illustration, 132–133 mortality computation, 39–40 population models, 40 population size estimation, 36–38 population trends use, 40–41 trend analysis, 38–39 Florida Manatee Sanctuary Act (1978), 41 Florinef, 708 Fluconazole adverse effects, 708 drug dosage cetaceans, 711 pinnipeds, 715 mycotic infection use, 349, 350 Flucytosine adverse effects, 708 cetacean drug dosage, 711 for mycotic infections, 350, 351
0839_frame_IDX Page 1035 Friday, May 25, 2001 9:37 AM
Index
Flumazenil, 670 Flumethasone, 721 Flunixin meglumine contraindications, 707 intensive care use, 699 Fluoroquinolones, 706 Fluoxetine, 994 Fluoxitine HCL, 715 Foam fractionation filtration for pool waters, 783 Folic acid cetacean drug dosage, 711 sea otter drug dosage, 719 Follicle-stimulating hormone (FSH) estrous cycle and, 196 reproduction regulation role, 194 stress response, 263 Follicular maturation and estrous cycle, 197 Food and Drug Administration (FDA), 118 Forceps, 628 Foreign material retrieval, 627 Forestomach evaluation by sonography, 606 Formularies, see Pharmaceuticals and formularies Foundation Directory, 123 Fractures, 532 French stranding networks contacts and programs, 54–55 FSA (Fur Seal Act (1966)), 760 FSH, see Follicle-stimulating hormone Fundic stomach evaluation by sonography, 606 Fungi diseases caused by, see Mycotic diseases public health significance of infections, 773 Fungi Imperfecti, 337 Furosemide adverse effects, 708–709 drug dosage cetaceans, 711 pinnipeds, 716 sea otters, 719 Fur seal (Arctocephalus spp.) bacterial diseases in, 320 brucellosis, 313 gastrointestinal, 327–328 blood transfusions, 692 fasting during lactation, 803 thermoneutral zone, 795 viral diseases in, 296 Fur Seal Act (FSA), 758, 760 Fusarium spp. lesions, diagnostic methods, and treatments, 344 mechanisms of pathogenesis, 339 FWS, see U.S. Fish and Wildlife Service
G Galapagos sea lion (Zalophus californianus wollebaeki) bacterial diseases in, 326 lactation energy requirements, 806
1035
Gastric nematodes in pinnipeds, 916 Gastrointestinal system of cetaceans, 146 cytology findings, cetaceans, 444 diseases helminth parasites, cetaceans, 361–362 helminth parasites, pinnipeds, 369–370 in pinnipeds and cetaceans, 327–328 fecal and urinary energy losses, 809–810 gastroenteritis, sea otters, 978 gross necropsy investigation, 454–455 intestines digestive system description, 145–146 intestinal erosions and obstructions, 533–534 intestinal volvulus, 534 pinniped diseases, 916–917 sonography clinical applications, 601, 605–608 walrus diseases, 934–935 Genetic analysis captive mammals applications hybrid detection, 276–277 paternity testing, 275–276 in necropsy, 453–454 sampling, 277–278 stranded mammals applications population identification, 273–274 social organization, 274–275 species identification, 273 techniques DNA fingerprinting, 272 DNA sequencing, 271–272 RFLP, 271 tandem repeats, 272 Genital tract gross anatomy, 147–148 microscopic anatomy, 149–150 non-infectious diseases, 534–535 Gentamicin cetacean drug dosage, 711 pinniped drug dosage, 716 sea otter drug dosage, 720 sirenian drug dosage, 718 Geomagnetic disturbances as explanation for live strandings, 86 German stranding networks contacts and programs, 54 GGT (γ-glutamyltransferase), 408–409 GH (growth hormone), 169 Giardia spp. anthropogenic origin, 5 in pinnipeds, 22, 368 public health significance, 774 Gingivitis, 532 Girella nigricans (opaleye perch), 301 GLDH (glutamate dehydrogenase), 407–408 Global Positioning System (GPS), 857 Globicephala spp. (pilot whale), see Pilot whale Globulins, 414–415 Glomerular disease, 534
0839_frame_IDX Page 1036 Friday, May 25, 2001 9:37 AM
1036
Glucagon, 186 Glucocorticoids adrenal gland studies and, 178 stress response components, 256, 259–260 stress response role, 180–181 Glucose, serum analytes and enzymes, 403–404 Glutamate dehydrogenase (GLDH), 407–408 γ-Glutamyltransferase (GGT), 408–409 Glutathione-S-transferase (GSH-T), 489–490 Gonadotropin-releasing hormone (GnRH) contraception for pinnipeds and, 202–203 estrous cycle and, 196 reproduction regulation role, 194 stress response, 263 Gonyaulax spp., 493, 494 Gonyautoxin-1, 21, 494 Goosebeak, 144 GPS (Global Positioning System), 857 Graded histopathological findings of stranded cetaceans, 93 Graduate degree programs Web sites, 109–110 Grampus griseus, see Risso’s dolphins Grantsnet Web site, 123 Granulomata, 535 Granulomatous infections, see Nocardia Granulomatous nodules, 536 Grateful Med Web site, 118, 119 Gray seal (Halichoerus grypus) bacterial diseases in, 22 Brucella, 314 respiratory, 325 contaminants’ effects on organ systems, 472, 473, 474 melatonin concentrations in, 168 mercury concentration in liver, 478 nutrition, 813 organohalogens accumulation in, 488 parasites of, 368 reproduction energy requirements, 805 social organization, 274 thermoneutral zone, 794 TH fluctuations and, 177 viral diseases in herpesviruses, 292, 293, 294 morbillivirus, 296 poxvirus, 287, 288 Gray whale (Eschrichtius robustus) caliciviruses in, 300 fasting during lactation, 803 hematology and biochemistry values, 393 Great sperm whale (Physeter catadon), 6–7; see also Sperm whale Greecian stranding networks contacts and programs, 55 Greenhouse gas effect, 5 Griseofulvin pinniped drug dosage, 716 sea otter drug dosage, 720
CRC Handbook of Marine Mammal Medicine
Gross anatomy adrenal glands, 148 caval sphincter, 144 circulatory structures, 151–152 diaphragm, 142 digestive system, 145–147 external features, 138–139 genital tract, 147–148 heart, 142 illustrations bottlenose dolphin, 136–137 California sea lion, 130–131 Florida manatee, 132–133 harbor seal, 134–135 larynx, 144 liver, 145 lungs, 143 mediastinum, 143 parathyroids, 144 pericardium, 142 pleura, 143 skeletal muscles, superficial, 141–142 skeleton bone marrow, 158 chevron bones, 156 limb complexes, 156–157 locomotion, 154 overview, 153–154 ribs, 155 sexual dimorphisms, 157 sternum, 155–156 vertebrae, 154–155, 156 thermal insult protection, reproductive organs, 152–153 thymus, 143 thyroids, 143 urinary tract, 147 Gross necropsy acoustic pathology, 459–460 age estimation, 454–455 anthropogenic causes of death, 458, 460 biotoxins specimen collection, 468 carcass condition classification, 451 carcass condition code, 453 carcass examination, 457–458 chain of custody form, 452 chemical pollutants speciman collection, 466 examinations and specimen collection, 450 genetic analysis, 453–454 harmful algal blooms, 465, 467 histopatholgy, 458–459 infectious diseases, 460, 462, 463 life-history data protocols, 461 morphometrics, 453 non-infectious diseases, 464–465 overview, 449–450 parasitology, 462–463 reproductive status, 456–457
0839_frame_IDX Page 1037 Friday, May 25, 2001 9:37 AM
Index
stomach contents protocol, 454 virology, 462 Growth hormone (GH), 169 GSH-T (glutathione-S-transferase), 489–490 Guadalupe fur seal (Arctocephalus townsendi), 692 Gum recession, 532 Gunshot cases, 458 Gymnodinium breve, 5, 493, 951
H Haematophagus megapterae, 359 Hageman factor (Factor XII) activity, 421 Halarachne spp. in pinnipeds, 372 in sea otters, 978 Halichoerus grypus, see Gray seal Halocercus, 443 Halogenated organics, see also Contaminants accumulation and variability, 482–484 octachlorostyrene, 487 organochlorine pesticides and metabolites, 484–485 organochlorines effects on metabolism, 488–490 PBDEs, 487 PCDDs, 486 PCDFs, 486 PCTs, 487 polybrominated biphenyls, 487 polychlorinated byphenyls, 485–486 polychlorinated napthalenes, 486 polychloroquaterphenyls, 486 TCPMe, 486 TCPMeOH, 486 Haloperidol, 716 Halothane for cetaceans, 660 euthanasia use, 734 for otariids, 667 Hand-rearing and artificial milk formulas cetaceans, 829–831 manatees, 843–845 pinnipeds elephant seals, 836–838 harbor seals, 832–834 sea lions, 839–840 walruses, 841–842 polar bears, 847–848 sea otters, 845–846 Haptoglobins (Hp), 264 Harassment definition, 761 Harbor porpoise (Phocoena phocoena) bacterial disease in, 313–314 contaminants’ effects on organ systems, 475 epizootic cause, 16 non-infectious diseases endocrine system, 535 genitourinary system, 535 musculoskeletal system, 532
1037
organochlorine reproductive function effects, 492 thermoneutral zone, 795 viral diseases in herpesviruses, 292, 294 morbillivirus, 296 papillomavirus, 19, 289–290, 291 water requirements, 800 Harbor seal (Phoca vitulina) bacterial diseases in Brucella, 22, 314 dermatological, 326 gastrointestinal, 327 Leptospira, 770 leptospirosis, 320 respiratory, 325 urogenital, 327 clinical pathology coronavirus case, 427–428 hematology and biochemistry values, 394–395 contaminants’ effects on organ systems, 474, 475 contraception, 203 cortisol levels, 178 dehydration indications, 690 estrous cycle, 195 external features, 139 gross anatomy illustration, 134–135 hand-rearing and artificial milk formulas, 832–834 heat increment of feeding, 808 hepatobiliary system disorders markers, 410 immunodiagnostics of, 241 lactation energy requirements, 805, 806 liver, 148 locomotion metabolic expenditure, 796 mercury concentration in liver, 478 non-infectious diseases endocrine system, 535 genitourinary system, 534 integumentary system, 532 organochlorine effects on, 490, 491 parasites of, 367, 370, 372 parental similarity vs. fitness and, 275 population structure determination, 274 pregnancy determination, 199 reproduction energy requirements, 805 reproductive characteristics, 196 sentinel role, 6 species identification techniques, 273 spinal cord, 151 stress response, 260 thermoneutral zone, 794 TH fluctuations and, 177 thyroid gland, 176 thyroids, 145 viral diseases in hepadnavirus, 302 herpesviruses, 292, 293 influenza, 298, 299 poxvirus, 286, 287, 288
0839_frame_IDX Page 1038 Friday, May 25, 2001 9:37 AM
1038
water requirements, 800 Harm definition, 761 Harmful algal blooms, 465, 467; see also Red tides Harpacticus pulex, 953 Harp seal (Pagophilus groenlandicus) bacterial diseases in, 22, 314 contaminants’ effects on organ systems, 472 heat increment of feeding, 808 integument microanatomy, 140 melatonin concentrations in, 168 milk fat content, 807 nutrition thiamine deficiency, 813 vitamin deficiencies, 815 organochlorine effects on, 490 parasites of, 368 reproduction energy requirements, 805 stress response, 260 thermoneutral zone, 794 TH fluctuations and, 177 thyroid gland, 176 viral diseases in herpesviruses, 293 morbillivirus, 296, 298 Haul-out area size for pinnipeds, 907 Hawaiian monk seal (Monachus schauinslandi) biotoxins effects on, 496 contraception, 203 estrous cycle, 196, 197 non-infectious diseases, 535 viral diseases in, 300 Hb (blood hemoglobin content), see Erythrocytes, evaluation HCB (hexachlorobenzene), 485 HCH, 485 HCT (hematocrit), 413; see also Erythrocytes, evaluation Health on the Net (HON), 119–120 Heart, see Cardiovascular system Heartworm infestations in pinnipeds, 919 in walruses, 935 Heat increment of feeding (HIF), 807–808 Hector’s dolphin (Cephalorhynchus hectori), 286 Helicobacter, 19 Helminths, see also Cestodes; Nematodes; Trematodes in manatees, 373 in pinnipeds, 369–372 in polar bears, 374 in sea otters, 373–374 Hematocrit (HCT), 413; see also Erythrocytes, evaluation Hematology, see also Blood overview, 390 references, 397–399 values per species, 391–396 Hematopoietic system, 150 Hemorrhagic encephalopathy, 536
CRC Handbook of Marine Mammal Medicine
Hemostatic parameters, see also Blood blood types, 420 disorders screening, 420–421 prothrombin time, 421–422 Hepadnavirus, 302 Hepadnavirus hepatitis, 426–427 Hepatic cirrhosis, 408 Hepatic lipidosis, 534 Hepatitis, 534, 916 Hepatobiliary system disorders markers γ-glutamyltransferase, 408–409 alanine aminotransferase, 406 alkaline phosphatase, 409–410 aspartate aminotransferase, 407 bile acids, 411 bilirubin, 410–411 glutamate dehydrogenase, 407–408 lactate dehydrogenase, 408 sorbitol dehydrogenase, 407–408 Heptachlor epoxide, 484, 485 Herpesviruses clinical signs, 293–294 diagnosis, 294–295 differentials, 295 epidemiology, 295 host range, 292–293 pathology, 294 in pinnipeds, 913, 920 public health significance, 295 in sea otters, 979 therapy, 294 virology, 293 Herpesvirus-like particles observations of, 21–22 serodiagnostics of, 244 Hetacillin, 720 Heterophils and leukocytes evaluation, 401 Hexachlorobenzene (HCB), 485 Hexamita, 360 Hibernation induction trigger (HIT), 990 HIF (heat increment of feeding), 807–808 Histopatholgy, 458–459 Histoplasma capsulatum lesions, diagnostic methods, and treatments, 348 mechanisms of pathogenesis, 338 Histriophoca fasciata (ribbon seal), 293 HIT (hibernation induction trigger), 990 HON (Health on the Net), 119–120 Hong Kong stranding networks contacts and programs, 55–56 Hooded seal (Cystophora cristata) bacterial diseases in, 22, 314 organochlorine effects on, 490 viral diseases in herpesviruses, 293 morbillivirus, 296 Hookworms (Ancylostomatidae), 369, 919
0839_frame_IDX Page 1039 Friday, May 25, 2001 9:37 AM
1039
Index
Hormones hypothalamus-pituitary and, 169 in polar bears, 992–993 reproduction regulation role, 194 Hp (haptoglobins), 264 Huckins, Olga Owens, 3 Human chorionic gonadotropin, 712 Human contraction of diseases caliciviruses, 302 Erysipelothrix, 316 herpesviruses, 295 influenza, 300 Nocardia, 322 papillomavirus, 291 poxvirus, 289 Humpback whale (Megaptera novaeangliae) biotoxins effects on, 494 parasites of, 359 population structure determination, 274 stranding, 69 Hybrid determination, 276–277 Hydration status, 690 Hydrocephalus in pinnipeds, 921 Hydrocortisone, 720 Hydrogen peroxide, 716 Hydrurga leptonyx, see Leopard seal Hyperadrenocorticisim, 535 Hyperglobulinemia, 416 Hyperkalemia, 416 Hyperkeratosis, 531 Hypernatremia, 416 Hyperthermia after anesthesia administration otariids, 669 phocids, 676 protecting against, 908 sea otters, 974–975 Hypochlorus acid, 785 Hypoglycemia sea otters, 974 therapy for, 693 Hypomagnesemia, 416 Hypomineralization of teeth, 532 Hyponatremia described, 814–815 in phocids, 416 in pinnipeds, 922 Hypothalamus-pituitary endocrinology, 169 reproduction regulation role, 193–194 Hypothermia after anesthesia administration odobenids, 680 otariids, 669 phocids, 675–676 protecting against, 908 sea otters, 975 Hypothyroidism in pinnipeds, 920
Hypoventilation, 676 Hypovitaminosis A in polar bears, 1003 Hypoxia, 680
I IAAAM (International Association for Aquatic Animal Medicine), 905 IATA (International Air Transport Association), 881 ICH (infectious canine hepatitis), 291 Icterus, 410 Imaging software, 557 Imipenem, 712 Immobilon, 732, 733 Immune system adaptive response, 238–239 clinical evaluation of disorders abnormality determination, 246–248 classification scheme, 246 evaluation flowchart, 248 immunodeficiency syndromes identification, 246, 247 contaminants’ effects on, 475–476 cytokines, 239–240 immunodiagnostics cellular immunity, 241–242 functional immune testing, 242–243 humoral immunity, 243 inflammation, 240–241 immunosuppression and organochlorines, 491 inflammatory response, 238 innate immunity, 238 serodiagnostics enzyme-linked immunosorbent assay, 245 importance of controls, 243–244 precipitation/agglutination techniques, 244–245 serum/virus neutralization test, 244 total immunoglobulin, 245 Immunocompetence effects of halogenated organics, 491–493 Immunoglobulins, 243 Immunology, see Immune system Implantable tags for cetaceans, 866 Index methods for population trends estimation, 37 Index surveys of manatees, 38 Indomethacin, 716 Infectious canine hepatitis (ICH), 291 Infectious diseases, see also Bacterial diseases and infections; Viral diseases and infections gross necropsy, 460, 462 manatees, 952–953 in polar bears, 1001 Inflammation markers erythrocyte sedimentation rate, 422 serum iron, 422–423 Influenza described, 298–300 in pinnipeds, 918
0839_frame_IDX Page 1040 Friday, May 25, 2001 9:37 AM
1040
public health significance, 300, 769 Infrared technology, see Thermal imaging Ingested energy, 803–804 Inhalation anesthesia cetaceans, 660 odobenids, 680 otariids, 667 phocids, 675 polar bears, 999–1000 sea otters, 683 Inhibin, 194 Inia geoffrensis (Amazon river dolphin) non-infectious diseases, 532 water requirements, 799–800 Injectable agents for euthanasia barbiturates, 732 etorphine, 732–733 paralytics, 733 route of administration, 731–732 T-61, 733 Insemination techniques for cetacean reproduction, 223–224 Insufflation, 636–637 Insulin, 186, 720 Integument contaminants’ effects on, 472 microanatomy, 139–140 non-infectious diseases, 531–532 pinniped diseases, 913–915 polar bear diseases, 1002–1003 Intensive care analgesics, 699 antibiotics, 698–699 blood transfusion, 692 central nervous system, 696 miscellaneous therapeutic agents, 699 nutritional therapy appetite stimulants, 694 emaciation, 693–694 hypoglycemia, 693 patient evaluation, 689–690 records and instructions, 689 rehydration, 690–692 reproductive emergencies, 697–698 respiratory emergencies, 695 support equipment, 699–700 trauma, 695–696 wound management, 696 International Air Transport Association (IATA), 881 International Association for Aquatic Animal Medicine (IAAAM), 905 International Biological Programme, 803 Internet use for information gathering, see Electronic information gathering Internships and residencies at aquaria or rehabilitation centers, 108–109 at institutions or agencies, 109 matched internships, 107–108
CRC Handbook of Marine Mammal Medicine
matched residencies, 108 at zoos, 109 Interviews for jobs, preparation advice, 103–106 Intestines, small and large, see also Gastrointestinal system digestive system description, 145–146 intestinal erosions and obstructions, 533–534 intestinal volvulus, 534 Intra-abdominal radio transmitters, 871–873 Intracellular hemosiderin, 533 Isoetharine, 716 Isoflurane for cetaceans, 660 euthanasia use, 734 for otariids, 667, 669 Isoniazid, 716 Isoproterenol pinniped drug dosage, 716 sea otter drug dosage, 720 Isreali stranding networks contacts and programs, 56 Isuprel, 716 Italian stranding networks contacts and programs, 56–57 Itraconazole adverse effects, 708 drug dosage cetaceans, 712 pinnipeds, 716 sirenians, 718 mycotic infection use, 349, 350, 351 Ivermectin adverse effects, 708 drug dosage cetaceans, 712 pinnipeds, 716 polar bears, 721 sea otters, 720 sirenians, 718 for parasite treatment, 359, 371 cetaceans, 362, 366 pinnipeds, 370
J Japanese stranding networks contacts and programs, 57–58 Job search methods action plan, 102 communication skills, 106–107 important attributes of applicants, 106 interview preparation, 103–106 jobs locations research, 102–103 self-assessment, 99–100 skill categorization, 100–101 Jobs in marine mammal medicine, see Careers in marine mammal medicine Journals and textbooks, online, 121–122
0839_frame_IDX Page 1041 Friday, May 25, 2001 9:37 AM
1041
Index
K Kaopectate, 720 Keloidal blastomycosis, 773 Kepone, 485 Ketamine for otariids, 666, 669 for phocids, 674 for polar bears, 997–998 for sea otters, 683 Ketamine/xylazine, 998 Ketoconazole adverse effects, 708 drug dosage cetaceans, 712 pinnipeds, 716 sea otters, 720 mycotic infection use, 350, 351 Kidney-associated serum analytes creatinine, 412 urea nitrogen, 411–412 Kidneys contaminants’ effects on, 472–473 described, 147 lead concentrations in, 481 reniculate, 147, 149 sonography clinical applications, 609–611 Killer whale (Orcinus orca) bacterial diseases in Brucella, 313–314 Nocardia, 322 clinical pathology hematology and biochemistry values, 392 infection cases, 425–426 cortisol levels, 178 cytology, see Cytology, cetacean estrous cycle, 208 hyponatremia care, 696 lactational suppression of estrus, 209 leukocytes evaluation, 403 non-infectious diseases cardiovascular system, 535 musculoskeletal system, 532 ovarian physiology, 208 parasites of, 359 PCB toxicity and, 486 population structure determination, 274 pregnancy, 211 pregnancy diagnosis, 212 pseudopregnancy, 210 reproductive cycle, 206 reproductive maturity, 204 reproductive parameters, 213–214 seasonality of fertility, 217 sexual maturity, 215–216 thermal imaging applications, 647 viral diseases in papillomavirus, 19, 290
poxvirus, 286 Klebsiella, 312 Kyaroikeus cetarius cytology findings in respiratory tract, 443 species range, 359
L Lacazia loboi epidemiology, 338 lesions, diagnostic methods, and treatments, 348 mechanisms of pathogenesis, 339 Lacey Act (1901), 758, 761 Lactate dehydrogenase (LDH), 408 Lactated Ringer’s, 720 Lactation and nursing energy requirements, 805–807 estrus suppression and, 209 manatees, 942 milk fat content of polar bears, 991–992 milk fat content of sea otters, 807 pinnipeds, 200 sea otters, 966 Lagenorhynchus acutus, see Atlantic white-sided dolphin Lagenorhynchus albirostris (Atlantic white-beaked dolphin) bacterial diseases in, 317 viral diseases in morbillivirus, 296 rhabdoviruses, 303 Lagenorhynchus obliquidens, see Pacific white-sided dolphin Lagenorhynchus obscurus, see Dusky dolphin Laparoscopy, 621 Large roundworms (Anisakidae), 361–362, 369 Larynx, 144 Law, animal, see Federal legislation and regulations LDH (lactate dehydrogenase), 408 Lead, 481 Legislation, federal, see Federal legislation and regulations Leopard seal (Hydrurga leptonyx) bacterial diseases in, 325 parasites of, 368 viral diseases in, 296 Leptonychotes weddellii, see Weddell seal Leptospira spp. in pinnipeds, 320–321, 919 public health significance, 770 in walruses, 935 Leptospires, 920 Leptospirosis in pinnipeds, 320–321, 919 in polar bears, 1001 reproductive impact, 490 Leukocytes evaluation age and, 402 basophils, 402
0839_frame_IDX Page 1042 Friday, May 25, 2001 9:37 AM
1042
counts in colon/rectum, 445 cytology interpretation, 442 in respiratory tract, 443 in stomach, 444 stress recognition tool, 261 in urine, 446 disease and, 403 eosinophils, 401–402 heterophils, 401 inflammatory disease indication, 902–903 lymphocytes, 402 monocytes, 402 neutrophils, 401 Leuprolide acetate for dolphin contraception, 218 drug dosage cetaceans, 712 pinnipeds, 716 sea otters, 720 Levamisole adverse effects, 708 drug dosage pinnipeds, 716 sea otters, 720 LH, see Lutenizing hormone Lice (Anoplura sp.), 372, 915 Lidocaine HCL cetacean drug dosage, 712 for cetaceans, 657 sea otter drug dosage, 720 Lighting conditions for captive animals, 781 Light sources in endoscopy, 626–627 Limb complexes, 156–157 Lincomycin, 720 Lindane, 485 Lipase, 405 Lipids organohalogens accumulation in, 483–484 PCB accumulation in, 486 serum analytes and enzymes, 404 Lissodelphis borealis (northern right whale dolphin), 359 Listing by cetaceans, 898 Listservs, marine mammal related, 120–121 Liver contaminants’ effects on, 472–473 function, 146 gross anatomy, 145 helminth parasites in cetaceans, 365 helminth parasites in pinnipeds, 372 hepatobiliary system disorders markers γ-glutamyltransferase, 408–409 alanine aminotransferase, 406 alkaline phosphatase, 409–410 aspartate aminotransferase, 407 bile acids, 411 bilirubin, 410–411 glutamate dehydrogenase, 407–408
CRC Handbook of Marine Mammal Medicine
lactate dehydrogenase, 408 sorbitol dehydrogenase, 407–408 lead concentrations in, 481 mercury concentration in, 478–480 microscopic anatomy, 148 sonography clinical applications, 601–604 Lobodon carcinophagus (crabeater seal), 296 Lobomycosis, 339 Lobo’s disease, 773 Locomotion metabolic rate and, 796–799 skeleton and, 154 walruses, 928 Lontra canadensis (river otter), 19 Lordosis, 532 LSD (lysergic acid diethylamide), 498 Lungs, see also Respiratory system gross anatomy, 143 sonography clinical applications, 597–600 Lungworm (Parafilaroides decorus) bacterial diseases and, 314 infestations in cetaceans, 365 in pinnipeds, 370 viral disease and, 301 Lutenizing hormone (LH) in cetaceans, 169 estrous cycle and, 196 reproduction regulation role, 194 stress response, 263 Lymphocytes, 402–403 Lymphoid system microscopic anatomy, 150 non-infectious diseases, 536 Lyris Web site, 121 Lysergic acid diethylamide (LSD), 498
M Macrophages in stomach, 444 Magnesium, 419 Magnetic resonance imaging (MRI) anatomy, dolphin, 587–589 described, 560 indications for use, 563 limitations, 568 Maintenance energy, 804 Major histocompatibility proteins (MHC), 239 Malassezia pachydermatis, 344 Maldivian stranding networks contacts and programs, 58 Maltan stranding networks contacts and programs, 58–59 Mammary glands and integument microanatomy, 140; see also Lactation and nursing Manatee (Trichechus manatus), see also Florida manatee age estimation, 455–456 aldosterone levels, 181 analgesics use, 699
0839_frame_IDX Page 1043 Friday, May 25, 2001 9:37 AM
Index
anatomy, 941 antibiotics use, 698 behavior, 942 biotoxins effects on, 493–494 boat trauma treatment, see Boat related maimings circulatory structures, 151 clinical pathology blood collection, 387 encephalomyelitis case, 429 enterocolitis case, 428–429 hematology and biochemistry values, 396 references, 398 contaminants’ effects on organ systems, 472, 473, 475 diaphragm, 142 digestive efficiencies, 810 digestive system, 146 emerging and resurging diseases in, 22–23 entanglement in fishing line, 957–958 environmental diseases brevetoxicosis, 951 cold stress syndrome, 951–952 external features, 138–139 FWS jurisdiction, 742 hand-rearing and artificial milk formulas, 843–845 hepatobiliary system disorders markers, 406, 407, 411 husbandry anesthesia, 950 diagnostic techniques, 946–948 habitat requirements, 942 nutrition, 943–944 physical examination, 946 restraint, handling, and transport, 944–945 therapeutics, 948–950 water requirements, 942–943 infectious diseases, 952–953 lactation energy requirements, 806 leukocytes evaluation, 401–403 liver, 145 mediastinum, 143 metabolic rate, 793 natural history, 939–941 neonatal disease, 953–954 neoplasia, 953 nutritional therapy, 693, 694 osmoregulatory hormones, 184 parasites of, 372–373 pectoral limb complex, 157 physiology, 941–942 population structure determination, 274 rehydration, 691 reproductive emergencies, 697 respiratory emergencies, 695 salinity and pH requirements, 781 serum analytes and enzymes, 405 species status under ESA, 749 spinal cord, 151
1043
tagging and tracking branding, 867 radio telemetry use and attachment, 867–868 tagging advantages, 868, 870 thermal imaging applications, 646 thermal insult protection during diving, 153 thermoneutral zone, 796 thyroxine levels, 170 urinary tract, 147 vertebrae, 156 viral diseases in, 296 water requirements, 799–801 MAP (mean arterial blood pressure), 661 Marine Mammal Commission (MMC), 743, 761 Marine Mammal Health and Stranding Response Act, 69–70, 753–754 Marine Mammal Health and Stranding Response Program, 450 Marine Mammal Net, 121 Marine Mammal Pathobiology Laboratory, 954 Marine Mammal Protection Act (MMPA) (1972) about, 742, 761 hunting regulations, sea otters, 967 incidental take exemptions and permits, 750, 752–753 manatee protection from, 41 marine mammal collections and, 7–8 moratorium on taking, 750 reauthorizations of, 753 species included, 750 strandings and health, 753–754 Marine Mammal Unusual Mortality Event (MMUME) contingency plan need, 69–70 cooperative response need, 77–78 definition, 70–71 events since 1992, 72–73 help avenues for public, 79 responses in U.S. elements of response, 74 expert working group, 71, 74 funding, 76–77, 78 national contingency plan, 71 process steps, 74–76 Title IV results, 78 MarMam Web site, 120 Marshall Space Flight Center, NASA, 857 Mass strandings, cetaceans current investigations into, 86–87 definition, 83 disposition of animals, 92, 94 east coast of U.S. (1987–1999), 84–85 evaluation of, 87 graded histopathological findings, 93 management of, 88–92 serum chemistry findings, 91 theories to explain, 83, 85–86 treatment precautions, 94 Maternal recognition of pregnancy (MRP), 197
0839_frame_IDX Page 1044 Friday, May 25, 2001 9:37 AM
1044
Mean arterial blood pressure (MAP), 661 Measles virus (MV), 296 Mebendazole, 721 Mechanical filtration methods for pool waters, 782 Medetomidine, 670 Medetomidine-ketamine, 666, 669 Mediastinum gross anatomy, 143 sonography clinical applications, 597 Medline Web site, 118, 119 Medroxyprogesterone acetate, 720 Megaptera novaeangliae, see Humpback whale Megestrol acetate cetacean drug dosage, 712 for nutritional therapy, 694 pinniped drug dosage, 716 Melatonin pineal gland and, 167–168 reproductive cycle and, 195 Meperidine cetacean drug dosage, 712 for cetaceans, 657 intensive care use, 699 for odobenids, 677 for sea otters, 683 Mercury concentrations in livers, 478 gross distribution within livers, 479 tolerance of marine mammals, 479 toxicity studies, 479–480 Merozoites, 23 Mesenteries, 146 Metabolic rate, 792–794 Methoxyflurane, 734 Methylmercuric chloride, 480 Methylprednisolone, 720 Metoclopramide HCL, 720 Metronidazole cetacean drug dosage, 712 pinniped drug dosage, 716 sea otter drug dosage, 720 sirenian drug dosage, 718 Mexican stranding networks contacts and programs, 59–60 MFO (mixed-function oxidase) system, 488–490 MHC (major histocompatibility proteins), 239 Microphallus pirum, 978 Microsatellite analysis genetic analysis and, 272 parental similarity vs. fitness and, 275 social organization examination using, 274 Microscopic anatomy adrenal glands, 150 digestive system, 148–149 genital tract, 149–150 hematopoietic system, 150 integument, 139–140 liver, 148
CRC Handbook of Marine Mammal Medicine
lymphoid system, 150 parathyroids, 145 respiratory system, 144 thymus, 145 thyroids, 145 urinary tract, 149 Microsporum canis, 345 Midazolam for cetaceans, 660 euthanasia use, 732 for odobenids, 677 for otariids, 670 for phocids, 674 for polar bears, 994 Milbemycin oxime, 721 Mineralocorticoids, 260 Mineral oil, 718 Minimally invasive surgery (MIS) described, 621–622 indications, 622, 624 techniques access, 637–638 cannulas, 638–639 in cetaceans, 640 closure, 636–640 insufflation, 636–637 in other marine mammals, 640 trocars, 638–639 Mink (Mustela vison), 200 Minke whale (Balaenoptera acutorostrata) bacterial diseases in, 314 contaminants’ effects on organ systems, 474 insemination techniques, 225 lactation energy requirements, 805 organochlorine effects on, 490 species identification techniques, 273 viral diseases in, 298 Minocycline, 713 Mirex, 485 Mirounga angustirostris, see Elephant seal Mirounga leonina, see Southern elephant seal MIS, see Minimally invasive surgery Mites in pinnipeds, 372 Mitochondrial DNA (mtDNA), 271–272 Mixed-function oxidase (MFO) system, 488–490 MMC (Marine Mammal Commission), 743, 761 MMPA, see Marine Mammal Protection Act MMUME, see Marine Mammal Unusual Mortality Event Mobbing behavior in pinnipeds, 203 Molting cortisol levels and, 178 energy requirements, 807 fasting during, 801 integument microanatomy and, 140 organotins concentrations and, 482 TH fluctuations and, 176 Monachus schauinslandi, see Hawaiian monk seal
0839_frame_IDX Page 1045 Friday, May 25, 2001 9:37 AM
1045
Index
Monk seal (Monachus monachus) biotoxins effects on, 494 integument microanatomy, 140 mass mortality case, 16 molt energy requirements, 807 viral diseases in, 296 Monoclonal antibodies for bacterial disease treatment, 318 Monocytes and leukocytes evaluation, 402 Monocytosis, 403 Monodon monoceros (narwhales), 290 Monorygma grimaldii, 366 Moraxella, 327 Morbillivirus infections and epizootics in cetaceans, 16, 18–19 description and treatment, 296–298 lack of connection to organochlorines, 492–493 in manatees, 952 in polar bears, 1001 public health significance, 769 serodiagnostics of, 244 in walruses, 935 Morganella morganii, 951 Mortality rate data, 39 Mouth, contaminants’ effects on, 472 MRI, see Magnetic resonance imaging MRP (maternal recognition of pregnancy), 197 mtDNA (mitochondrial DNA), 271–272 Mucor spp., 951 Musculoskeletal system, see also Skeleton diagnostic imaging techniques, 571 gross anatomy, 141–142 lead concentrations in, 481 non-infectious diseases, 532–533 pinniped diseases, 915–916 sonography clinical applications, 616 Mustela vison (mink), 200 Mustelidea, see Sea otter MV (measles virus), 296 Mycobacterial disease, 319–320 Mycobacterium spp. in manatees, 952 in pinnipeds, 918 public health significance, 771–772 Mycoplasma infections, 772–773 Mycotic diseases clinical diagnosis, 340, 349 clinical manifestations, 339 common fungi, 337–338 epidemiology mechanisms of pathogenesis, 338–339 modes of transportation, 338 host species table, 341–348 lesions, diagnostic methods, and treatments Apophysomyces elegans, 345–346 Aspergillus fumigatus, 341–342 Blastomyces dermatitidis, 347 Candida, 342–343
Cladophialophora bantiana, 343 Coccidioides immitis, 347 Cryptococcus neoformans, 343–344 Deuteromyces, 344 Fusarium, 344 Histoplasma capsulatum, 348 Lacazia loboi, 348 Malassezia pachydermatis, 344 Microsporum canis, 345 Rhizomucor pusillus, 346 Saksenaea vasiformis, 346 Scedosporium apiospermum, 345 Sporothrix schenckii, 345 Trichophyton, 345 Trichosporon pullulans, 345 zygomycetes, 345–347 in polar bears, 1001–1002 therapeutics, 349–351 Myocardial lesions, 536
N NADPH cytochrome c reductase, 489 Narwhale (Monodon monoceros), 290 NASA, 857 Nasal mite (Halarachne miroungae), 978 Nasitrema spp. as cause of strandings, 365 cytology findings in respiratory tract, 443 cytology findings in stomach, 444 identifying, 358 implemented in loss of acoutic ability, 86 National Animal Disease Laboratory, 311 National Cancer Institute Web site, 118, 119 National Climatic Data Center (NCDC), 5 National Conservation Training Center (NCTC), 119 National Marine Fisheries Service (NMFS) addresses, 751 ESA and, 745 jurisdiction, 742 purpose of, 761 tagging and tracking, 857 National Marine Mammal Laboratory, 121 National Marine Mammal Tissue Bank (NMMTB), 465 National Marine Services (NMS), 450 National Oceanic and Atmospheric Administration (NOAA), 761 National Research Council (NRC), 803 Natural markings as tags, 851 NCDC (National Climatic Data Center), 5 Necropsy, see Gross necropsy Necrotizing myopathies, 533 Necrotizing pancreatitis, 404–405 Nematodes, see also Helminths in cetaceans, 361–362, 365–366 in manatees, 373 in pinnipeds, 369, 916 in polar bears, 374
0839_frame_IDX Page 1046 Friday, May 25, 2001 9:37 AM
1046
in sea otters, 373 treatment of, 359 Neomycin pinniped drug dosage, 716 sea otter drug dosage, 720 Neonatal disease in manatees, 953–954 Neophoca cinerea (Australian sea lion), 320 Neophocoena phocoenoides (finless porpoises), 325–326 Neoplasia in California sea lions, 21 in manatees, 953 in polar bears, 1000–1001 prevalence and types, 522, 525–528 in sea otters, 979 Neosaxiton, 494 Neosaxitoxin, 21 Nervous system, see Central nervous system The Netherlands stranding networks contacts and programs, 60–61 NetVet Web site, 119 Neurohypophyseal hormones, 169 Neurological dysfunction encephalitis etiology unclear, 537 in manatees, 952 in pinnipeds, 368, 921, 922 encephalomyelitis, 429 helminth parasites in cetaceans, 365–366 hydrocephalus in pinnipeds, 921 parasites as cause of strandings, 365, 366, 368 in pinnipeds, 19 Neutropenia due to drug interactions, 707 Neutrophils cytology findings in respiratory tract, 443 cytology findings in stomach, 444 leukocytes evaluation, 401, 403 New Zealand fur seal (Arctocephalus forsteri), 320; see also Fur seal New Zealand sea lion (Phocarctos hookeri), 296 New Zealand stranding networks contacts and programs, 61 Niclosamide, 721 Nitrofurantoin, 721 Nitrous oxide for cetaceans, 660 sea otter drug dosage, 720 NMFS, see National Marine Fisheries Service NMMTB (National Marine Mammal Tissue Bank), 465 NMS (National Marine Services), 450 NOAA (National Oceanic and Atmospheric Administration), 761 Nocardia in cetaceans, 322, 324–325 clinical signs, 321–322 in pinnipeds, 325 species infected, 323 Noise pollution, 5 Nonachlors, 485
CRC Handbook of Marine Mammal Medicine
Non-infectious diseases cardiovascular system, 535–536 congenital defects, 521–522, 523–524 dental system, 532–533 digestive system, 533–534 endocrine system, 535 genitourinary system, 534–535 gross necropsy, 464–465 integumentary system, 531–532 lymphoid system, 536 musculoskeletal system, 532–533 neoplasia, 522, 525–528 nervous system, 536–537 reproductive system, 534–535 respiratory system, 533 special senses, 536–537 trauma anthropogenic, 530–531 interspecific, 528–530 intraspecific, 522, 524, 528 Nonshivering thermogenesis (NST), 168 Norepinephrine and dive response, 178 Northern elephant seal, see Elephant seal Northern fur seal (Callorhinus ursinus) bacterial diseases in Erysipelothrix, 318 Leptospira, 770 leptospirosis, 321 urogenital, 327 contaminants’ effects on organ systems, 474, 476 estrous cycle and, 196, 197 hand-rearing and artificial milk formulas, 839–840 lactation energy requirements, 805, 806 neurological disfunction in, 19 non-infectious diseases digestive system, 534 genitourinary system, 534 lymphoid system, 536 musculoskeletal system, 533 parasites of, 368, 369, 372 pineal gland size, 167 pregnancy determination, 199 renin-antiotensin system, 185 stress response, 260 viral diseases in caliciviruses, 300 herpesviruses, 293 poxvirus, 286–288 water requirements, 800 Northern right whale dolphin (Lissodelphis borealis), 359 NRC (National Research Council), 803 NST (nonshivering thermogenesis), 168 Nudacotyle undicola, 953 Nursing, see Lactation and nursing Nutrition and energetics bioenergetic scheme fecal and urinary energy losses, 809–810
0839_frame_IDX Page 1047 Friday, May 25, 2001 9:37 AM
Index
gross energy requirements calculation, 810–811 heat increment of feeding, 807–808 ingested energy, 803–804 lactation, 805–807 maintenance energy, 804 molt, 807 production, 804–805 disorders hyponatremia, 814–815 in polar bears, 1000 scombroid poisoining, 816–817 thiamine deficiency, 813–814 vitamin deficiencies, 815–816 energy requirements locomotion, 796–799 maintenance costs, 792 metabolic rate, 792–794 thermoregulation, 794–796 fasting and starvation, 801–803 manatees, 943–944 nutritional therapy appetite stimulants, 694 emaciation, 693–694 hypoglycemia, 693 pinnipeds, 908 polar bears adult, 990–991 geriatrics, 992 juveniles, pregnant, lactating, 991–992 prey seasonality of composition, 813 species consumed in captivity, 811–812 sea otters, 967–968, 971 walrus diet, 929 water requirements, 799–801 Nyotran, 349 Nystatin cetacean drug dosage, 713 mycotic infection use, 349, 350 pinniped drug dosage, 716
O Obstructive emphysema, 533 OC, see Organochlorine (OC) chemicals Octachlorostyrene (OCS), 487, 488 Ocular lesions, 536; see also Ophthalmology Odobenus rosmarus spp., 314 Odontocetes, see also Dolphins; Porpoises; Whales digestive system, 148–149 organochlorine effects on, 489 organohalogens accumulation in, 487 Office International des Epizooties (OIE), 881 Ofloxacin, 713 OIE (Office International des Epizooties), 881 Oil constituents, 496 contact exposure effects, 496–498
1047
exposure problems for sea otters, 976–977 oral-dosing studies, 498 petroleum constitutents in animals, 498–499 OMNI (Organized Medical Networked Information), 119–120 Omphalophlebitis, 1001 Opaleye perch (Girella nigricans), 301 Ophthalmology ocular lesions, 536 pinniped diseases, 920–921 sonography imaging, 616 walrus diseases, 932–933 Optimum Sustainable Population (OSP), 750 Orcas, see Killer whale Orcinus orca, see Killer whale Organized Medical Networked Information (OMNI), 119–120 Organochlorine (OC) chemicals captive animals studies of, 8 contamination by, 4–5 pesticides and metabolites, 484–485 toxicity studies, 488–490 Organohalogens and adrenal hyperfunction, 264 Organotins, 481–482 Orthohalarachne, 372 Orthopoxvirus, 288 Orthosplanchnus fraterculus, 978 Osmoregulatory hormones atrial natriuretic peptide, 185 renin-antiotensin system, 185 vasopressin, 183–184 OSP (Optimum Sustainable Population), 750 Osteomyelitis, 532 OT, see Oxytocin Otaria byronia (South American sea lion), 286, 288 Otariids, see also Pinnipeds; specific species anesthesia emergencies, 669–670 immobilizing agents used, 663–665 induction, 662, 666 inhalation anesthesia, 667 intubation, 666–667 monitoring, 668 support, 668–669 clinical pathology blood collection, 385 references, 397 estrous cycle, 195, 196 fasting during lactation, 803 integument microanatomy, 140 lactation energy requirements, 806 lactation in, 200 lymphoid and hematopoietic system, 150 molt energy requirements, 807 parasites of, 372 pleura and lungs, 143 respiratory system, 144 salinity and pH requirements, 781
0839_frame_IDX Page 1048 Friday, May 25, 2001 9:37 AM
1048
sexual dimorphisms, 157 Otostrongylus circumlitus in northern elephant seals, 536 in pinnipeds, 918, 919 Ovaries described, 148 sonography imaging, 612–615 Ovulation hormonal regulation of, 194 manipulation and control, cetaceans induction, 221–222 synchronization, 222–223 Oxacillin, 720 Oxychlordane, 485 Oxytetracycline polar bear drug dosage, 721 sirenian drug dosage, 718 Oxytocin (OT) dystocia treatment and, 697 lactation in pinnipeds and, 200 pinniped drug dosage, 716 purpose of, 169 reproduction regulation role, 194 sea otter drug dosage, 720 Ozone use for water sterilization, 785–786
P Pacific harbor seal, 294, 295; see also Harbor seal Pacific walrus (Odobenus rosmarus divergens), 300; see also Walrus Pacific white-sided dolphin (Lagenorhynchus obliquidens) clinical pathology, 426–427 fasting, 803 hepatobiliary system disorders markers, 406, 410 non-infectious diseases cardiovascular system, 536 endocrine system, 535 genitourinary system, 534 thermoneutral zone, 795 thyroxine levels, 174 viral diseases in, 302, 426–427 Packed cell volume (PCV), see Erythrocytes, evaluation Pagophilus groenlandicus, see Harp seal Pain relief, 699 Pancreas endocrine system and, 185–186 helminth parasites in pinnipeds, 372 location, 146 pancreatitis, acute, 405, 534 serum analytes and enzymes, 404 sonography clinical applications, 605 Panophthalmitis, 536 Pan-tropical spotted dolphin (Stenella attenuata), 534 Papillomavirus animals found in, 19 description and treatment, 289–291
CRC Handbook of Marine Mammal Medicine
in manatees, 952 PAR (pathogen antimicrobial resistance), 24 Parafilaroides decorus (lungworm) in pinnipeds, 918 viral disease and, 301 Paralytic agents for euthansia, 733 Paralytic shellfish poisoning (PSP), 494–495 Paramyxoviridae, see Morbillivirus infections Parapoxvirus, 288 Parasitic diseases of cetacea ectoparasites, 367 helminths, 361–367 protozoa, 359–361 of manatees, 952–953 of pinnipeds ectoparasites, 372 helminths, 369–372 protozoa, 367–368 of polar bears, 374, 1002 removal and fixation of parasites, 357–358 of sea otters, 978 apicomplexans, 373 helminths, 373–374 of sirenia apicomplexans, 372 helminths, 373 treatment, 359 Parasitology, 462–463 Parathyroid, 144, 145, 416 Parathyroid hormone (PTH), 416 Parturition induction in cetaceans, 212, 215 in pinnipeds, 199 Passive integrated transponder (PIT) manatee tagging and tracking, 866 sea otters tagging and tracking, 870 Passive tags, 851, 862 Pasteurella, 312 Pasteurella hemolytica, 312 Pasteurella multocida in cetaceans, 315 in polar bears, 1001 retrovirus and, 302 in sea otters, 971 Pasteurellosis, 315–316 Paternity testing techniques, 275–276 Pathogen antimicrobial resistance (PAR), 24 Pathogen-specific antibodies measurement, see Serodiagnostics PBBs (polybrominated biphenyls), 488; see also Organochlorine (OC) chemicals PBDEs (polybrominated diphenyl ethers), 487, 488 PCBs (polybrominated biphenyls), 485–486 PCDDs (polychlorinated dibenzo-p-dioxins), 486, 489 PCDFs, 486 PCNs (polychlorinated napthalenes), 486 PCP (porcine zona pellucida vaccine), 201
0839_frame_IDX Page 1049 Friday, May 25, 2001 9:37 AM
Index
PCQs (polychloroquaterphenyls), 486 PCTs (polychlorinated terphenyls), 487, 488 PCV (packed cell volume), see Erythrocytes, evaluation PDV (phocine distemper virus), 7, 273, 289, 293, 296, 917 Pectoral limb complex, 156–157 Pelecanus occidentalis (brown pelican), 494 Pelvic limb complex, 157 Penicillin pinniped drug dosage, 717 polar bear drug dosage, 721 sea otter drug dosage, 720 sirenian drug dosage, 718 Pentobarbitol use for euthanasia, 732 Pentoxyresorufin-o-depentylase (PROD), 489 Pepto-Bismol cetacean drug dosage, 713 polar bear drug dosage, 721 Pericardium, 142 Periodontitis, 532 Peritonitis, 1001 Permits under ESA, 745 MMPA, 750, 752–753 for observations, 765 public display, 764–765 research and enhancement, 763–764, 765 stranding networks, 762–763 Peruvian stranding networks contacts and programs, 62 Peste-des-petits ruminants virus (PPRV), 296 Pethidine, 674 Phalacrocorax penicillatus (Brandt’s cormorants), 495 Pharmaceuticals and formularies cautions for, 703–704 dose scaling, 705 drug dosages cetaceans, 709–714 pinnipeds, 714–718 polar bear, 721–722 sea otters, 719–721 sirenians, 718–719 drug interactions aminoglycosides, 707 antacids, 705–706 antifungals, 708 antiparasitic drugs, 708 azithromycin, 707 cephalosporins, 707 chloramphenicol, 707 cimetidine, 705–706 diuretics, 708 florfenicol, 707 fluoroquinolones, 706 rifampin, 707 steroids, 708 sulfonamides, 707 tetracyclines, 706 drug routes, 704–705
1049
Phenobarbital pinniped drug dosage, 717 sea otter drug dosage, 720 Phenylephrine, 720 Phenytoin cetacean drug dosage, 713 pinniped drug dosage, 717 sea otter drug dosage, 720 PhHV-1 (phocid herpesvirus type-1), 7, 293, 295 PhHV-2 (phocid herpesvirus-2), 294, 295 Phoca caspica (Caspian seal), 296 Phoca hispida, see Ringed seal Phoca largha (spotted seals) trauma treatment, 695 viral diseases in, 293 Phocarctos hookeri (New Zealand sea lions), 296 Phoca sibirica (Baikal seal), 184 Phoca vitulina, see Harbor seal Phocid herpesvirus-2 (PhHV-2), 294, 295 Phocid herpesvirus type-1 (PhHV-1), 7, 293, 295, 918 Phocids, see also Pinnipeds; Seals; specific species aldosterone levels, 181 anesthesia emergencies, 676–677 immobilizing agents used, 671–673 induction, 670, 674 inhalation anesthesia, 675 intubation, 674–675 monitoring, 675 support, 675–676 clinical pathology blood collection, 385 references, 397–398 estrous cycle, 195, 196 fasting during lactation, 802 lactation energy requirements, 806 lactation in, 200 lymphoid and hematopoietic system, 150 molt energy requirements, 807 nutrition, 814–815 pleura and lungs, 143 reproductive tract, 150 respiratory system, 144 spinal cord, 151 thermal insult protection during diving, 153 TH fluctuations and, 176 Phocine distemper virus (PDV), 7, 273, 289, 293, 296, 917 Phocoena phocoena (harbor porpoise), see Harbor porpoise Phocoena sinus (vaquita), 313 Phocoena spinipinnis (Burmeister’s porpoises), 19 Phocoenoides dalli (Dall’s porpoises), 474 Phosphorus, 419 Photoperiod embryonic diapause in pinnipeds, 198 lighting in captivity considerations, 908 melatonin and, 167
0839_frame_IDX Page 1050 Friday, May 25, 2001 9:37 AM
1050
reproductive cycle and, 195 in cetaceans, 205 in pinnipeds, 193 Phyllobothrium delphini, 366 in pinnipeds, 372 Physeter catadon (great sperm whale), 6–7 Physeter macrocephalus (sperm whales), see Sperm whale Pilot whale (Globicephala macrorhynchus) bacterial diseases in Brucella, 313–314 Nocardia, 322 corpora albicantia in, 210 diseases in, 18 graded histopathological findings, 93 hematology and biochemistry values, 392 hepatobiliary system disorders markers, 406, 409 mass strandings investigation, 86–87 mercury concentration in liver, 478 non-infectious diseases digestive system, 534 endocrine system, 535 genitourinary system, 534 lymphoid system, 536 musculoskeletal system, 532 organohalogens accumulation in, 488 parasites of, 359 rehydration, 691 stress response, 259–260 viral diseases in influenza, 298, 299 morbillivirus, 296, 298 Pineal gland described, 151 endocrinology, 167–168 reproduction regulation role, 195 Pinniped reproduction abnormalities, 203 contraception and control of aggression females, 202 males, 202–203 purpose of, 201 energy requirements, 805 female embryonic diapause and reactivation, 198 estrous cycle, 196–197 implantation, 198–199 lactation, 200 milk collection, 200 parturition induction, 199 pregnancy and pseudopregnancy, 197 pregnancy diagnosis, 199 reproductive cycle, 195–196 male anatomy, 200–201 seasonality of fertility, 201–202 sexual maturity, 201 reproductive tract, 150
CRC Handbook of Marine Mammal Medicine
Pinnipeds, see also specific species adrenal glands, 150, 178 age estimation, 455–456 aldosterone concentrations, 182 bacterial diseases in Brucella, 314 Erysipelothrix, 318–319 gastrointestinal, 327–328 leptospirosis, 320–321 mycobacterial disease, 319–320 pasteurellosis, 315–316 respiratory, 325 urogenital, 327 vibriosis, 315 cadmium toxicosis, 480–481 clinical pathology hematology and biochemistry values, 394–395 references, 397–398 cortisol concentrations, 179–180 diagnostic imaging applications radiographic anatomy, 581–585 radiographic pathology, 585–586 diagnostic imaging techniques for, 566 diagnostic techniques, 910–911 digestive efficiencies, 809 digestive system, 145, 146, 148–149 diseases in cardiovascular system, 919 digestive system, 916–917 endocrine system, 920 eyes, 920–921 integumentary system, 913–915 musculoskeletal system, 915–916 nervous system, 921–922 respiratory system, 917–918 urogenital system, 919–920 drug dosages, 714–718 electrolyte disorders, 416 emerging and resurging diseases in bacterial, 22 domoic acid-induced, 19, 21 table of, 20–21 urogenital neoplasia examination, 21–22 fasting, 801 hand-rearing and artificial milk formulas elephant seals, 836–838 harbor seals, 832–834 sea lions, 839–840 walruses, 841–842 heart and pericardium gross, 142 hepatobiliary system disorders markers, 406, 408 husbandry, 907–908 hyponatremia care, 696 integument microanatomy, 140 liver, 148 lymphoid and hematopoietic system, 150 nervous system, 151 non-infectious diseases
0839_frame_IDX Page 1051 Friday, May 25, 2001 9:37 AM
Index
cardiovascular system, 535–536 musculoskeletal system, 532 nervous system, 536 nutrition therapy for, 694 vitamin deficiencies, 816 organohalogens accumulation in, 487 parasites of apicomplexans, 367–368 ectoparasites, 372 flagellates, 368 helminths, 369–372 PCB toxicity and, 486 physical examination, 909–910 pleura and lungs, 143 regulating agency, 742 reproduction, see Pinniped reproduction restraint, 908–909 salinity and pH requirements, 781 skeletal muscles use for swimming, 141 space requirements, 780 species status under ESA, 748–749 tagging and tracking branding, 858 flipper tags, 858 impact of devices, 862 tag location and attachment, 858–859 TDR location and attachment, 861–862 telemetry device location and attachment, 859–861 therapeutic techniques, 911–912 thermal imaging applications, 646 thermal insult protection during diving, 153 thyroids, 143, 145 thyroxine concentrations, 171–172 thyroxine levels, 170 transport of, 888 triiodothyronine concentrations, 173–174 viral diseases in herpesviruses, 295 morbillivirus, 296–298 poxvirus, 286–288 Piperazine pinniped drug dosage, 717 polar bear drug dosage, 721 Pitressin, 720 Pituitary gland, 151 Placentonema, 366 Plasma fibrinogen, 902 Plasma prekallikrein (Fletcher Factor activity), 421 Platform Transmitter Terminals (PTTs), 855–857 Plesiomonas, 327 Pleura, 143 Pleuritis in polar bears, 1001 PMV (porpoise morbillivirus), 16, 296 Pneumonia in cetaceans, 325–326 due to non-infectious diseases, 533
1051
indications in cetaceans, 901 in pinnipeds, 325–326, 918 Pneumothorax therapy, 695 Pod cohesion as factor in mass strandings, 86 Point-contact sterilization, 784 Polar bear (Ursus maritimus), 24 anesthesia carfentanil, 999 delivery and monitoring, 996–997 etorphine, 999 fentanyl citrate, 999 inhalation agents, 999–1000 ketamine, 997–998 ketamine/xylazine, 998 telazol/medetomidine, 999 tiletamine HCL, 998–999 zolazepam HCL, 998–999 behavior, 994 clinical pathology blood collection, 390 hematology and biochemistry values, 396 references, 399 contaminants’ effects on organ systems, 474, 476 cortisol concentrations, 180 dehydration indications, 690 drug dosages, 721–722 emerging and resurging diseases in, 24 endocrine system reproductive hormones, 992–993 thyroid hormones, 993 FWS jurisdiction, 742 hand-rearing and artificial milk formulas, 847–848 hepatobiliary system disorders markers, 407, 408, 409, 411 housing, 993–994 milk fat content, 807 natural history, 989 non-infectious diseases, 532 nutrition adult, 990–991 geriatrics, 992 juveniles, pregnant, lactating, 991–992 organohalogens accumulation in, 487 parasitic diseases of, 374 PCB toxicity and, 486 physical examination, 994–995 physiology, 989–990 rehydration, 691 reproduction, 201, 992 restraint, 996 serum analytes and enzymes, 404 space requirements, 780 species status under ESA, 749 systemic diseases dental, 1003 developmental/anomalous, 1000 infectious, 1001–1002 neoplasia, 1000–1001
0839_frame_IDX Page 1052 Friday, May 25, 2001 9:37 AM
1052
nutritional, 1000 skin, 1002–1003 tagging and tracking, 874 thermal imaging applications, 649 thermal insult protection, reproductive organs, 201 thyroxine concentrations, 172 thyroxine levels, 170 toxins, 1003 transport of, 889 trauma treatment, 696, 1003 triiodothyronine concentrations, 174 venipuncture, 995 viral diseases in, 296 zoonoses, 1003 Pollution, 5; see also Toxicology Polybrominated biphenyls (PBBs), 488; see also Organochlorine (OC) chemicals Polybrominated diphenyl ethers (PBDEs), 488 Polychlorinated biphenyls (PCBs), 485–486 Polychlorinated camphenes, 485 Polychlorinated dibenzo-p-dioxins (PCDDs), 489 Polychlorinated napthalenes (PCNs), 486 Polychlorinated terphenyls (PCTs), 488 Polychloroquaterphenyls (PCQs), 486 Polychromasia, 399–400 Polymorphus spp., 23 Pool water, see Water and environment quality Population identification techniques, 273–274 Population trends estimation, 37 Population viability analysis (PVA), 40 Porcine zona pellucida vaccine (PCP), 201 Porpoise morbillivirus (PMV), 16, 296 Porpoises, see also Cetaceans; Odontocetes; specific species digestive system, 148–149 hepatobiliary system disorders markers, 407, 410 organochlorine effects on, 489 organohalogens accumulation in, 487 water requirements, 800 Posaconazole, 351 Postthoracic vertebrae, 156 Potassium, 416–417, 721 Poxviruses clinical signs, 287 diagnosis, 288 differential, 288–289 epidemiology, 289 host range, 286 pathology, 287–288 public health significance, 289, 768 therapy, 287 PPRV (peste-des-petits ruminants virus), 296 Praziquantel drug dosage cetaceans, 713 pinnipeds, 717 sea otters, 720
CRC Handbook of Marine Mammal Medicine
sirenians, 718 nematode treatment, cetaceans, 362 nematode treatment, pinnepeds, 370 for parasite treatment, 359 trematode treatment, cetaceans, 365 trematode treatment, pinnipeds, 372 Precipitation/agglutination techniques, 244–245 Prednisolone adverse effects, 708 cetacean drug dosage, 713 polar bear drug dosage, 721 Prednisone, 370 Pregnancy, see Reproduction Pregnant mare serum gonadotropin, 713 Prey and nutrition seasonality of composition, 813 species consumed in captivity, 811–812 Pricetrema, 370 Primidone, 717 PRL, see Prolactin Procaine penicillin G, 713 PROD (pentoxyresorufin-o-depentylase), 489 Profenal, 717 Professional societies and organizations, 112–113 Profilicollis altmani, 978 Progesterone photoperiod changes and, 198 in pinnipeds, 197, 198 reproduction regulation role, 194 Prolactin (PRL) hypothalamus-pituitary and, 169 lactation in pinnipeds and, 200 reproductive cycle and, 195 Prolactin-releasing factor, 200 ProMED Web site, 118 Propofol, 660 Propriopromazine, 717 Prostaglandin F2α ovulation synchronization and, 222 reproduction regulation role, 194 Prothrombin time, 421–422 Protozoans, 23 Pseudaliids, 366 Pseudomonas spp. cetacean infection case study, 425 dermatological disease and, 326 in manatees, 951 in sea lions, 326 septicemia and, 312 in walruses, 935 Pseudonitzschia australis, 5, 7, 19, 21 Pseudorabies, 952 Pseudorca crassidens, see False killer whale PSP (paralytic shellfish poisoning), 494–495 PTH (parathyroid hormone), 416 PTTs (Platform Transmitter Terminals), 855–857 Ptychodiscus brevis, 5, 493
0839_frame_IDX Page 1053 Friday, May 25, 2001 9:37 AM
Index
Public health significance of marine diseases bacterial infections Brucella, 771 Clostridium, 770 Coxiella burnetti, 772 Edwardsiella, 770 Erysipelothrix, 771 Leptospira, 770 mixed infections, 772 Mycoboacterium, 771–772 Streptococcus, 770–771 Vibrio, 769–770 fungal infections, 773 human contraction of diseases caliciviruses, 302 Erysipelothrix, 316 herpesviruses, 295 influenza, 300 Nocardia, 322 papillomavirus, 291 poxvirus, 289 mycoplasma infections, 772–773 potential for transmission, 774–775 protozoal infections, 773–774 scope of concerns, 767–768 viral infections caliciviruses, 768–769 influenza, 769 poxviruses, 289, 768 rabies, 769 PubMed Web site, 118, 119 Pulmonary aspergillosis, 337 Pulse rates of transmitters, 852 Pupil dilation after anesthesia administration, 670 Pustular folliculitis, 931 PVA (population viability analysis), 40 Pylorus, 606 Pyometra, 698 Pyridoxine, 717
Q Quinolones, 312
R Rabies found in ringed seal, 303 in manatees, 952 in polar bears, 1001, 1003 public health significance, 769 Radiology, see also Diagnostic imaging; Ultrasonography clinical applications dolphin, 574–581 pinnipeds, 581–586 computed tomography, 558–559, 586 computer radiography, 557–558
1053
indications for abdominal radiography, 561–562 for bone films, 562–563 CT and MRI use, 563 positive-contrast media use, 563 for thoracic radiographs, 561 limitations facilities and equipment, 566–568 portable diagnostic units, 565–566 risks of removing animals from water, 565 magnetic resonance imaging, 558, 560, 587–589 techniques, 568–573 Radio tags for pinniped tracking, 859 Radio telemetry use and attachment, manatees, 867–868 Ranitidine cetacean drug dosage, 713 sea otter drug dosage, 720 RAS (renin-antiotensin system), 185 Red blood cells (RBC), see Erythrocytes, evaluation Red deer (Cervus elaphus), 275 Red tides, 19, 21, 493, 951; see also Harmful algal blooms Regulating agencies, 742–743; see also Federal legislation and regulations Rehydration, 690–692 Renal calculi in pinnipeds, 920 Renal toxicity due to drug interactions, 707 Reniculate kidneys, 147, 149 Renin-antiotensin system (RAS), 185 Reproduction abnormalities, 534–535 in cetaceans, see Cetacean reproduction contaminants’ effects on, 473–474 emergency care, 697–698 energy requirements, 804–805 halogenated organics effects on, 490–491 manatees, 941 necropsy examination of status, 456–457 non-infectious diseases of, 534–535 PCB toxicity and, 486 physiology, 193–195 in pinnipeds, see Pinniped reproduction polar bears, 992 sea otters, 965–966 stress response indicators, 263 tracts female, 147–148 male, 148 sonography clinical applications, 611–616 thermal insult protection, male, 152–153 Respiratory system contaminants’ effects on, 472 cytology findings, cetaceans, 442–443 disease described, 325–326 diagnostic imaging techniques, 571 helminth parasites, cetaceans, 365–366 helminth parasites, pinnipeds, 370–372 emergency care, 695
0839_frame_IDX Page 1054 Friday, May 25, 2001 9:37 AM
1054
lungs gross anatomy, 143 sonography clinical applications, 597–600 microscopic anatomy, 144 non-infectious diseases, 533 pinniped diseases, 917–918 Restraint of marine mammals, see also Transport of marine mammals of polar bears, 996 sea otters chemical, 973 physical, 971–972 Restriction fragment length polymorphisms (RFLP), 271 Reticulocyte counts, 902 Retrovirus, 302–303 Return to sea as stranding response, 94 RFLP (restriction fragment length polymorphisms), 271 Rhabdoviruses, 303; see also Rabies Rhizomucor pusillus, 346 Rhodococcus equi, 327 Ribbon seal (Histriophoca fasciata), 293 Ribs, 155 Rickets, 532, 1000 Rifampin adverse effects, 707 cetacean drug dosage, 713 pinniped drug dosage, 717 Rigid telescopes, 626 Rinderpest virus (RPV), 296 Ringed seal (Phoca hispida) aldosterone levels, 181 bacterial diseases in, 22, 314 contaminants’ effects on organ systems, 473 heat increment of feeding, 808 non-infectious diseases, 534 organochlorine reproductive function effects, 490 osmoregulatory hormones, 184 parasites of, 368 stress response, 260 thermoneutral zone, 794 viral diseases in herpesviruses, 293 rhabdoviruses, 303 Risso’s dolphin (Grampus griseus) hepatobiliary system disorders markers, 410 mercury concentration in liver, 478 parasites of, 360 parathyroids, 144, 145 tremotode as cause of strandings, 365 River dolphins, 532 River otter (Lontra canadensis), 19 Ronnel, 717 Roto-radio tag, 863–864 RPV (rinderpest virus), 296
CRC Handbook of Marine Mammal Medicine
S Sacral vertebrae, 156 Saksenaea vasiformis, 346 Salinity and pH in waters, 781 Salivary glands, 146 Saliva sample collection and handling, 166 Salmonella spp. gastrointestinal disease and, 327 in pinnipeds, 916 in sea otters, 971 septicemia and, 312 Sampling and collection bacterial diseases, 310–311 blood from cetaceans, 90, 385 equipment and processing, 384–385 manatees, 946–947 pinnipeds, 910 polar bears, 995 sample handling, 166 sites, 385–390 cytology, cetacean aspirates from masses, 439 fecal, 439 gastric, 438–439 guidelines, 438 respiratory tract, 438 urinary tract, 439 genetic analysis techniques, 277–278 necropsy examinations and specimen collection, 450 parasite removal and fixation, 357–358 semen, cetacean, 219–221 tissues, 167 urine, 166–167 manatees, 947 pinnipeds, 910–911 virus isolation, 285–286 San Diego Library Consortium, 118 San Miguel Island, Calif., 7 San Miguel sea lion virus (SMSV) described, 300–302 in manatees, 952 in pinnipeds, 913, 920 public health significance, 768–769 serodiagnostics of, 244 in walruses, 935 Sarcocystis spp. anthropogenic origin, 5 in pinnipeds, 22, 368, 916 in sea otters, 23, 429, 978 species range, 359 Sarcodina, 360 Sarcoptes (mites), 372 Satellite telemetry cetaceans tagging and tracking, 863 description and uses, 855–857 pinnipeds tagging and tracking, 860
0839_frame_IDX Page 1055 Friday, May 25, 2001 9:37 AM
Index
Saxidomus giganteus (butter clam), 494 Saxitoxin, 494 Scedosporium apiospermum, 345 Scientific societies and membership organizations, 112–113 Scolex pleuronectis, 367 Scoliosis, 532, 533 Scombroid poisoning, 816–817 SDH (sorbitol dehydrogenase), 407–408 Sea Grant colleges, 122 Seal finger, 771, 772–773 Sea lion (Otaria byronia) caval sphincter, 144 digestive system, 146 external features, 138 hyponatremia care, 696 liver, 145 locomotion, 154 muscle mass, 141 neoplasic pathogenesis studies, 7 PCDDs concentrations in, 487 pectoral limb complex, 157 pelvic limb complex, 157 rehydration, 692 water requirements, 800 Seal louse (Echinophthirius horridus), 372, 919 Seal pox, 913 Seals, see also Phocids; Pinnipeds; specific species caval sphincter, 144 digestive system, 146 external features, 139 hepatobiliary system disorders markers, 407 locomotion, 154 muscle mass, 141 population structure determination, 274 water requirements, 800 Sea otter (Enhydra lutris) anatomy, 963–964 anesthesia emergencies, 684 immobilizing agents used, 682 induction, 681, 683 inhalation anesthesia, 683 intubation, 683 monitoring and support, 683 classification, 962 clinical examination, 973–974 clinical pathology blood collection, 387 hematology and biochemistry values, 396 references, 399 cortisol concentrations, 180 dehydration indications, 690 dentistry, 980 digestive efficiencies, 810 digestive system, 146 drug dosages, 719–721 emerging and resurging diseases in, 23–24
1055
feeding and metabolism, 967–968 FWS jurisdiction, 742 hand-rearing and artificial milk formulas, 845–846 heat increment of feeding, 808 hepatobiliary system disorders markers, 407, 408, 411 history, 961–962 husbandry, 969–970 integument microanatomy, 140 leukocytes evaluation, 403 lymphoid and hematopoietic system, 150 medical problems bacterial infections, 979 capture myopathy, 979 clinical chemistry abnormalities, 977–978 coat condition loss, 975–976 gastroenteritis, 978 herpesviruses, 979 hyperthermia, 974–975 hypoglycemia, 974 hypothermia, 975 neoplasia, 979 oil exposure, 976–977 parasites, 978 milk fat content, 807 mortality in the wild, 967 non-infectious diseases, 534 nutrition, captive, 971 organohalogens accumulation in, 488 parasitic diseases of apicomplexans, 373 helminths, 373–374 PCB toxicity and, 486 population structure determination, 274 preventive medicine, 980 rehydration, 691, 692 reproduction, 965–966 restraint chemical, 973 physical, 971–972 social organization, 965, 970 space requirements, 780 species status under ESA, 749 surgery, 979–980 tagging and tracking attachment methods, 870–871 intra-abdominal radio transmitters, 871–873 PITs, 870 thermal insult protection, reproductive organs, 201 thyroxine concentrations, 172 thyroxine levels, 174 transport of, 888–889 vision, 965 water requirements, 800 Secondary sexual characteristics in pinnipeds, 201 Sedative agents cetaceans, 657 manatees, 945
0839_frame_IDX Page 1056 Friday, May 25, 2001 9:37 AM
1056
for phocids, 909 Sei whale (Balaenoptera borealis) bacterial diseases in, 314 parasites of, 359 viral diseases in adenoviruses, 291 caliciviruses, 300 Selenium mercury concentration and, 479, 482 sea otter drug dosage, 720 Semen collection and storage, 219–221 Sense organs and contaminants’ effects on, 472 Sentinel systems characteristics of, 3–4 ecosystem changes detected by, 4–5 marine mammals, use as biomarkers, 6 data handling, 8 effects of anthropogenic noise, 7 for fish stock analysis, 6 hearing and sound production capabilities, 6–7 stranded mammals information, 7 useful characteristics, 5–6 value of mammals in collections, 7–8 Septicemia, 312 Septic metritis, 952 Serodiagnostics enzyme-linked immunosorbent assay, 245 importance of controls, 243–244 precipitation/agglutination techniques, 244–245 serum/virus neutralization test, 244 total immunoglobulin, 245 Serum albumin, 902 Serum amyloid A, 241 Serum analytes and enzymes amylase, 405 cholesterol, 404–405 creatinine phosphokinase, 420 glucose, 403–404 lipase, 405 lipids, 404 pancreatic enzymes, 404 triglycerides, 404–405 trypsin-like immunoreactivity, 405 uric acid, 420 Serum chemistry findings of stranded cetaceans, 91 Serum iron concentration immunodiagnostics and, 241 inflammation marker, 422–423 inflammatory disease indication, 902 Serum protein electrophoresis (SPEP), 414 Serum proteins albumins, 414 globulins, 414–415 hematocrit, 413 total plasma protein, 413 Serum urea nitrogen (BUN), 411–412 Serum/virus neutralization test (SNT/VNT), 244
CRC Handbook of Marine Mammal Medicine
Service Argos system, 855–857 Sevoflurane, 667 Sexual dimorphisms, 157 Shark wounds in pinnipeds, 915 Short-finned pilot whale (Globicephala macrorhynchus), see Pilot whale Siderotic plaques, 536 Silent Spring (Carson), 3 Silver, 482 Simonsiella, 443 Sinus diseases, 365–366 Sirenians, see also Manatee aldosterone concentrations, 182 anesthesia, 681 cortisol concentrations, 180 digestive system, 147, 149 drug dosages, 718–719 hepatobiliary system disorders markers, 406 integument microanatomy, 140 mercury concentration in liver, 478 metabolic rate, 793 musculoskeletal system, 141–142 parasitic diseases of apicomplexans, 372 helminths, 373 PCB toxicity and, 486 pleura and lungs, 143 salinity and pH requirements, 781 skeletal muscles use for swimming, 141 space requirements, 780 species status under ESA, 749 thyroxine concentrations, 172 transport of, 889 triiodothyronine concentrations, 174 Skeleton, see also Musculoskeletal system bone marrow, 158 chevron bones, 156 contaminants’ effects on, 474 limb complexes, 156–157 locomotion, 154 overview, 153–154 ribs, 155 sexual dimorphisms, 157 sternum, 155–156 vertebrae, 154–155, 156 Skin of marine mammals, see Integument SMSV, see San Miguel sea lion virus SNOMED (Systemized Nomenclature of Human and Veterinary Medicine), 119 SNT/VNT (serum/virus neutralization test), 244 Soay sheep (Aries aries), 275 Social behavior assessing in cetaceans, 898–899 manatees, 940 polar bears, 994 sea otters, 965, 970 Sodium, 416 Sodium bicarbonate, 720
0839_frame_IDX Page 1057 Friday, May 25, 2001 9:37 AM
Index
Sodium chloride, 717 Sodium iodine, 720 Software, imaging, 557 Somatotropin, 169 Sonography, 561, 562; see also Ultrasonography Sorbitol dehydrogenase (SDH), 407–408 South American sea lion (Otaria byronia), 286, 288 Southern elephant seal (Mirounga leonina), see also Elephant seal fasting during lactation, 803 integument microanatomy, 140 melatonin concentrations in, 167–168 Southern fur seal (Arctocephalus australis), 22; see also Fur seal Southern sea otter, see Sea otter Spaeck finger, 772–773 Spanish stranding networks contacts and programs, 62 Species identification techniques, 273 SPEP (serum protein electrophoresis), 414 Sperm whale (Physeter macrocephalus) lactation energy requirements, 805 non-infectious diseases, 532 nutrition, 816 parasites of, 359–360 population structure determination, 274 sexual dimorphisms, 157 viral diseases in caliciviruses, 300 papillomavirus, 19, 289–290 Spinal cord, 151 Spinner dolphin (Stenella longirostris) bacterial diseases in, 322 non-infectious diseases, 536 parasites of, 360 thermoneutral zone, 795 Spleen, 146, 604–605 Sporothrix schenckii, 345 Spotted dolphin (Delphinus spp.), 795 Spotted seal (Phoca largha) trauma treatment, 695 viral diseases in, 293 Spumavirus, 302–303 St. Lawrence estuary, 7 Stanozolol, 720 Staphylococcus spp. cetacean infection case study, 425 in manatees, 951, 952 pneumonia in dolphins and, 326 septicemia and, 312 in walruses, 931 Starvation, 801–803 Steller sea lion (Eumetopias jubatus) bacterial diseases in, 327 fasting during lactation, 803 hand-rearing and artificial milk formulas, 839–840 heat increment of feeding, 808 hematology and biochemistry values, 394–395
1057
non-infectious diseases integumentary system, 531 lymphoid system, 536 organotins concentrations, 482 parasites of, 369 reproduction energy requirements, 805 sentinel role, 6 viral diseases in caliciviruses, 300 herpesviruses, 293 Stenella attenuata (pan-tropical spotted dolphin), 534 Stenella coeruleoalba, see Striped dolphin Stenella longirostris, see Spinner dolphin Stenurus, 358 Sterilization methods for pool waters, 784–786 Sternum, 155–156 Steroids adverse effects, 708 Stillbirth in cetaceans, 219 Stomach, see Gastrointestinal system Stranding networks contacts and programs worldwide Argentina, 46–47 Australia, 47–48 Belgium, 48–49 Brazil, 49–50 Canada, 51 Caribbean, 52–53 Croatia, 52 Denmark, 53 France, 54–55 Germany, 54 Greece, 55 Hong Kong, 55–56 Israel, 56 Italy, 56–57 Japan, 57–58 Maldives, 58 Malta, 58–59 Mexico, 59–60 The Netherlands, 60–61 New Zealand, 61 Peru, 62 Spain, 62 Sweden, 63 Ukraine, 63 United Kingdom, 64–65 United States, 65–66 criteria for euthanasia, 729–730 genetic analysis population identification, 273–274 social organization, 274–275 species identification, 273 mass strandings, cetaceans current investigations into, 86–87 definition, 83 disposition of animals, 92, 94 east coast of U.S. (1987-99), 84–85 evaluation of, 87
0839_frame_IDX Page 1058 Friday, May 25, 2001 9:37 AM
1058
graded histopathological findings, 93 management of, 88–92 serum chemistry findings, 91 theories to explain, 83, 85–86 treatment precautions, 94 MMPA oversight, 753–754 objectives of, 45–46 parasites as cause of, 365, 366, 368 permits, 762–763 Streptococcus cetacean infection case study, 425 public health significance, 770–771 septicemia and, 312 Streptomycin, 713 Stress in marine mammals acute response indicators, 262 chronic response indicators, 263–264 future research needs, 264–265 response and regulation baseline data collection, 254 endocrine factors, see Endocrinology, stress response components immunological factors, 261 neurological factors, 255 stressors, 253–254 Striped dolphin (Stenella coeruleoalba) bacterial diseases in, 313–314 contaminants’ effects on organ systems, 475 diseases in, 18 mercury concentration in liver, 478 non-infectious diseases, 535 parasites of, 359, 360 PCB toxicity and, 486 viral diseases in, 296 Strobilocephalus triangularis, 362 Sucralfate, 713 Sulfasalazine, 718 Sulfonamides adverse effects, 707 Surgery on cetaceans, 903 Survey definition, 37 Swedish stranding networks contacts and programs, 63 Systemized Nomenclature of Human and Veterinary Medicine (SNOMED), 119
T T-61 for euthanasia use, 733 Tagging and tracking active tags, 852 antennas, 852–853 cetaceans attachment methods, 863, 864, 866 branding, 862 data loggers, 863 impact of devices, 866 implantable tags, 866 passive tags, 862 roto-radio tag, 863–864
CRC Handbook of Marine Mammal Medicine
telemetry device location and attachment, 862–863 companies and addresses, 854 GPS, 857 manatees branding, 867 PITs, 866 radio telemetry use and attachment, 867–868 tagging advantages, 868, 870 passive tags, 851 pinnipeds branding, 858 flipper tags, 858 impact of devices, 862 tag location and attachment, 858–859 TDR location and attachment, 861–862 telemetry device location and attachment, 859–861 polar bear, 874 satellite telemetry, 855–857 sea otters attachment methods, 870–871 intra-abdominal radio transmitters, 871–873 PITs, 870 systems ranges, 853 time-depth recorders, 853 transmitters, 853 Tandem repeats, 272 Tapeworms in cetaceans, 362 TBG (thyroid binding globulin), 170 TCDD (tetrachlorodibenzo-p-dioxin), 489 TCPMe, 486 TCPMeOH, 486 TDRs (time-depth recorders), 853 Teeth, see also Dental system age estimation use, 455–456 anesthesia for extractions, 657 polar bears, 995, 1003 sea otters, 963 walruses, 928 Telazol/medetomidine, 999 Telemetry devices cetacean tagging and tracking, 862–863 pinniped tagging and tracking, 859–861 Temperature anesthesia depth determination and, 661 for pinnipeds in captivity, 908 ranges for marine mammals, 780 Terbinafin, 351 Terbutaline, 670 Testes pinnipeds, 200–201 position of, 148 Testosterone in bottlenose dolphins, 216 in pinnipeds, 201 Tetracaine, 657 Tetrachlorodibenzo-p-dioxin (TCDD), 489
0839_frame_IDX Page 1059 Friday, May 25, 2001 9:37 AM
1059
Index
Tetracycline adverse effects, 706 drug dosage cetaceans, 713 pinnipeds, 717 polar bears, 721 sea otters, 721 sirenians, 718 seal finger treatment, 773 Theophylline, 717 Theragra chalcogramma (walleye pollock), 6 Thermal imaging cameras, 645 clinical applications cetaceans, 647–649 general techniques, 645–646 manatees, 646 pinnipeds, 646 polar bear, 649 history, 644–645 techniques, 643 Web sites, 650 Thermal insult to pinniped testes, 201 Thermography, see Thermal imaging Thermoneutral zone (TNZ), 794–796 Thiacetarsamide, 717 Thiamine nutritional deficiency, 813–814, 922 polar bear drug dosage, 721 sea otter drug dosage, 721 sirenian drug dosage, 718 Thiamine HCL cetacean drug dosage, 713 pinniped drug dosage, 717 Thiopental for cetaceans, 660 for odobenids, 677 Thoracic imaging radiograph diagnostic imaging, 571 sonography clinical applications, 596, 600–601 Thoracic lymphadenopathy, 601 Thoracoscopy, 621 Thrombocytopenia, 707 Thrombosis, 536 Thymus gross anatomy, 143 microscopic anatomy, 145 Thyroid binding globulin (TBG), 170 Thyroid gland (thyroid hormone (TH)) circulating concentrations of thyroxine, 171–172 concentrations of fT4, 175–176 fluctuations in TH levels, 176–177 gross anatomy, 143 hormones stress response components, 260–261 hormone storage, 170 microscopic anatomy, 145 polar bear hormones, 993 protein binding, 170
reverse T3, 174, 176 thyroxine levels, 170, 174 triiodothyronine concentrations, 173–174 Thyroid-stimulating hormone (TSH), 169, 722 Thyrotropin-releasing hormone (TRH), 200 Thyroxine, 170, 722; see also Thyroid gland Tiletamine HCL, 998–999 Time-depth recorders (TDRs) cetacean tagging and tracking, 863 pinniped tagging and tracking, 861–862 tagging and tracking, 853 Tissue sample collection and handling, 167 Title IV, MMPA purpose of, 70, 71 results, 78 TLI (trypsin-like immunoreactivity), 405 TNZ (thermoneutral zone), 794–796 Toothed whales, see Cetaceans; Whales Total carbon dioxide, 417–418 Total immunoglobulin test, 245 Total plasma protein (TPP), 413 Toxaphene, 485 Toxic algal blooms, 22–23; see also Red tides Toxicology biotoxins brevetoxin, 493–494 ciguatera, 496 domoic acid, 495 paralytic shellfish poisoning, 494–495 classes of toxicants, 477 contaminants’ effects on organ systems cardiovascular, 472 central nervous, 473 endocrine, 474–475 immune, 475–476 integument, 472 kidneys, 472–473 liver, 472–473 mouth, 472 reproductive, 473–474 respiratory, 472 sense organs, 472 skeleton, 474 diagnosis, 501–502 dose scaling, 499 elements arsenic, 482 cadmium, 480–481 copper, 482 lead, 481 mercury, 478–479 organotins, 481–482 selenium, 482 silver, 482 vanadium, 482 halogenated organics accumulation and variability, 482–484 endocrine function effects, 490–491
0839_frame_IDX Page 1060 Friday, May 25, 2001 9:37 AM
1060
CRC Handbook of Marine Mammal Medicine
epizootics effects, 491–493 immunocompetence effects, 491–493 octachlorostyrene, 487, 488 organochlorine pesticides and metabolites, 484–485 organochlorines effects on metabolism, 488–490 PBDEs, 487, 488 PCDDs, 486 PCDFs, 486 PCTs, 487, 488 polybrominated biphenyls, 487, 488 polychlorinated byphenyls, 485–486 polychlorinated napthalenes, 486 polychloroquaterphenyls, 486 reproduction function effects, 490–491 TCPMe, 486 TCPMeOH, 486 oil constituents, 496 contact exposure effects, 496–498 oral-dosing studies, 498 petroleum constitutents in animals, 498–499 overview, 471, 477 reference databases, 500 treatment, 499–501 Toxoplasma gondii anthropogenic origin, 5 in manatees, 372, 953 public health significance, 773–774 in sea otters, 23–24, 978 species range, 359, 360, 368 TPP (total plasma protein), 413 Transmitters for tagging and tracking, 853 Transport of marine mammals cetaceans body support techniques, 882 equipment requirements, 886–887 ground and air, 885–886, 887 improvements in, 884 removal from transport unit, 888 stretcher use, 884–885 temperature control, 882, 884 water requirements, 885 history of, 882 medical considerations, 889–890 pinnipeds, 888 polar bear, 889 regulations, 881 sea otters, 888–889, 969, 970 sirenians, 889 Trauma therapy, 695–696 non-infectious diseases anthropogenic, 530–531 interspecific, 528–530 intraspecific, 522, 524, 528 Trematodes, see also Helminths in cetaceans, 362, 365 in manatees, 373
in pinnipeds, 370, 372 removal and fixation, 358 in sea otters, 373 treatment of, 359 Treponema spp., 536 TRH (thyrotropin-releasing hormone), 200 Trichechus inunguis (Amazonian manatee), see Manatee Trichechus manatus (West Indian manatee), see Manatee Trichechus manatus latirostris, see Florida manatee Trichechus senegalensis (West African manatee), see Manatee Trichinella spp. in pinnipeds, 372 in polar bears, 374, 1002 Trichinosis, 1003 Trichophyton lesions, diagnostic methods, and treatments, 345 in walruses, 931 Trichosporon pullulans, 345 Triglycerides, 404–405 Trimethoprimsulfadiazine cetacean drug dosage, 713 pinniped drug dosage, 717 sea otter drug dosage, 721 Trocars, 638–639 Trypsin-like immunoreactivity (TLI), 405 TSH (thyroid-stimulating hormone), 169, 722 Tuberculous mycobacterial infections, 319 Tubular nephrosis, 534 Tursiops truncatus, see Bottlenose dolphin Tusk infections and trauma, walruses, 933 TwoCal HN, 694 Tylosin, 714
U UDP-glucouronyl transferase (UDP-GT), 490 UGFNAB (ultrasound-guided fine-needle aspiration biopsy), 600, 601 Ukrainian stranding networks contacts and programs, 63 Ultrasonography, see also Diagnostic imaging; Radiology clinical applications abdominal imaging, 601 biliary system, 602 body condition, 618 eyes, 616 gastrointestinal tract, 605–608 heart, 596–597 liver, 601–604 lungs, 597–600 mediastinum, 597 musculoskeletal system, 616 pancreas, 605 reproductive tract, 611–616 spleen, 604–605 thoracic imaging, 596
0839_frame_IDX Page 1061 Friday, May 25, 2001 9:37 AM
Index
thoracic lymph nodes, 600–601 urinary tract, 609–611 indications, 593–594 limitations, 594 techniques equipment and preparation, 594–595 image orientation, 595 Ultrasound-guided fine-needle aspiration biopsy (UGFNAB), 600 Uncinaria, 369, 916 United Kingdom stranding networks contacts and programs, 64–65 United States stranding networks contacts and programs, 65–66 University of Michigan School of Information and Library Sciences, 118 Urea nitrogen, 411–412 Uric acid, 420 Urinary tract cytology findings, cetaceans, 445–446 energy losses, 809–810 gross anatomy, 147 microscopic anatomy, 149 non-infectious diseases, 534–535 sonography clinical applications, 609–611 urinalysis, 423 urine sample collection, manatees, 947 urine sample collection, pinnipeds, 910–911 urine sample collection and handling, 166–167 Urogenital system diseases of, 327 helminth parasites in cetaceans, 366 pinniped diseases, 919–920 Ursidae, see Polar bear Ursus maritimus, see Polar bear US Code (USC), 742 U.S. Department of Agriculture (USDA) sample collection and handling and, 311 transport regulations, 881 Web site, 118 U.S. Environmental Protection Agency (EPA), 785 U.S. Fish and Wildlife Service (FWS) addresses, 751–752 jurisdiction, 742 scope of, 450, 761–762 Web site, 119, 745 U.S. National Library of Medicine, 118, 119 Uterus, 147, 616 UV sterilization, 784
V Vaccinations, 317, 318 Valvular endocarditis, 536 Vanadium, 482 Vancomycin, 714 Vaquita (Phocoena sinus), 313 Vasoactive intestinal polypeptide, 200
1061
Vasopressin, 169, 183–184 Vertebrae, 154–155, 156 Vertebral column, 154 VHF frequency bands, 852 VHF radio telemetry, 863 Vibrio in manatees, 951 public health significance, 769–770 in sea otters, 971 septicemia and, 312 Vibriosis, 314–315 Video monitors/recorders in endoscopy, 630 Video systems, underwater, 857 Viral diseases and infections about, 19 adenoviruses, 291–292 caliciviruses, 300–302 coronavirus, 302 hepadnavirus, 302 herpesviruses clinical signs, 293–294 diagnosis, 294–295 differentials, 295 epidemiology, 295 host range, 292–293 pathology, 294 public health significance, 295 therapy, 294 virology, 293 influenza, 298–300 morbillivirus in cetaceans, 16, 18–19 description and treatment, 296–298 lack of connection to organochlorines, 492–493 in manatees, 952 in polar bears, 1001 public health significance, 769 serodiagnostics of, 244 in walruses, 935 papillomavirus, 289–291 in polar bears, 1001 poxvirus clinical signs, 287 diagnosis, 288 differential, 288–289 epidemiology, 289 host range, 286 pathology, 287–288 public health significance, 289 therapy, 287 public health significance calicivirus, 768–769 influenza, 769 poxvirus, 289, 768 rabies, 769 retrovirus, 302–303 rhabdovirus, 303 virus isolation, 285–286
0839_frame_IDX Page 1062 Friday, May 25, 2001 9:37 AM
1062
Virology gross necropsy, 462 Vitamin A content in polar bears, 991, 1000 drug dosage pinnipeds, 717 polar bears, 722 Vitamin B, 721 Vitamin C, see Ascorbic acid Vitamin deficiencies, 815–816 Vitamin E cetacean drug dosage, 714 pinniped drug dosage, 718 polar bear drug dosage, 722 sea otter drug dosage, 721 sirenian drug dosage, 719 Voriconazole, 351
W Walleye pollock (Theragra chalcogramma), 6 Walrus (Odobenus rosmarus) analgesics use, 699 bacterial diseases in, 314 biology, 927 clinical pathology blood collection, 385 hematology and biochemistry values, 394–395 diagnostic techniques, 930–931 diet, 929 estrous cycle, 195 FWS jurisdiction, 742 hand-rearing and artificial milk formulas, 841–842 integument microanatomy, 140 lactation energy requirements, 806 medical problems bacterial, yeasts, and fungal isolates, 932 dermatological, 931–932 foreign bodies, 934 intestinal disease, 934–935 miscellaneous diseases, 935 ophthalmology, 932–933 tusk infections and trauma, 933 parasites of, 372 physical examination, 929–930 reproduction, 928 reproductive characteristics, 196 respiratory emergencies, 695 restraint, 930 sedation and general anesthesia, 930 specimen examination, 930–931 thermal imaging applications, 646 viral diseases in caliciviruses, 300 herpesviruses, 293 Water and environment quality contaminants’ effects on organ systems, 472, 473, 475
CRC Handbook of Marine Mammal Medicine
contamination by feral and domestic cat populations, 24 environmental considerations, 779–781 filtration methods biological, 781–782 flocculation, 782 foam fractionation, 783 mechanical, 782 microorganisms as indicators, 783–784 pool size for pinnipeds, 907 sterilization methods, 784–785 ozone use, 785–786 WBCs (white blood cells), see Leukocytes evaluation Web sites Argos, 855 for career start advanced training programs, 111–112 fellowships, 112 graduate degree programs, 109–110 internships and residencies, 107–109 related programs, 110–111 scientific societies and membership organizations, 112–113 CITES, 760 CT scanning information, 558 federal regulations, 742 FWS, 745, 753 MRI applications, 560 NASA, 857 NCDC, 5 NMFS, 745, 753 parasite removal and fixation advice, 358 thermal imaging, 650 toxicology reference databases, 500 Weddell seal (Leptonychotes weddellii) adrenal glands studies, 178 bacterial diseases in, 313 cortisol concentrations, 181 cortisol levels, 178 molt energy requirements, 807 non-infectious diseases cardiovascular system, 535 genitourinary system, 534 osmoregulatory hormones, 183–184 viral diseases in, 292–293 West African manatee (Trichechus senegalensis), see Manatee West Indian manatee (Trichechus manatus), see Manatee WGMMUME (Working Group on Marine Mammal Unusual Mortality Events), 71, 450 Whale lice, 367 Whale Net Web site, 121 Whales, see also Cetaceans; specific species baleen whales age estimation, 455–456 mercury concentration in liver, 478 parasites of, 366
0839_frame_IDX Page 1063 Friday, May 25, 2001 9:37 AM
1063
Index
PCB toxicity and, 486 sexual dimorphisms, 157 trematode treatment, 365 digestive system, 145, 148–149 organochlorine effects on, 489 organohalogens accumulation in, 487 sentinel role, 4 trematode treatment, 365 White-beaked dolphin (Lagenorhynchus albirostris) bacterial diseases in, 317 viral diseases in morbillivirus, 296 rhabdoviruses, 303 White blood cells (WBCs), see Leukocytes evaluation White-sided dolphin, see Atlantic white-sided dolphin Wildlife Health Web site, 120 Working Group on Marine Mammal Unusual Mortality Events (WGMMUME), 71, 450 Wound management, 696
X Xylazine euthanasia use, 732 for otariids, 670 Xylazine-ketamine, 666
Y Yeasts in urine, 446 Yohimbine, 670 Yunan paiyio, 692
Z Zalophus californianus, see California sea lion Zalophus californianus wollebaeki (Galapagos sea lion), 326 Zinc chlorhexidate gel, 721 Zolazepam, 670 Zolazepam HCL, 998–999 Zolazepam-tiletamine (ZT) for odobenids, 677, 680 for otariids, 666 for phocids, 674 Zygomycete infections clinical diagnosis, 349 lesions, diagnostic methods, and treatments, 345–347 mechanisms of pathogenesis, 339 species affected, 338
0839_frame_IDX Page 1064 Friday, May 25, 2001 9:37 AM
