Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, 3rd Edition

FE R R E T S, R A B B I T S, and R O D E N T S C l i nical Medicine and Surgery T H I R D

E D I T I O N

Katherine E. Quesenberry DVM, MPH, Diplomate ABVP (Avian) Service Head Avian and Exotic Pet Service The Animal Medical Center New York, New York

James W. Carpenter MS, DVM, Diplomate ACZM Professor of Zoological Medicine Department of Clinical Sciences College of Veterinary Medicine Kansas State University Manhattan, Kansas

3251 Riverport Lane St. Louis, Missouri 63043

FERRETS, RABBITS, AND RODENTS: CLINICAL MEDICINE AND SURGERY Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

ISBN: 978-1-4160-6621-7

Some material was previously published. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Images in Chapter 34 © Stephen J. Divers. Notice Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods, they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. With respect to any drug or pharmaceutical products identified, readers are advised to check the most current information provided (i) on procedures featured or (ii) by the manufacturer of each product to be administered, to verify the recommended dose or formula, the method and duration of administration, and contraindications. It is the responsibility of practitioners, relying on their own experience and knowledge of their patients, to make diagnoses, to determine dosages and the best treatment for each individual patient, and to take all appropriate safety precautions. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Previous editions copyrighted 2004 and 1997. Library of Congress Cataloging-in-Publication Data Ferrets, rabbits, and rodents : clinical medicine and surgery / [edited by] Katherine E. Quesenberry, James W. Carpenter. — 3rd ed. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4160-6621-7 (pbk. : alk. paper) I. Quesenberry, Katherine E. II. Carpenter, James W. (James Wyman), 1945[DNLM: 1. Ferrets—surgery. 2. Rabbits—surgery. 3. Rodentia—surgery. 4. Surgical Procedures, Operative—veterinary. SF 997.3] 636.932’2—dc23 2011039100 Vice President and Publisher: Linda Duncan Publisher, Veterinary Medicine: Penny Rudolph Associate Developmental Editor: Brandi Graham Publishing Services Manager: Catherine Jackson Senior Project Manager: Carol O’Connell Design Direction: Paula Catalano Printed in the United States of America Last digit is the print number:  9  8  7  6  5  4  3  2  1

Contributors Sean Aiken, DVM, MS, Diplomate ACVS Veterinary Specialty Hospital San Diego, California Natalie Antinoff, DVM, Diplomate ABVP (Avian) Gulf Coast Veterinary Specialists Avian and Exotics Houston, Texas Heather W. Barron, DVM, Diplomate ABVP (Avian) Professor and Chair Department of Veterinary Clinical Sciences School of Veterinary Medicine St. Matthew’s University Grand Cayman, Cayman Islands, British West Indies Louise Bauck, DVM, MVSc Professor of Biology Department of Math and Science Brenau University Gainesville, Georgia Teresa Bradley Bays, DVM, CVA Director Belton Animal Clinic and Exotic Care Center Animal Urgent Care of Cass County Belton, Missouri Judith A. Bell, DVM, PhD Department of Population Medicine Ontario Veterinary College University of Guelph Guelph, Ontario, Canada R. Avery Bennett, DVM, MS, Diplomate ACVS Chief of Surgery The Animal Medical Center New York, New York Cynthia R. Bishop, DVM Assistant Professor Department of Veterinary Clinical Sciences Seattle Pacific University Seattle, Washington Cynthia Brown, DVM, Diplomate ABVP (Avian) Avian and Exotic Medicine New England Veterinary Medical Center Mystic, Connecticut Susan A. Brown, DVM Rosehaven Exotic Animal Veterinary Service North Aurora, Illinois Michelle L. Campbell-Ward, BSc, BVSc (Hons I), DZooMed, MRCVS Taronga Western Plains Zoo Dubbo, NSW, Australia

Vittorio Capello, DVM, Diplomate ABVP (Exotic Companion Mammal), Diplomate ECZM (Small Mammal) Exotic Companion Mammal Medicine and Surgery Clinica Veterinaria S.Siro Clinica Veterinaria Gran Sasso Milano, Italy Stephen J. Divers, BSc (Hons), BVetMed, Diplomate ACZM, Diplomate ECZM (Herpetology), Diplomate ZooMed, FRCVS Professor of Zoological Medicine Department of Small Animal Medicine & Surgery College of Veterinary Medicine University of Georgia Athens, Georgia Thomas M. Donnelly, BVSc, Diplomate ACLAM Warren Institute Ossining, New York Peter G. Fisher, DVM Director Pet Care Veterinary Hospital Virginia Beach, Virginia Anthony J. Fischetti, DVM, MS, Diplomate ACVR Department Head of Diagnostic Imaging The Animal Medical Center New York, New York James G. Fox, DVM, MS, Diplomate ACLAM Professor and Director Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, Massachusetts Carley J. Giovanella, DVM, Diplomate ACVIM (Neurology) Gulf Coast Veterinary Neurology and Neurosurgery Houston, Texas Jennifer Graham, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal), Diplomate ACZM Avian and Exotic Medicine Angell Animal Medical Center Boston, Massachusetts Michelle G. Hawkins, VMD, Diplomate ABVP (Avian) Associate Professor, Companion Avian and Exotic Pet Medicine Department of Medicine and Epidemiology School of Veterinary Medicine University of California, Davis Davis, California Laurie Hess, DVM, Diplomate ABVP (Avian) Veterinary Center for Birds and Exotics Bedford Hills, New York

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CONTRIBUTORS

Heidi L. Hoefer, DVM, Diplomate ABVP (Avian) Island Exotic Veterinary Care Huntington Station, New York; Consultant and Adjunct Clinician Exotics Emergency and Critical Care Long Island Veterinary Specialists Plainview, New York Sharon M. Huston, DVM, Diplomate ACVIM (Cardiology) San Diego Veterinary Cardiology San Diego, California Evelyn Ivey, DVM, Diplomate ABVP (Avian) Four Corners Veterinary Hospital Concord, California Jeffrey R. Jenkins, DVM, Diplomate ABVP (Avian) Avian and Exotic Animal Hospital San Diego, California

Marla Lichtenberger, DVM, Diplomate ACVECC Department of Emergency and Critical Care Milwaukee Emergency Center for Animals Greenfield, Wisconsin Teresa Lightfoot, DVM, Diplomate ABVP (Avian) Avian and Exotic Service BluePearl Veterinary Partners Tampa, Florida Andrew S. Loar, DVM, Diplomate ACVIM ALX Laboratories The Animal Medical Center New York, New York Lori Ludwig, VMD, MS, Diplomate ACVS Veterinary Surgical Care, LLC Mt. Pleasant, South Carolina

Cathy A. Johnson-Delaney, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal) Avian and Exotic Animal Medicine Center Kirkland, Washington

Douglas R. Mader, MS, DVM, Diplomate ABVP (Canine and Feline) Marathon Veterinary Hospital Marathon Sea Turtle Hospital Conch Republic

Amy S. Kapatkin, DVM, MS, Diplomate ACVS Associate Professor of Orthopedic Surgery Department of Surgical and Radiological Sciences College of Veterinary Medicine University of California–Davis Davis, California

Christoph Mans, MedVet Special Species Health Service Department of Surgical Sciences School of Veterinary Medicine University of Wisconsin Madison, Wisconsin

Eric Klaphake, DVM, Diplomate ACZM, Diplomate ABVP (Avian) Animal Medical Center Bozeman, Montana

Mark A. Mitchell, DVM, MS, PhD, Diplomate ECZM (Herpetology) Professor, Zoological Medicine College of Veterinary Medicine University of Illinois Urbana, Illinois

Marc S. Kraus, DVM, Diplomate ACVIM (Cardiology, Internal Medicine) Senior Lecturer Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, New York Pamela Ming-Show Lee, DVM, MS Cardiology Service The Animal Medical Center New York, New York Angela M. Lennox, DVM, Diplomate ABVP (Avian) Avian and Exotic Animal Clinic Indianapolis, Indiana; Adjunct Assistant Professor Department of Veterinary Clinical Sciences School of Veterinary Medicine Purdue University West Lafayette, Indiana

James K. Morrisey, DVM, Diplomate ABVP (Avian) Senior Lecturer Exotic and Wildlife Medicine Department of Clinical Sciences College of Veterinary Medicine Cornell University Ithaca, New York Robert D. Ness, DVM Ness Exotic Wellness Center Lisle, Illinois Barbara L. Oglesbee, DVM, Diplomate ABVP (Avian) Clinical Associate Professor Department of Veterinary Clinical Science College of Veterinary Medicine The Ohio State University Capital Veterinary Referral and Emergency Center Columbus, Ohio

CONTRIBUTORS Connie Orcutt, DVM, Diplomate ABVP (Avian, Exotic Companion Mammals) Avian and Exotic Animal Medicine Putnam Veterinary Clinic Topsfield, Massachusetts Peter J. Pascoe, BVSc, Diplomate ACVA, DVA, Diplomate ECVAA Professor Department of Surgical and Radiological Sciences School of Veterinary Medicine University of California, Davis Davis, California Joanne Paul-Murphy, DVM, Diplomate ACZM Professor Department of Companion Avian and Exotic Pets School of Veterinary Medicine University of California, Davis Davis, California Anthony A. Pilny, DVM, Diplomate ABVP (Avian) Avian and Exotic Pet Medicine Veterinary Internal Medicine and Allergy Specialists New York, New York Christal G. Pollock, DVM, Diplomate ABVP (Avian) Veterinary Consultant Lafeber Company Cleveland, Ohio Lauren V. Powers, DVM, Diplomate ABVP (Avian) Avian and Exotic Pet Service Carolina Veterinary Specialists Huntersville, North Carolina Karen L. Rosenthal, DVM, MS Associate Professor of Special Species Medicine School of Veterinary Medicine University of Pennsylvania Philadelphia, Pennsylvania Jonathan Rubinstein, DVM Avian and Exotic Service BluePearl Veterinary Partners Tampa, Florida Andrea Siegel, DVM ALX Laboratories The Animal Medical Center New York, New York Kathy Tater, DVM, Diplomate ACVD Master’s of Public Health Candidate in Quantitative Methods Harvard School of Public Health; Clinical Assistant Professor Cummings School of Veterinary Medicine Tufts University North Grafton, Massachusetts

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Thomas N. Tully, Jr., DVM, MS, Diplomate ABVP (Avian), Diplomate ECZM (Avian) Professor of Zoological Medicine Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University Baton Rouge, Louisiana Alexandra van der Woerdt, DVM, MS, Diplomate ACVO, Diplomate ECVO Staff Ophthalmologist The Animal Medical Center New York, New York David Vella, BSc, BVSc (Hons), Diplomate ABVP (Exotic Companion Mammal) North Shore Veterinary Specialist Centre; Animal Referral Hospital Sydney, New South Wales, Australia James Walberg, DVM, Diplomate ACVP Consultant Department of Pathology The Animal Medical Center New York, New York Bruce H. Williams, DVM, Diplomate ACVP Senior Pathologist Veterinary Pathology Service Joint Pathology Center Washington, D.C. Nicole R. Wyre, DVM, Diplomate ABVP (Avian) Chief, Special Species Section Matthew J. Ryan Veterinary Hospital School of Veterinary Medicine University of Pennsylvania Philadelphia, Pennsylvania Ashley Zehnder, DVM, Diplomate ABVP (Avian) Post Doctoral Scholar Department of Comparative Medicine Stanford University Stanford, California

I dedicate this book to all of my friends and very close colleagues who have supported me through some very difficult times in these last 5 years. Without their encouragement and support, pulling this book together in the face of both personal and professional challenges would not have been possible. In particular, I thank Connie Orcutt, Laurie Hess, Tom Donnelly, Heidi Hoefer, Susan Orosz, and my co-editor, Jim Carpenter, all of whom I have worked with and learned from for many years in this profession. I thank my sister, Marcia Quesenberry, for her unwavering support, encouragement, and love. I also give special thanks and love to my children, Zachary and Chelsea Messinger, who are always at the center of my life and being. Katherine E. Quesenberry I wish to acknowledge all those who have contributed to our knowledge and understanding of small mammal medicine; Dr. Kathy Quesenberry for graciously cajoling me into collaborating on yet another edition of the Pink Book; and the many colleagues, interns and residents, and students who have inspired my professional life. I also wish to thank veterinary students Caitlin Burrell, Richard Brooksby, and Amy Guersey for their “office assistance,” and especially Dr. Chris Marion for his editorial assistance in the preparation of this edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery. I would like to dedicate this book to my family (wife, Terry; son, Michael; and daughter, Erin, and her family–husband, Steve, and kids, Kylie, Hayden, and Asher) who have supported me as I pursue my passion for zoological medicine. James W. Carpenter

Preface In the 15 years since the first edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, the specialty of exotic pet mammal medicine and surgery has become more mainstream in veterinary practice, and, as a matter of fact, has become an integral component of most small animal clinical practices. The knowledge base has expanded tremendously as interest in small mammals has prompted research in these animals not as just as laboratory species, but as companion animals. As information about these species has become much more accessible via Internet websites and chat groups, the public has increasingly recognized that these species are valued as pets and deserving of high quality veterinary care. Many pet owners look beyond the financial value of these animals and expect state of the art veterinary care at the same level as that given to dogs and cats. The numbers of small mammals, particularly pet rabbits, chinchillas, and guinea pigs, presented to veterinarians for health care has grown steadily, as a consequence of more accessible information about their care to the public as well as the increase in spending on companion animal care in general. As veterinarians, we therefore must be able to provide a high level of medical and surgical care, based on a solid knowledge base, for these pets. The number of books, publications, and websites that are now devoted to the husbandry and veterinary care of small pet mammals is enormous. Whereas previously only a few veterinary texts were published about these species, now there are many books, serial publications, and journal articles available on various topics ranging from medicine, surgery, imaging, and clinical techniques to behavior. Some of this work is original, and some works only present the same material reworded into different formats.

With this third edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, we have tried to stay true to our original mission of publishing up-to-date information in a reader-friendly, comprehensive yet concise format. All of the chapters have been updated, with new authors on many of the topics and new chapters added on “Emergency and Critical Care” and “Behavior.” As the information about these species has exploded, we have tried to focus on the most pertinent and reliable information to present to our readers. Our authors are among the most respected in veterinary medicine and encompass a broad range of specialists in exotic pet mammals, internal medicine, surgery, critical care, and laboratory animal medicine. We are proud to present this third edition of Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery to our colleagues and readers, and we are especially pleased that this edition is in color. We are very grateful to the chapter authors and to our team at Elsevier, especially Penny Rudolph, Brandi Graham, and Carol O’Connell, for their patience and very hard work in bringing this publication together. We are confident that the format, presentation, information, and reliability of the “Pink Book” will continue to set it apart as the standard in this subspecialty of veterinary medicine.

Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian)

James W. Carpenter, MS, DVM, Diplomate ACZM

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SECTION ONE

Ferrets

CHAPTER

1

Basic Anatomy, Physiology, and Husbandry

 

Lauren V. Powers, DVM, Diplomate ABVP (Avian), and Susan A. Brown, DVM

Domestication History Uses Anatomy and Physiology Integument Gastrointestinal System Urogenital System Cardiovascular and Lymphatic Systems Respiratory System Endocrine System Musculoskeletal System Neurologic System and Special Senses Physiology and Reproduction Physiology Body Size and Seasonal Weight Variation Reproduction Husbandry Housing Environmental Enrichment Nutrition

DOMESTICATION HISTORY Ferrets belong to the family Mustelidae and are related to weasels, mink, otters, badgers, stoats, and martens. There are currently three living species of ferrets (also known as polecats in Europe and Asia): the European polecat (Mustela putorius), the Steppe or Siberian polecat (Mustela eversmanni), and the blackfooted ferret (Mustela nigripes). All three species live primarily solitary social lives and are very efficient hunters supporting Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

their obligate carnivore lifestyle. The European polecat is found in various areas from the Atlantic to the Ural mountains and dwells along the edges of woodlands and wetlands.12 The Siberian polecat is found in Eurasia from the thirtieth to the sixtieth degree of latitude, may be larger than the European polecat, and lives primarily in open areas such as steppes, slopes of ravines, and semi-deserts.12 The black-footed ferret is native to the prairies of North America. It almost became extinct in the wild because of habitat destruction and the decimation of its main food source, the prairie dog, from poisoning and hunting.12 Currently, captive breeding and reintroduction programs are under way in an attempt to reestablish the black-footed ferret into its native range. It is illegal to own this endangered species. The origin of the domestic ferret, which is traditionally referred to as Mustela putorius furo, is shrouded in mystery. The Latin name translates loosely as “mouse-eating (mustela) smelly (putorius) thief (furo).” Currently there is a move toward using nomenclature that differentiates the wild progenitors of a domesticated species from the domesticate, and some mammologists are moving toward referring to the domesticated ferret as Mustela furo.5,11 The domesticated ferret may have originated from either the Siberian or the European polecat, or possibly both.5,8,33 It is difficult to find archaeological evidence of domestication, possibly because of the ferret’s small skeleton, which may have deteriorated rapidly or was indistinguishable from wild ferrets living in the environment, or the lack of paraphernalia associated with the ferret, making them archaeologically unimportant.5 Ferrets have been domesticated for approximately 2,000 to 3,000 years.5 The first clear reference to domestication is in the writings of the Spaniard Isidore of Seville in 622 ad.5 There is a high probability that ferrets were brought by Romans or Normans during their invasions, but there are currently no references that irrefutably link Romans and Greeks with domestication of ferrets.5 It is likely that ferrets 1

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SECTION I  Ferrets

were first domesticated over a wide area somewhere in the south to southeastern portion of Europe near the Mediterranean.5 Over the centuries, numerous references have been made to the use of ferrets in Europe for rodent control in homes, farms, and ships and for hunting rabbits both for damage control and for human food, as well as for their pelts. The domestic ferret was introduced into Australia from Europe in the 1800s to control the populations of European rabbits that had been previously released.19 Fortunately, enough other predators, such as foxes, dingoes, and hawks, preyed on the ferret so that feral populations never developed.19 However, when they were introduced into New Zealand for the same reason in the late 1800s along with stoats and weasels, there were no predators to control their numbers.19 Feral populations of domestic ferrets therefore developed and are still present today.8,19 The impact of feral ferrets on native wildlife has been controversial. The domestic ferret was probably introduced into the United States from Europe by the shipping industry in the 1700s. They may have come as pets or as hunting companions.8,19 Ferrets were also used for their pelts and the town of New London, Ohio, became known as Ferretville because of its huge breeding population of ferrets around 1915.19 At the turn of the twentieth century, hunting with ferrets was banned in some states to protect against destruction of the native rabbit population.19 The pet ferret has changed from its wild progenitor in the process of domestication in the areas of physiology, reproduction, and behavior. Reproductively, ferrets mate two or more times a year compared to polecats at generally once a year.5 The ferret’s litter size average of eight is larger than the polecat’s at six.5 Ferret coat color has changed, as is the case with most animals that are domesticated. Albino ferrets have been bred for centuries and are often preferred for hunting because of their high visibility. Albino ferrets may also have alterations in their vision and hearing.5 Other changes include a 15% to 20% smaller cranial capacity, a wider postorbital constriction, and dental crowding.5 Behaviorally, ferrets appear to be more gregarious than their wild counterparts, but this may be due to juvenilization, which is a common side effect of domestication that allows ferrets to live in “litter groups” rather than their more naturally solitary lifestyle.5 The most noticeable behavioral change is a loss of the innate fear of humans, as well as a lack of fear toward unfamiliar objects in their environment.5

USES Historically, humans have not domesticated animals primarily for the purpose of companionship. Animals needed to serve an economic purpose and the same holds true for the ferret. Early references to ferrets record their use for rodent or rabbit control.8,19 Ferrets are efficient little predators that can bring down prey quite a bit larger than themselves and can maneuver in small spaces more effectively than cats. Ferrets were used on ships in colonial days to control the rat populations.8 In the early 1900s, the U.S. Department of Agriculture encouraged the use of ferrets as a means of controlling rabbits, raccoons, gophers, mice, and rats around granaries and farms.8 One needed only to call the local “ferret master” to bring out his ferrets, which were set loose to do their work and then recaptured to work another day. Large facilities kept their own ferrets on site. Ferrets are still used for rodent and rabbit control in some areas of Europe and Australia today. However, hunting with ferrets is prohibited in the United States.

Ferrets have long been used to hunt rabbits—not only for control, but as a food source for human beings. “Ferreting” was a common sport in the United Kingdom and many other areas of Europe. It is still practiced today but to a much lesser degree. Ferrets are released in a rabbit warren area, where they investigate burrows and flush out rabbits. The rabbits are then caught in nets or by dogs or shot by the waiting hunter as they exit their burrows. Domestic ferrets have been bred for their pelts. A coat made of ferret fur is referred to as fitch. Ferret hair has also been used in other products such as artist’s brushes.19 Ferret fur never really took hold in the United States, but it still exists in a few areas of Northern Europe. An entertainment peculiar to English pubs and still found in a few isolated areas of the United Kingdom is called ferret-­ legging. This is a sport in which a man securely ties his trouser legs closed at the ankles and then places two ferrets, each with a full set of teeth, into his trousers. He then securely ties the trousers closed at the waist. The contest is to see how long he can stand having the ferrets in his trousers. If a ferret bites, it can only be dislodged from the outside of the trousers. The record of 5 hours and 26 minutes was set by a 72-year-old Yorkshire man.8 Domestic ferrets have also been used to transport cables through long stretches of conduit. They have been used to string cable for oilmen of the North Sea, for camera crews, in jets, and for telephone companies.8 Ferrets have been used in biomedical research since the early 1900s, when they were used to study human influenza and other viral diseases.8 Today ferrets are used in the fields of virology, reproductive physiology, anatomy, endocrinology, and toxicology.8 Although the use of ferrets in research is very distasteful to some, much of the information gained has directly benefited the pet ferret as well. The main use for ferrets today, however, particularly in the United States, is as a companion animal. Their popularity has increased dramatically over the past few decades. There has been a proliferation of ferret organizations dedicated to the wellbeing of this pet. It is difficult to say when the first ferret was kept strictly as a pet, but it is hard to imagine people in the distant past not feeling some attraction to the engaging personality of this animal. Ferrets make suitable pets for many people. They are small, clean, and very interactive with human beings and each other. However, as with all companion animals, the prospective owner should be educated on their husbandry requirements and behavior. For instance, ferrets (as with most pets) are not suitable for children younger than 6 years. Another consideration is that the majority of ferrets in the United States will likely be afflicted by one or more neoplastic diseases as they age. In addition, certain legal restrictions relate to the ownership of ferrets. Ferrets are still not considered domestic animals in most areas of the United States despite their long history of domestication. In some areas, owning a ferret as a pet is illegal, and in other areas permits must be obtained for ownership. With the advent of an approved rabies vaccine for the domestic ferret, restrictions on their use as pets have been lifted in many parts of the United States. However, in some localities, even if the ferret is appropriately vaccinated, it can be seized and destroyed if it bites a human being. Veterinarians should therefore be familiar with legislation not only in their state, but in their specific county or city regarding the keeping of ferrets before they engage in ferret veterinary care.

CHAPTER 1  Basic Anatomy, Physiology, and Husbandry

ANATOMY AND PHYSIOLOGY The basic body plan of the domestic ferret is similar to that of other carnivores. The following is a brief review of the clinically relevant anatomic and physiologic features of the ferret. Skeletal anatomy is depicted in Figure 1-1, visceral anatomy in Figure 1-2, and normal radiographic anatomy in Figures 1-3 and 1-4. For radiographic views of select pathologic conditions, anatomy, see Chapter 35. Selected physiologic values are detailed in Table 1-1. The reader is directed to publications containing more extensive reviews of ferret anatomy and physiology.7,9,10,17,31

INTEGUMENT Coat The domestic ferret possesses a fine undercoat and coarse, long guard hairs that provide excellent insulation.7 There are no specific breeds of ferrets, but color and pattern standards exist (Fig. 1-5). The American Ferret Association recognizes the following color standards for the purposes of show and breeding—albino, black, black sable, champagne, chocolate, dark-eyed white, and sable. Recognized pattern standards for these colors include solid, standard, color point (Siamese), blaze, panda, roan, and dark-eyed white. Mask configuration can change from season to season and from year to year, making photography an unreliable method of individual identification. Ferrets living outdoors tend to be darker in color.20 Ferrets undergo a heavy shed in the spring and the fall as seasonal weight changes occur. The coat may be shorter in summer months and longer in the fall, and lighter in color in the winter and darker in the fall. Sexually altered ferrets of either gender

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have a less dramatic molt and color change than intact animals. Clients should be warned that fur shaved for a procedure may not be replaced for weeks to months, and that the fur may initially discolor the skin a bluish hue before erupting and may have a different color or texture than surrounding fur.

Skin and Associated Glands The thick skin and muscle found on the neck and shoulders of a ferret protect it from trauma during fighting and mating. Ferrets have very active sebaceous glands, which account for their strong musky odor.14 During the breeding season, intact animals have increased sebaceous secretions; this increase results in a noticeable increase in odor, yellow to orange discoloration of the undercoat, and oily fur.14 Ferrets lack sweat glands, and in part for this reason they are very susceptible to heat prostration.14,15,25

Anal Glands Ferrets possess a pair of well-developed anal glands, as do all mustelids. These glands produce a serous yellow liquid with a powerful odor. Ferrets that are frightened or threatened can express their anal glands but, unlike skunks, are unable to project the fluid over long distances.14,15 The anal gland ducts are located at about 4 o’clock and 8 o’clock and open into the anal canal. The glands are typically about 10 mm × 5 mm in size.20 Striated external anal sphincter muscle encloses the duct of each anal sac.7,15,20 Ferrets raised at large commercial breeding facilities in the United States are routinely descented between 5 and 6 weeks of age. This is despite the fact that the majority of the odor from a ferret arises from the sebaceous glands.14

Fig. 1-1  Skeletal anatomy of a ferret. 1, Calvaria; 2, hyoid apparatus; 3, larynx; 4, seven cervical vertebrae; 5, clavicle; 6, scapula; 7, 15 thoracic vertebrae; 8, five lumbar vertebrae; 9, three sacral vertebrae; 10, 18 caudal vertebrae; 11, first rib; 12, manubrium; 13, sternum; 14, xiphoid process; 15, humerus; 16, radius; 17, ulna; 18, carpal bones; 19, accessory carpal bone; 20, metacarpal bones; 21, ilium; 22, ischium; 23, pubis; 24, femur; 25, patella; 26, fabella; 27, tibia; 28, fibula; 29, tarsal bones; 30, calcaneus; 31, metatarsal bones; 32, talus; 33, os penis. (Adapted from An NQ, Evans HE. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:14.)

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SECTION I  Ferrets

Fig. 1-2, see legend on opposite page.

CHAPTER 1  Basic Anatomy, Physiology, and Husbandry

5

Fig. 1-2  A, Ventral aspect of the viscera of a ferret in situ. B, Anatomy of the viscera and most important blood vessels as seen after removal of the lungs, liver, and gastrointestinal tract. 1, Larynx; 2, trachea; 3, right cranial lobe of lung; 4, left cranial lobe of lung; 5, right middle lobe of lung; 6, right caudal lobe of lung; 7, left caudal lobe of lung; 8, heart; 9, diaphragm; 10, quadrate lobe of liver; 11, right medial lobe of liver; 12, left medial lobe of liver; 13, left lateral lobe of liver; 14, right lateral lobe of liver; 15, stomach; 16, right kidney; 17, spleen; 18, pancreas; 19, duodenum; 20, transverse colon; 21, jejunoileum; 22, descending colon; 23, uterus; 24, ureter; 25, urinary bladder; 26, right common carotid artery; 27, left common carotid artery; 28, vertebral artery; 29, costocervical artery; 30, superficial cervical artery; 31, axillary artery; 32, right subclavian artery; 33, right internal thoracic artery; 34, left internal thoracic artery; 35, branch to thymus; 36, left subclavian artery; 37, brachiocephalic (innominate) artery; 38, cranial vena cava; 39, aortic arch; 40, right atrium; 41, pulmonary trunk; 42, left atrium; 43, right ventricle; 44, left ventricle; 45, caudal vena cava; 46, aorta; 47, esophagus; 48, hepatic veins; 49, celiac artery; 50, cranial mesenteric artery; 51, left adrenolumbar vein; 52, left adrenal gland; 53, right adrenal gland; 54, left renal artery and vein; 55, left kidney; 56, suspensory ligament of ovary; 57, left ovarian artery and vein; 58, left ovary; 59, left deep circumflex iliac artery and vein; 60, caudal mesenteric artery; 61, broad ligament of uterus; 62, left external iliac artery; 63, right common iliac vein; 64, left internal iliac artery; 65, rectum. (Adapted from An NQ, Evans HE. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. Philadelphia: Lea & Febiger; 1988:14.)

1 2 3 4

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5 2

2 6

7 12

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13 9 8

10

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A

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B Fig. 1-3  A, Ventrodorsal radiograph of a 1-year-old, spayed female ferret. Note normal positioning of thoracic and abdominal viscera. B, Same radiograph as (A): 1, trachea (endotracheal tube within lumen); 2, lung; 3, cranial mediastinum; 4, left primary bronchus; 5, heart; 6, liver; 7, stomach; 8, spleen; 9, left kidney; 10, urinary bladder; 11, right primary bronchus; 12, small intestine; 13, right kidney. (Silverman S, Tell LA. Radiology of rodents, rabbits, and ferrets: an atlas of normal anatomy and positioning. St. Louis: Elsevier Saunders; 2005:233.)

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SECTION I  Ferrets

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A 1

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3

4

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10

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B 12

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8

14

Fig. 1-4  A, Lateral (right lateral recumbency) radiograph of a 1-year-old, spayed female ferret. Note normal positioning of thoracic and abdominal viscera. B, Same radiograph as (A): 1, trachea (endotracheal tube within lumen); 2, lung; 3, pulmonary vasculature; 4, bronchus; 5, pulmonary vein; 6, stomach; 7, kidney; 8, spleen; 9, colon; 10, intrathoracic adipose tissue; 11, heart; 12, liver; 13, small intestine; 14, urinary bladder. (Silverman S, Tell LA. Radiology of rodents, rabbits, and ferrets: an atlas of normal anatomy and positioning. St. Louis: Elsevier Saunders; 2005:232.)

GASTROINTESTINAL SYSTEM Teeth and Salivary Glands The ferret dentition is typical of carnivores, consisting of long curved canine teeth and resilient molars and premolars. The permanent teeth erupt between 50 and 74 days of age. There are 34 adult teeth, and the dental formula of the adult ferret is 2(I33 C11 P33 M12).14,15,18,20 The incisors and canine teeth possess a single root. The premolars have two roots each except for the upper third premolar (carnassial tooth), which has three roots. The upper molar and first lower molar have three roots, and the tiny second lower molar has only one.7 This formula differs slightly from other carnivores in that ferrets have three rather than four premolars.14,18 Supernumerary incisors are common.3,14,20 Ferrets possess five major pairs of salivary glands—the parotid, mandibular, sublingual, molar, and zygomatic glands.15,20

Esophagus, Stomach, and Intestines The muscle of the ferret esophagus is striated along the entire length cranial to the diaphragm.15 There is not a true gastroesophageal sphincter, and ferrets are able to vomit.15,18,20 However, in contrast to other carnivores, ferrets with gastrointestinal obstruction usually are not presented with a history of vomiting.20 The stomach is relatively simple, being roughly J-shaped and consisting of a cardia, body, pyloric antrum, and fundus.14,20 The stomach is separated from the liver by the lesser omentum.7 The stomach is capable of enormous distention18,20 and can easily hold 50 mL/kg or more.20 The small intestine is comparatively short, with reported lengths of 182 to 198 cm in the adult, and with a ratio of intestinal length to body length of about 5:1.7,15,18,28 This short intestinal length contributes to the comparatively short gastrointestinal transit time of 3 to 4 hours in the adult ferret.4

CHAPTER 1  Basic Anatomy, Physiology, and Husbandry

7

  Table 1-1     Selected Biologic Values for the Domestic Ferret9,14,15,20 Parameter

Gender

Value

Body weight

Intact male Intact female Neutered, both genders

1-2 kg 0.6-1.0 kg 0.8-1.2 kg

Life span Rectal temperature Heart rate Blood volume Systolic blood pressure, awake Respiratory rate Tidal volume Stomach capacity Gastrointestinal transit time Urinary output Bladder capacity Urine pH Puberty

Reproductive life span

Male Female Male Female

Male Female Male Female

Gestation Litter size Birth weight Eyes and ears open Weaning age Maintenance fluid needs Maintenance caloric needs

6-12 yr 101.8°F (range 100-104°F [37.8-40°C]) 200-400 beats/min 60 mL 40 mL 161 mm Hg 133 mm Hg 33-36 breaths/min 10-11 mL/kg 50 mL/kg when distended 2.5 to 3.6 hr (meat); liquids may reach rectum within 1 hr 1 mL/hr (range 0.33-5.8 mL/hr) Approximately 5 mL/kg (higher under increased pressure) 6.5-7.5 9 months (as early as 23 wk with photoperiod manipulation) 8-12 months (as early as 16 wk with photoperiod manipulation) Throughout life 2-5 yr 41 days (range 39-42 days) 8 kits (range 1-18 kits) 6-12 g 28-34 days 6-8 wk Unknown, estimated at 60 mL/kg/day 200-300 kcal/kg/day

indistinguishable and are coiled together as the jejunoileum.15 The ferret lacks a cecum and appendix, and therefore the ileocolic junction is grossly indistinguishable but is typically defined as the region at which the ileocolic and jejunal arteries join.15,18 The large intestine is about 10 cm in length, consisting of the colon, rectum, and anus. There is an ascending, transverse, and descending colon.7

Liver, Gallbladder, and Pancreas

Fig. 1-5  Sable coloring of a domestic ferret. (Photography subjects provided by J. Ball.) The duodenum is about 10 cm in length and consists of three portions—the shorter (2 cm) cranial portion, the descending portion (5 cm), and the ascending portion (3 cm). The mesoduodenum encloses the right limb of the pancreas and a portion of the lesser omentum.7 The jejunum and ileum are

The liver is relatively large in the ferret and consists of six lobes—left lateral, left medial, quadrate, right medial, right lateral, and caudate.7,15,18,20 The gallbladder sits in a fossa between the quadrate and right medial liver lobes and averages 2 cm in length and 1 cm in width.7,15,20 The cystic duct joins the left, right, and central hepatic ducts to form the common bile duct, although variations of this formula exist.7 The pancreas is V-shaped and divided into right and left lobes connected by a body that lies close to the pyloris and is contained within the mesoduodenum.7 The left lobe extends along the dorsal caudal stomach and medial to the spleen. The right lobe closely follows the descending duodenum. Ducts from the left and right lobes connect to form the common pancreatic duct that joins the bile duct, which opens into the duodenal lumen about 3 cm caudal to the cranial duodenal flexure.7

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SECTION I  Ferrets

UROGENITAL SYSTEM Kidneys, Ureters, and Bladder The kidneys are bean-shaped in the ferret and average 2.4 to 3.0 cm in length, 1.20 to 1.35 cm in width, and 1.10 to 1.35 in thickness.7 Both kidneys are retroperitoneal and covered by a thin peritoneal membrane. The cranial margin of the right kidney sits in a fossa of the caudate lobe of the liver.7 The ureters pass from the renal pelvis and extend caudally along the ventral aspect of the psoas muscle, entering the dorsolateral bladder just caudal to the its neck.7 The urinary bladder sits ventrally in the abdomen just cranial to the pelvic inlet. Although the bladder is small, it can easily hold 10 mL of urine at low pressure.15,31

Male Reproductive Tract The reproductive tract of the intact male (hob) resembles that of the dog, with a palpable os penis.14,15 Unlike the dog, however, the tip to the os penis is J-shaped, making urethral catherization difficult.15 The prepuce is a fold of skin reflected over the penis, containing fur externally but bare within.7 The preputial opening lies just caudal to the umbilical area on the ventral abdomen. The scrotum lies just caudal to the caudal margin of the os penis.7 The prostate gland is the single accessory reproductive gland in the male ferret. It is a fusiform structure that surrounds the proximal urethra and measures approximately 1.5 cm long and 0.6 cm across.16 Each ductus deferens opens into the urethra at the level of the prostate.7

Female Reproductive Tract The female reproductive tract of the intact female ferret (jill) closely resembles that of other carnivores. Two long, tapering uterine horns are present, as well as a short uterine body and a single cervix.7,15,29 The ovaries are paired and located just caudal to the kidneys. The ovary is attached to the wall of the abdominal cavity by the suspensory ligament cranially and to the uterine horn caudally by the proper ligament. The uterus is suspended by the broad and round ligaments. The urethra opens into the vaginal floor at the urethral orifice.7 The vulva consists of the vestibule, clitoris, and labia, and is located in the perineum ventral to the anus. In the nonestrous jill, the urogenital opening appears as a slit. During estrus, the vulva can swell considerably and resemble a doughnut. There are three to five pairs of nipples present in both jills and hobs.14,15

CARDIOVASCULAR AND LYMPHATIC SYSTEMS Heart and Blood Vessels The heart lies in a comparatively caudal location within the thoracic cavity, between the sixth and eighth ribs, with the apex to the left.15,18,20 The cardiodiaphragmatic ligament can contain varying amounts of fat.15A single brachiocephalic artery exits the aorta just proximal to the left subclavian artery. At the level of the thoracic inlet, this single artery divides into the right and left common carotid arteries and the right subclavian artery.7,18 This variation of the brachiocephalic artery is speculated to assist in maintaining cerebral blood flow during extreme rotation of the head and neck.32

Lymphatic Structures The thymus is located in the cranial mediastinum and can vary in size with age.7 Mediastinal lymph nodes are also present.15 The mediastinum is believed to be complete in ferrets.15 The

ferret possesses a robust system of lymph nodes and organs. The palatine tonsil is a flattened and ovoid structure that lies in the tonsillar fossa lateral to the ventral sulcus of the soft palate.7,15 The mandibular lymph node lies just rostral to the mandibular salivary gland and can easily be confused with this structure. The abdominal cavity has several prominent lymph nodes, including a prominent, palpable node at the root of the mesentery.27 The spleen sits in the left hypogastric region and parallels the greater curvature of the stomach.7 The reported normal size of the ferret spleen is 5.1 cm in length and 1.8 cm in width.7 However, benign extramedullary hematopoiesis is common in adult ferrets and results in a moderately to severely enlarged spleen.15 When enlarged, the spleen can extend diagonally from the upper left to the lower right abdomen, crossing the midline.

RESPIRATORY SYSTEM The lungs, contained within the slender and elongated thoracic cavity, are comparatively long in ferrets. The right lung is divided into cranial, middle, caudal, and accessory lobes. The left lung consists of a cranial and caudal lobe.7,14,15,18,20 Ferret lungs have a remarkably large filling capacity at about three times the predicted value for body size.14,18,30 Pinpoint yellow foci on the surface of the lungs observed at necropsy are foci of alveolar histiocytosis, the significance of which is unknown.14 The trachea is wide and very long, which results in comparatively lower airway resistance.14 The trachea bifurcates at the fifth intercostal space.15

ENDOCRINE SYSTEM Adrenal Glands The adrenal glands are located near each kidney, embedded in fat and covered by peritoneum.7 Each gland lies ventral to its ipsilateral adrenolumbar artery. The right gland lies in close apposition to the caudal vena cava and is draped by the caudate lobe of the liver. The right adrenal gland is slightly larger and longer than the left, at approximately 8 to 11 mm in length in one study.13 In another study, the adrenal gland length in females ranged from 5.0 to 10.0 mm for the left and 5.0 to 10.0 mm for the right; in males, it was 7.0 to 10.5 mm for the left and 7.5 to 13.5 for the right.26 Blood supply to either gland arises from the ipsilateral renal artery, with branches arising directly from the aorta, as well as the right adrenolumbar artery for the right adrenal gland.13 Variation to the adrenal blood supply exists. Accessory adrenal tissue can occasionally be found.7,15

Thyroid and Parathyroid Glands The thyroid gland is located ventrally along the neck between the third and eleventh tracheal rings, with each lobe positioned just lateral to the trachea.7,15 The parathyroid glands are small, pinkish structures that lie along the medial surface of the cranial portion of the thyroid, contacting the trachea at the fourth to fifth tracheal ring.7,15 The glands may be paired, occasionally single, on either side.7

MUSCULOSKELETAL SYSTEM The domestic ferret possesses a slender, elongated body, allowing it to fit through narrow spaces in pursuit of prey.7,29 The skeleton is lightweight but very flexible and strong.7,18 The skull is long and lacks sutures in the adult and the nasal openings are small compared to other mammals.7,20 Like all mustelids, ferrets

CHAPTER 1  Basic Anatomy, Physiology, and Husbandry possess strong muscles of mastication and can inflict a powerful and unrelenting bite.20 The vertebral formula of the ferret is C7, T15, L6 (5 or 7), S3, Cd18.7,15,18 The thorax is comparatively large with a narrow thoracic inlet and small first ribs.7,18,20 There are 15 pairs of ribs (occasionally 14), the first 10 of which are attached to the sternum and the last 5 comprising the costal arch.7,18 The spine is remarkably flexible, allowing the ferret to easily turn 180 degrees in a narrow passage. Despite possessing short legs, ferrets can climb remarkably well. Each foot possesses five digits with non-retractable claws. There are three phalangeal bones for each digit except the first digit, which has two.

NEUROLOGIC SYSTEM AND SPECIAL SENSES Brain and Spinal Cord The ferret brain has a typical mammalian design, and has been extensively reviewed elsewhere.17 The spinal cord and peripheral nerves of the ferret appear to be similar to those of the dog.15 The cauda equina begins at about the level of the last lumbar vertebra in the ferret.15

Special Senses Sight. The ferret eye has a prominent third eyelid, large cornea, horizontal elliptical pupillary opening, and a spherical lens.18,24 The eyes are open at 4 to 5 weeks after birth.18 Ocular movements are less pronounced in the ferret than in dogs and cats, and ferrets appear to track moving objects with movements of the head.15 Dorsal and ventral nasolacrimal puncta are present, although the dorsal punctum is smaller.15 The retina is similar to that of the dog and has a well-defined tapetum lucidum. The ferret is adapted to nocturnal living, and as such eyesight in the ferret is relatively poor compared to its olfactory and auditory senses.18 However, the ferret is a skilled hunter and can respond to movements of 25 to 45 cm/sec. Ferrets are believed to have limited color discrimination, at best.15,18,24 Hearing. The ferret pinna is set close to the head and points forward.7,18 The structure of the middle and inner ear is similar to that in the dog, although the ferret lacks a distinct tubular ear canal.18 Auditory function in the ferret is similar to the cat, although the auditory response may be more primitive.18 Although hearing range of ferrets is between 4 and 15 kHz (0.5 to 5 kHz in humans), lactating females can hear distress calls as high as 100 kHz from kits.15,18 Kits can hear by 32 days of age.18 Taste and Olfaction. Ferrets rely extensively on sense of smell.18 Ferrets have an elaborate nasal turbinate system like other carnivores, and appear to develop their olfactory preferences for food items during the first few months of life. These preferences may explain why diet changes in pet adult ferrets can be quite challenging.1 Taste sensation arises from the fungiform, vallate, and perhaps the foliate tongue papillae.15

PHYSIOLOGY AND REPRODUCTION PHYSIOLOGY Physiologic values of the domestic ferret are presented in Table 1-1.

BODY SIZE AND SEASONAL WEIGHT VARIATION Ferrets typically reach adult size by 6 months of age.7 Intact hobs weigh between 1 and 2 kg, in comparison to a weight between 0.6 and 1 kg for intact jills.18,23,29 If ferrets are neutered

9

before weaning, jills tend to become comparatively larger and hobs comparatively smaller, with a range of 0.8 to 1.2 kg for the two sexes. Neutered hobs lack the pronounced, muscular neck and shoulders characteristic of intact hobs. Ferrets tend to gain weight as winter approaches and lose weight in the spring.14 This seasonal weight fluctuation can be dramatic, with body weight variability approaching 40% in some individuals.10,15

REPRODUCTION Jills normally reach sexual maturity at 8 to 12 months of age, usually in the first spring after birth.10,14,22,29 Hobs typically reach puberty at about 9 months of age.29 The domestic ferret is seasonally polyestrus.14 Ferrets require alternating periods of long days and short days to have a normally functioning annual cycle.22 In both sexes, fertility increases as the days get longer. Spermatogenic activity in the hob occurs from December to July, and the testicles enlarge during this time. If not bred, intact jills will remain in persistent estrus from late March into early August.22,29 To an inexperienced observer, copulation appears violent, with the hob biting and dragging the jill by the  neck.14,29 A receptive jill will remain limp and not fight back. For successful mating, it is suggested to wait until the jill has been in estrus for 10 days before placing with the hob. The jill and the hob can be left together for up to 48 hours or be bred for shorter periods on 2 consecutive days.22,29 Jills are induced ovulators, requiring neck restraint and intromission, and ovulation generally occurs 30 to 36 hours after copulation.14,22,29 Gestation length in the domestic ferret is 41 days on average, with a range of 39 to 42 days. If fertilization does not occur after ovulation is induced, a pseudopregnancy lasting 40 to 42 days may occur.14,15,22 If ovulation is not induced mechanically or chemically, the jill will remain in estrus until a changing photoperiod occurs. Prolonged estrus introduces the risk of severe anemia due to bone marrow suppression caused by persistent hyperestrogenism.18 Jills deliver an average of 8 kits, with a range of 1 to 18. Kits weigh 6 to 12 grams at birth and are born blind and deaf with a thin coat of white fur.10,14,22 Jills raise the kits alone. Kits begin eating soft food by 21 days of age, often before their eyes open. Kits are generally weaned by 6 to 8 weeks of age.14,18

HUSBANDRY The following discussion of husbandry is an overview of the keeping of ferrets as pets. A wealth of information is now available on all these topics, providing more details. The literature also contains ample information about maintaining ferrets as laboratory animals; thus this topic is not addressed here. Behavior of the domestic ferret is discussed in Chapter 39.

HOUSING Ferrets can be housed either indoors or outdoors depending on the climatic conditions of the area. Ferrets are intelligent, curious animals that should not be continuously confined in a small cage. Pets need a safe play area where they can investigate a variety of objects, such as boxes, bags, and plastic pipes. Ferrets should be allowed a minimum of 2 hours a day of exercise. Lewington21 has an extensive description of an entire “ferretarium” and other outside enclosures for ferrets that are rich in environmental stimuli. A play or living area for ferrets must first be “ferret proofed”— that is, all holes to the outside or to areas from which the ferrets

10

SECTION I  Ferrets

cannot be retrieved must be blocked. In addition, ferrets like to burrow into the soft foam rubber of furniture and mattresses. Owners should be advised to cover the bottom of all couches, chairs, and mattresses with a piece of thin wood, or a sturdy wire mesh. The burrowing is not only destructive but also potentially life-threatening because ferrets may swallow the foam rubber and develop gastrointestinal obstructive disease. Reclining chairs have been implicated in the crushing deaths of many ferrets and should be removed from the environment. In addition, all access to foam or latex rubber items, such as dog and cat toys, athletic shoes, rubber bands, stereo speakers, headphones, and pipe insulation, should be eliminated. Ferrets will often chew these materials and ingest them. Ingestion of rubber foreign bodies is the most frequent cause of gastrointestinal obstruction, particularly in ferrets younger than 1 year. Up to two ferrets can use a wire cage of 24 × 24 × 18 inches as a home base when it is necessary to confine them. The floor can be either solid or wire. Glass tanks are not suitable for caging ferrets because they provide poor ventilation. Custom-built wooden cages can also be used, but care must be taken to protect corners, the lower third of walls, and the floor from contamination with urine and feces. A moisture-proof material such as linoleum or plastic or vinyl molding can be used for this purpose. If ferrets are kept outdoors, a portion of the cage should be shaded for protection from extremes of heat and cold, and a well-insulated nest box should be provided. Ferrets do not tolerate temperatures above 90°F (32°C), especially in the presence of high humidity, and may need to be brought indoors. In climates where the temperature drops below freezing, particularly if the temperature falls below 20°F (−7°C), a heated shelter is necessary. When caring for ferrets in a clinical setting, ensure that cages are escape proof. Ferrets have been known to squeeze between the bars of a standard dog or cat hospital cage. Using a piece of Plexiglas over cage bars that are too wide can keep the patient secure. Ferrets need a dark, enclosed sleeping area. This is also essential in the clinical setting, because the patient may become more anxious and stressed if denied access to such a “safe” area. Towels, old shirts, and cloth hats can be used, in addition to specific products designed for ferrets to sleep in, such as cloth tubes and tents. For the occasional ferret that insists on eating its cloth sleeping material, use a small cardboard, plastic, or wooden box with an access hole cut into it. Some owners use slings, hammocks, or shelves that are built into the cage to provide additional sleep and play areas. In a multiple-ferret household, at least one sleep area should be provided per ferret to allow the choice to be alone and to reduce fighting. Ferrets can be trained to use a litter box relatively easily. Because ferrets like to back up in corners to defecate or urinate, the litter box sides should be high enough to contain the excreted material. Pelleted litter material is recommended instead of clay or clumping litter. Because of the ferret’s short digestive transit time, the pet may not always reach the cage to use the litter box if it is not close by. Therefore owners should be advised to have several litter boxes available in various rooms of the house for use by the pet when it is uncaged.

ENVIRONMENTAL ENRICHMENT Environmental enrichment is of vital importance to the health and mental well-being of any animal kept in captivity. A sterile environment is an inhumane environment. The goal of enrichment should be to encourage healthy, natural behaviors and to

minimize injurious unnatural behaviors or even lack of activity. Consider these five areas when planning enrichment strategies: dietary, occupational activity, physical environment, sensory, and social. Specifics of ferret nutrition are discussed in the next section, but method of delivery of the food and the variety of food can be an enrichment in itself. Feeding the same diet, day in and day out, always available in a clearly visible container does not allow natural hunting behaviors. It is beneficial to alter feeding times, put food in different places, hide the food, give food choices, use food as part of positive reinforcement training, and rotate through a variety of food textures (hard to soft) and tastes to simulate more natural feeding patterns. Occupational therapy would include activities that stimulate physical exercise and mental stimulation. There are countless possibilities for creativity. For instance, ferrets like to be in underground burrows, so providing a network of tunnels and tubes made out of PVC pipe or other sturdy material stimulates movement. Ferrets also enjoy climbing up ramps and cloth- or carpetcovered objects. Digging boxes are greatly enjoyed and allow burrowing. Boxes filled with clean dirt can be given on a limitedtime basis in an area that is easy to clean up such as a bathtub or a secure outdoor enclosure. Some ferrets like to play in water, so an occasional trip to the bathtub with an inch of water in the bottom can be a special treat. Ferrets love the movement of a “prey” animal such as cat toys on a string or mechanical toys that can be introduced for a period of time into the environment. Enrichment of the physical environment includes the type and size of the cage or play area, the cage furniture, and the toys. Secure, private sleeping areas are very important to ferrets. Provide several choices with different textures and sizes. Have at least one hiding area per ferret that has an opening of a size the ferret can “defend” from other ferrets. Other furniture may include the climbing and hiding areas described under occupational activity. Toys for ferrets should not include any latex rubber toys intended for dogs or cats because these can end up as a gastrointestinal obstruction when ferrets chew on them. Instead, paper bags, cloth toys for cats or babies, or cat or human infant toys made of hard plastic or metal can be used. Toys that move or make noise are especially appreciated. Sensory enrichment is a part of many of the strategies already discussed. Stimulate olfactory, visual, tactile, auditory, and taste experiences. Food, toys, and the cage environment should be rich in sensory experiences. Using different scents, such as food scents, or safe essential oil scents on toys can turn them into a new item. If the ferret is an indoor pet, if at all possible arrange regular forays outside in a safe environment to promote further mental stimulation and exposure to novel experiences. Social enrichment can be fulfilled with contact with other ferrets and with other species, including humans. Play both contact and noncontact games with the ferret. Consider engaging in positive reinforcement training, which not only strengthens the bond between pet and caregiver, but also empowers the ferret to make choices, to continue to learn, and to be mentally stimulated.

NUTRITION Ferrets are obligate carnivores designed to eat whole, small prey animals including small to medium-sized mammals, birds, eggs, frogs, crustaceans, fish, worms, and insects. The ferret’s polecat ancestors would bring their kill home and store the excess in the den and eat small frequent meals rather than gorging.2

CHAPTER 1  Basic Anatomy, Physiology, and Husbandry Ferrets have a very short gastrointestinal tract with minimal gut flora and few brush border enzymes, so they cannot use carbohydrates efficiently or digest fiber.2 Ferrets in nature would only encounter carbohydrates as found in the partially digested stomach contents of their prey. Like many other obligate carnivores, young ferrets imprint on food by smell at a very young age and develop strong food preferences by the time they are a few months old. Therefore, regardless of the diet strategy that is chosen, ferrets should be exposed to a variety of food tastes, textures, smells, and different protein sources as juveniles so their diet will have more flexibility as an adult. This can be extremely helpful when ferrets experience medical conditions that may require restricted or altered diets at an older age. Ferrets should be fed a diet high in animal fat for energy, high in good-quality meat (not plant) protein, with minimal carbohydrate and fiber. A whole-prey diet or a balanced fresh or freeze-dried carnivore diet is the most appropriate for a ferret, and such diets are currently fed in many areas of the world with great success. Clean sources of prey food such as chicks, mice, and rats are now available in many areas of the country thanks to the reptile market, which uses these foods for carnivorous pets. If an owner does not want to feed a 100% whole-prey or raw diet, consider the occasional “treat” of a whole mouse or chick as valuable environmental enrichment. The stools of a ferret on a whole-prey diet are very firm and of low volume and odor. The most common diet fed to pet ferrets in the United States is dry kibble. Although there have been advancements in dry ferret food formulation, these diets still contain grain, which is necessary to hold the food in its solid shape. Very high levels of plant proteins in the diet can lead to urolithiasis.2 The hardness of the kibble may promote excessive dental wear and disease.6 Furthermore, excess dietary carbohydrates may affect the pancreas and may contribute to disease of the beta cells. Unfortunately, ferrets seem to enjoy sweet foods, and some commercial pet food companies have capitalized on this preference by producing ferret treats that are little more than sugar-coated grains. These treat foods are particularly dangerous to the health of the pet ferret. The stools of a ferret eating a dry kibble diet are formed but are soft, voluminous, and may contain visible undigested grain. If a dry diet is fed to the ferret, the owner should read the diet ingredients carefully. The crude protein should be 30% to 35% and composed primarily of high-quality meat sources, not grains; the fat content should be 15% to 20%.32 Dry food ingredients are listed on the label in descending order of their amount in the product. The first three ingredients of a ferret diet should be meat products. Because the diet is dry, it can be left out at all times. However, this is not very mentally stimulating and the ferret may establish stashes of food around the house, mimicking the storage of extra prey in its ancestral den. It is preferable not to leave the dry food out all the time, but rather to offer it two or three times a day and increase mental stimulation by varying the times and locations of feeding. Growing kits need 35% protein and 20% fat, and lactating females require 20% fat and twice the calories of the nonpregnant ferret.2 Acceptable supplemental foods to a dry diet include fresh human food grade raw organ or muscle meat and raw egg. Cooking the meat or eggs may not be absolutely necessary if they are fresh and are suitable for human consumption; however, raw or undercooked meat and eggs introduce the risk of certain enteric pathogens such as Salmonella species, Campylobacter jejuni, and E. coli. Adding a small amount of high-quality canned cat food

11

can add variety in texture, taste, and protein sources. Omega-3 oils, fish oils, or meat fat can be added to increase the fat content of the diet provided these additions are not allowed to become rancid, and are not fed in excess of about 20% of the diet. Dairy products have also been used as a fat and protein supplement, but some ferrets develop soft stools when fed these products. Even though ferrets enjoy eating fruits, they should only be given occasionally in very small amounts. Owners often overfeed these items, leading to a reduction in the consumption of a healthier diet and the overfeeding of sugars and fiber. Perhaps one of the best strategies for feeding a ferret is to offer a variety of food items throughout the ferret’s life, including a minimum of weekly whole-prey foods, daily high-quality ferret kibble, and small amounts of high-quality canned cat food or other meat-based treats fed two to three times a week. This diet would cover many nutritional bases, increase the flexibility of the ferret’s diet preferences, and is mentally enriching. Because of the short gastrointestinal transit time, fasting a ferret for longer than 3 hours is not necessary to check the fasting blood glucose level or to empty the GI tract for a surgical procedure. Ferrets older than 2 years in the United States are prone to develop insulinoma, and a longer fast could result in a serious hypoglycemic condition. Water should always be available in either a sipper bottle or a heavy crock-type bowl. Ferrets love to play in the water, so the bowl should not be easy to overturn. Supplements should not be added to the ferrets’ water supply.

References 1. Apfelbach R. Olfactory sign stimulus for prey selection in polecats. Zeitschrift fur Tierpsychol. 1973;33:270-273. 2. Bell JA. Ferret nutrition. Vet Clin North Am Exot Anim Pract. 1999;2:169-192. 3. Berkovitz BKB. Supernumerary deciduous incisors and the order of eruption of the incisor teeth in the albino ferret. J Zool Lond. 1968;155:445-449. 4. Bleavins MR, Aulerich RJ. Feed consumption and food passage time in mink (Mustela vison) and European ferrets (Mustela putorius furo). Lab Anim Sci. 1981;31:268-269. 5. Church B. Ferret-polecat domestication: genetic, taxonomic and phylogenetic relationships. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:122-150. 6. Church RR. The impact of diet on the dentition of the domesticated ferret. Exot DVM. 2007;9:30-39. 7. Evans HE, An NQ. Anatomy of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:19-69. 8. Fox JG. Taxonomy, history, and use. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:3-18. 9. Fox JG. Normal clinical and biologic parameters. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:183-210. 10. Fox JG, Bell JA. Growth, reproduction, and breeding. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:211-227. 11. Gentry A, Clutton-Brock J, Groves CP. The naming of wild animal species and their domestic derivatives. J Archaeol Sci. 2004;31:645-651. 12. Grzimek B. Grzimek’s encyclopedia of mammals. Vol 3. New York: McGraw-Hill; 1990;388–449. 13. Holmes RL. The adrenal glands of the ferret, Mustela putorius. J Anat. 1961;95:325-336.

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SECTION I  Ferrets

14. Hrapkiewicz K, Medina L. Clinical laboratory animal medicine: an introduction. 3rd ed. Ames, Iowa: Blackwell Publishing; 2007. 15. Ivey E, Morrisey J. Ferrets: examination and preventive medicine. Vet Clin North Am Exot Anim Pract. 1999;2:471-494. 16. Jacob S, Poddar S. Morphology and histochemistry of the ferret prostate. Acta Anat. 1986;125:268-273. 17. Lawes INC, Andrews PLR. Neuroanatomy of the ferret brain. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:71-102. 18. Lewington JH. Ferrets. In: O’Malley B, ed. Clinical anatomy and physiology of exotic species: structure and function of mammals, birds, reptiles and amphibians. Philadelphia: WB Saunders; 2005:237-261. 19. Lewington JH. Classification, history and current status of ferrets. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:3-14. 20. Lewington JH. External features and anatomy profile. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:15-33. 21. Lewington JH. Accommodation. In: Lewington JH, ed. Ferret husbandry, medicine and surgery. Philadelphia: WB Saunders; 2008:34-56. 22. Lindeberg H. Reproduction of the female ferret (Mustela putorius furo). Reprod Domest Anim. 2008;43(suppl 2):150-156. 23. MacDonald D. The velvet claw: a natural history of the carnivores. London: BBC Books; 1992.

24. Miller PE. Ferret ophthalmology. Semin Avian Exot Pet Med. 1997;6:146-151. 25. Moody KD, Bowman TA, Lang CM. Laboratory management of the ferret for biomedical research. Lab Anim Sci. 1985;35:272-279. 26. Neuwirth L, Collins B, Calderwood-Mays M, et  al. Adrenal ultrasonography correlated with histopathology in ferrets. Vet Radiol Ultrasound. 1997;38:69-74. 27. Paul-Murphy J, O’Brien RT, Spaeth A, et  al. Ultrasonography and fine needle aspirate cytology of the mesenteric lymph node in normal domestic ferrets (Mustela putorius furo). Vet Radiol Ultrasound. 1999;38:69-74. 28. Poddar S, Murgatroyd L. Morphological and histological study of the gastro-intestinal tract of the ferret. Acta Anat. 1976;96: 321-334. 29. Purcell K, Brown SA. Essentials of ferrets: a guide for practitioners. 2nd ed. Lakewood, Colorado: AAHA Press; 1999. 30. Vinegar A, Sinnett EE, Kosch PC, et al. Pulmonary physiology of the ferret and its potential as a model for inhalation toxicology. Lab Anim Sci. 1985;35:246-250. 31. Whary MT, Andrews PLR. Physiology of the ferret. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Baltimore: Williams & Wilkins; 1998:103-148. 32. Willis LS, Barrow MV. The ferret (Mustela putorius furo) as a laboratory animal. Lab Anim Sci. 1971;21:712-716. 33. Zeuner FE. A history of domesticated animals. New York: Harper & Row; 1963.

CHAPTER

2

Basic Approach to Veterinary Care

 

Katherine E. Quesenberry, DVM, MPH, Diplomate ABVP (Avian), and Connie Orcutt, DVM, Diplomate ABVP (Avian, Exotic Companion Mammal)

Restraint and Physical Examination Restraint Physical Examination Preventive Medicine Vaccinations Parasites Hospitalization Clinical and Treatment Techniques Venipuncture Intravenous Catheters Fluid Therapy Antibiotic and Drug Therapy Pain Management Nutritional Support Urine Collection and Urinalysis Urinary Catheterization Splenic Aspiration Bone Marrow Collection Tracheal Wash Blood Transfusion Blood Pressure Monitoring Diagnostic Peritoneal Lavage

Ferrets are commonly seen in many small animal veterinary practices. Special equipment needs are minimal, and the approach to handling ferrets is similar in many ways to that for dogs and cats. Ferret owners regularly seek veterinary care for a variety of reasons: ferrets need preventive vaccinations for canine distemper and rabies; ferrets have a relatively short life span compared with that of cats and dogs; ferrets in the United States and in some European countries have a high incidence of endocrine, gastrointestinal, and neoplastic diseases; and many of the diseases common to ferrets are not easily ignored by the pet owner (e.g., alopecia resulting from adrenal disease and hypoglycemic episodes caused by insulinoma). Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

RESTRAINT AND PHYSICAL EXAMINATION RESTRAINT Most ferrets are docile and can be easily examined without assistance. However, an assistant is usually needed when taking the rectal temperature, when administering injections or oral medications, or if an animal has a tendency to bite. Young ferrets often nip, and nursing females and ferrets that are handled infrequently may bite. Unlike dogs and cats, which growl, ferrets will bite without warning. Therefore always ask the owner if the ferret will bite before handling it and take precautions accordingly. Make sure to obtain the rabies vaccination history before physical examination, as reporting and rabies protocols for animal bites from vaccinated and unvaccinated ferrets differ (see below). Ferrets that are prone to bite and are not currently vaccinated for rabies may need tranquilization for procedures that require restraint. Depending on the ferret’s disposition, several basic manual restraint methods can be used for physical examination. For tractable animals, lightly restrain the ferret on the examination table. Examine the mucous membranes, oral cavity, head, and integument. Then pick the ferret up and use one hand for support under its body while using the second hand to auscultate the thorax and palpate the abdomen. The ferret can be scruffed at any time for vaccination, ear cleaning, or other procedures that may elicit an attempt to escape or bite. For a very active animal or one that bites, scruff the ferret at the back of its neck and suspend it with all four legs off the table (Fig. 2-1). Most ferrets become very relaxed with this hold, and the veterinarian is able to examine the oral cavity, head, and body; auscultate the chest; and palpate the abdomen easily. However, this method may not work for very fractious animals. To manually restrain a ferret for procedures such as venipuncture or ultrasound, hold it firmly by the scruff of its neck and around the hips without pulling the legs back. Most ferrets struggle if their legs are extended by pulling on the feet. Some animals can be distracted during a procedure by feeding a meat-based canned food (a/d Prescription Diet, Hill’s Pet Nutrition, Topeka, KS; Eukanuba Maximum-Calorie, The Iams 13

14

SECTION I  Ferrets

Fig. 2-1  Restrain an active ferret by scruffing the loose skin on the back of the neck. The ferret will relax and allow you to palpate the abdomen or administer a vaccine.

Company, Dayton, OH) or a small amount of a supplement such as ­FerreTone (8-in-1 Pet Products, Islandia, NY) by syringe. Avoid products containing sugar, which can affect blood glucose values, particularly in ferrets with insulinoma. For very fractious or anxious animals or for procedures requiring lengthy restraint, light tranquilization or sedation may be indicated (see Chapter 31).

PHYSICAL EXAMINATION Most ferrets strenuously object to having their temperature taken with a rectal thermometer. If a ferret struggles during the examination, the temperature taken at the end of the examination may be artificially high. Therefore measure the rectal temperature early in the physical examination with a flexible digital thermometer that is well lubricated. The normal range of rectal temperature of a ferret is 100.5°F to 102.5°F (38.0°C to 39.2°C); a mean of 102°F (38.8°C), with a wider range of 100°F to 104°F (37.8°C to 40.0°C), is also reported.21 Interestingly, in normal ferrets housed outdoors at a fur farm in very cold ambient temperatures (21°F [−6.1°C]), the mean 24-hour mean core body temperature measured by sterile thermosensitive data loggers implanted in their abdomens was 99.3°F (37.4°C), with a range of 97.3°F to 101.1°F (36.3°C to 38.4°C).41 The physical examination of a ferret is basically the same as that of any small mammal and can be performed quickly and efficiently if a few simple guidelines are followed. Observe the attitude and alertness of the animal. Ferrets may sleep in the carrier in the veterinary office; however, once awakened for the examination, a ferret should be alert and responsive. Assess hydration by observing the skin turgor of the eyelids, tenting of the skin at the back of the neck, and moistness of the oral mucous membranes. However, skin turgor can be difficult to evaluate in a cachectic animal. Estimate the capillary refill time by digitally pressing on the gingiva.

Examine the eyes, nose, ears, and facial symmetry. Cataracts can develop in both juvenile and adult animals. Retinal degeneration is another ophthalmic disorder seen in ferrets and may be indicated by abnormal pupil dilation. Inspect for nasal discharge and ask the owner about any history of sneezing or coughing. The ears may have a brown waxy discharge, but the presence of excessive brown exudate may indicate infestation with ear mites (Otodectes cynotis). Bruxism often indicates gastrointestinal discomfort. The teeth of ferrets should be clean and the gingiva pink. Dental tartar is commonly present in pet ferrets. The amount of plaque may be exacerbated by the feeding of soft foods or sugary treats, such as raisins, and is possibly related to a dry kibble diet.14 Tartar most commonly accumulates on the first and second maxillary premolars. Excessive dental tartar should be removed by dental techniques used in dogs and cats, and measures to prevent tartar buildup should be implemented. As a preventive, a pet dentifrice or tartar control toothpaste25,32 can be applied to the teeth to decrease formation of calculus. Gingivitis, which manifests as erythematous gingival tissue that sometimes bleeds, is a common sequela of excessive dental tartar. Ferrets often break off the tip of one or both canine teeth; however, they rarely exhibit clinical signs of sensitivity or pain associated with a fractured canine. If the tooth turns dark or the ferret exhibits sensitivity when eating, recommend a root canal or extraction, depending on the degree of damage to the tooth (see Chapter 32). Rarely, an infected root of a fractured canine can cause swelling of the ipsilateral submandibular lymph node. If swelling is present, dental radiographs, canine tooth extraction, and possibly lymph node biopsy are indicated. Observe the symmetry of the face. Although uncommon, salivary mucoceles occur in ferrets and are noticeable as a unilateral swelling on the side of the face, usually in the cheek or temporal area (see Chapter 3). Palpate the regional lymph nodes of the neck and axillary, popliteal, and inguinal areas. Nodes should be soft and may sometimes feel enlarged in large or overweight animals because of surrounding fat. Any degree of firmness or asymmetry in one or more nodes is suspicious and warrants a fine-needle aspirate or a biopsy. If two or more nodes are enlarged and firm, a full diagnostic workup is indicated. Auscultate the heart and lungs in a quiet room. Ferrets have a rapid heart rate (180 to 250 beats/min) and often a pronounced sinus arrhythmia. If a ferret is excited and has a very rapid heart rate, subtle murmurs may be missed. Valvular disease, cardiomyopathy, and congestive heart failure are seen in ferrets, and any murmur or abnormal heart rhythm should be investigated further (see Chapter 5). Palpate the abdomen while holding the ferret off the table, either by scruffing the neck or supporting the ferret with one hand. This allows the abdominal organs to displace downward, facilitating palpation. If the history is consistent with an intestinal foreign body or urinary blockage, palpate gently to avoid causing iatrogenic injury, such as a ruptured stomach or bladder. Palpate the cranial abdomen, paying particular attention to the presence of gas or any irregularly shaped mass in the stomach area, especially in ferrets with a history of vomiting, melena, or chronic weight loss. The spleen is commonly enlarged in ferrets; this may or may not be significant, depending on other clinical findings (see Chapter 5). Palpate a large spleen gently to avoid iatrogenic damage. A very enlarged spleen may indicate systemic disease or, very rarely, idiopathic hypersplenism, and

CHAPTER 2  Basic Approach to Veterinary Care further diagnostic workup is warranted. Always note any degree of splenic enlargement in the medical record so that this finding can be rechecked at future examinations. Examine the genital area, observing the size of the vulva in females. Vulvar enlargement in a spayed female is consistent with either adrenal disease or an ovarian remnant; the latter is rare. If the vulva is of normal size, show this to the owner so that any vulvar enlargement in the future will be noticed. Examine the preputial area and size of the testicles of male ferrets; preputial and testicular tumors are sometimes seen. Check the fur coat for evidence of alopecia. Alopecia of the tail tip is common in ferrets and may be incidental and transient or an early sign of adrenal disease. Symmetric, bilateral alopecia or thinning of the hair coat that begins at the tail base and progresses cranially is a common clinical finding in ferrets with adrenal disease. Examine the skin on the back and neck for evidence of scratching or alopecia. Pruritus may be present with adrenal disease (common) or with ectoparasites (fleas, Sarcoptes scabiei). Check closely visually and by searching through the hair coat with your fingers for evidence of skin masses. Mast cell tumors are common and can range in diameter from a few millimeters to over a centimeter. Often, the fur around a mast cell tumor is parted and matted with dark blood from the animal’s scratching. Other types of skin tumors, such as sebaceous adenomas and basal cell tumors, are also common (see Chapter 9). Perform an excisional biopsy of any lump found on the skin.

PREVENTIVE MEDICINE Young, recently purchased ferrets need serial distemper vaccinations until they are 13 to 14 weeks of age.2 Rabies vaccines should be given annually beginning at 3 months of age.15 ­Ferrets should be examined annually until they are 4 or 5 years of age; middle-aged and older animals should be examined twice yearly because of the high incidence of metabolic disease and neoplasia. Annual blood tests (consisting of a complete blood count and plasma or serum biochemical analysis) are recommended for older animals. Measure the blood glucose concentration twice yearly in healthy middle-aged and older ferrets; more frequent monitoring is needed in ferrets with insulinoma. An endocrine panel is indicated in ferrets with hair loss on the tail or other clinical signs suggestive of early adrenal disease (see Chapter 7). Testing for infectious diseases may be warranted, especially in new ferrets that will be introduced into a multi-ferret household or those that are taken to ferret shows. Currently, ferrets can be tested for Aleutian disease virus and ferret enteric coronavirus by polymerase chain reaction (PCR) testing (Michigan State University, Diagnostic Center for Population and Animal Health, www.animalhealth.msu.edu; Veterinary Molecular Diagnostics, www.vmdlabs.com). Serologic tests for Aleutian disease by enzyme-linked immunosorbent assay (ELISA) and counterimmunoelectrophoresis (CIEP) are also available (see Chapter 5).

VACCINATIONS Canine Distemper Ferrets must be vaccinated against canine distemper virus. Currently, one vaccine is approved by the U.S. Department of Agriculture for use in ferrets: PureVax (Merial, Athens, GA). Because PureVax is a canarypox-vectored recombinant vaccine

15

it does not contain adjuvants or the complete distemper virus; thus many of the postvaccination risks have been reduced. This product has a wide safety margin and has proved effective in protecting ferrets against canine distemper infection.58 Another distemper vaccine that was widely used previously (Fervac-D, United Vaccines, Inc., Madison, WI) is no longer available. Fervac-D was a modified live virus vaccine propagated in avian cell lines. Another modified live canine distemper vaccine (Galaxy D, Merck/Schering-Plough Animal Health, Whitehouse Station, NJ) has been studied for safety and efficacy in ferrets. This product, derived from the Onderstepoort distemper strain and attenuated in a primate cell line, proved effective in preventing canine distemper in young ferrets challenged after serial vaccination.64 However, duration of immunity with this product is not known, and its use in clinical animals is extralabel, requiring informed owner consent. Although no vaccine reactions were reported in the study, the incidence of vaccine reactions with Galaxy D is unknown because experience with repeated longterm use in ferrets has been limited.64 Because of the possibility of vaccine-induced disease, especially in immunosuppressed or sick ferrets, do not use combination canine vaccines or vaccines of ferret cell or low-passage canine cell origin. In young ferrets, the half-life of maternal antibody to canine distemper virus is 9.43 days.2 Vaccinate young ferrets for distemper at 8 weeks of age, then give two additional boosters at 3-week intervals for a total of three vaccinations. Give booster vaccines annually.

Rabies All ferrets should be vaccinated against rabies.15 A killed rabies vaccine is approved for use in ferrets (Imrab-3 or Imrab-3 TF, Merial, Duluth, GA) and is effective in producing immunity for at least 1 year.55 Current recommendations are to vaccinate healthy ferrets at 3 months of age at a dose of 1 mL administered subcutaneously. Give booster vaccinations annually. Titers develop within 30 days of rabies vaccination.55 In ferrets that were experimentally inoculated intramuscularly with skunk-origin rabies virus, the mean incubation period was 33 days and the mean morbidity period was 4 to 5 days.42 Clinical signs were ascending paralysis, ataxia, cachexia, bladder atony, fever, hyperactivity, tremors, and paresthesia. Virus antigen was present in the brain tissue of all ferrets with clinical signs of rabies, and virus was isolated from the salivary gland of one ferret. In a similar study of ferrets inoculated with a raccoon rabies isolate, the mean incubation period was 28 days. Virus was isolated from the salivary glands of 63% of rabid ferrets, and 47% shed virus in saliva. Virus excretion began from 2 days before until 6 days after the onset of illness.43 In an earlier study of ferrets with experimentally induced rabies, only mild clinical signs were observed before death.7 Infected ferrets exhibited restlessness and apathy, and some showed paresis. Sick animals did not attempt to bite when threatened, and virus was not excreted in the submaxillary salivary glands of animals that died. In this study, the authors concluded that ferrets are 50,000 times less susceptible to rabies than fox and 300 times less susceptible than hares. In another study, ferrets that were fed up to 25 carcasses of mice infected with rabies did not develop the disease; in contrast, skunks become fatally infected after the consumption of only one carcass.4 Ferrets are considered currently immunized 28 days after the initial rabies vaccination and immediately after a booster vaccination.15 If a healthy pet ferret bites a person, current

16

SECTION I  Ferrets

recommendations of the Compendium of Animal Rabies Prevention and Control are to confine and observe the animal for 10 days; the ferret should not be vaccinated during this period.15 If signs of illness develop, this should be reported to the local health department and a veterinarian should evaluate the animal. If signs suggest rabies, the ferret must be euthanized and protocols for rabies evaluation should be followed. If a stray ferret bites a person, the ferret must be euthanized and submitted immediately for rabies testing. For a vaccinated ferret exposed to a possible rabid animal, recommendations are to revaccinate the ferret immediately and quarantine for 45 days. An unvaccinated animal that is exposed to a rabid animal should be euthanized immediately and submitted for rabies testing. See the website of the Centers for Disease Control and Prevention (www.cdc. gov/mmwr/preview/mmwrhtml/rr5702a1.htm) or the National Association of Public Health Veterinarians (www.nasphv.org/) for specific guidelines.

Vaccine-Associated Adverse Events In ferrets, adverse events associated with vaccination are primarily type I hypersensitivity reactions or anaphylaxis.37 Type I hypersensitivity reactions involve lymphoid tissue associated with mucosal surfaces (skin, intestines, and lungs) and result from the interaction of antigen and immunoglobulin E in mast cells or basophils. Ferrets with mild reactions may exhibit pruritus and skin erythema. More severe reactions are typified by vomiting, diarrhea, piloerection, hyperthermia, cardiovascular collapse, or death. Vaccine reactions are most common after distemper vaccination but may also occur after rabies vaccination. In a study of vaccine reactions in 3,857 ferrets, the incidence of adverse events associated with rabies vaccine alone, distemper vaccine alone, and rabies and distemper vaccines together were 0.51%, 1.0%, and 0.85%, respectively. The incidence of adverse events did not differ significantly among these three groups; however, the cumulative number of distemper vaccinations received was significantly associated with the occurrence of an adverse event. The distemper vaccines used in this population of ferrets were PureVax and Fervac D; however, the two distemper vaccines were grouped collectively in the analysis, and the incidence of adverse events associated with the individual distemper vaccines was not reported. Sex, age, and body weight were not associated with occurrence of an adverse event. All reactions occurred immediately after vaccination and most commonly consisted of vomiting and diarrhea. In another study of 143 ferrets, the incidence of adverse events after administering distemper (5.9%) (Fervac D), rabies (5.6%) (Imrab-3), or both vaccines (5.6%) was not significantly different. In a 2001 report of vaccine reactions in ferrets reported to the United States Pharmacopeia Veterinary Practitioners’ Reporting Program, 65% (54 of 83) of reports involved administration of FerVac D; 24% (20 of 83) involved concomitant administration of FerVac D and Imrab; and 11% (9 of 83) involved administration of Imrab alone (PureVax was not approved for use at the time these data were collected).37 According to Merial’s product information, the incidence of vaccine reactions with PureVax is 0.3%. No data are available for products not licensed for use in ferrets. Veterinarians are not required to report vaccine-associated adverse events, and surveillance of these events is passive, relying on voluntary reporting by practitioners.37 Vaccine-associated adverse events can be reported to the Center for Biologics, U.S. Department of Agriculture (1-800-752-6255;  www.aphis. usda.gov/animal_health/vet_biologics/vb_adverse_event.shtml).

Always follow the manufacturer’s instructions for vaccine administration and inform the owner of the possibility of a reaction before vaccinating. Have the owner monitor the ferret in the waiting area for 30 minutes or more after vaccination with any product. As stated, most reactions occur almost immediately after vaccination. If a ferret has an adverse reaction, administer an antihistamine (e.g., diphenhydramine hydrochloride [Benadryl, ParkeDavis, Morris Plains, NJ], 0.5 to 2.0 mg/kg intravenously [IV] or intramuscularly [IM]), epinephrine (20 μg/kg IV, IM, subcutaneously [SC], or intratracheally), or a short-acting corticosteroid (e.g., dexamethasone sodium phosphate, 1 to 2 mg/kg IV or IM), and give supportive care. For any biologic product, veterinarians must assess risk versus benefit of vaccination. The treatment options for ferrets that have had a vaccine reaction include not vaccinating if the risk of exposure is minimal; administering diphenhydramine (2 mg/kg orally [PO] or SC) at least 15 minutes before vaccination; or, for distemper, administering a different product. Vaccine injection-site sarcomas have been described in ferrets.39,40 In one report, 7 of 10 fibrosarcomas in ferrets were from locations used for vaccination.39 Fibrosarcomas from injection sites had a high degree of cellular pleomorphism and similar histologic, immunohistochemical, and ultrastructural features as those reported for feline vaccine-associated sarcomas. In the reported cases in ferrets, no definitive association could be made between the fibrosarcoma and the type of vaccine. In cats, adjuvanted vaccines are most likely to be involved in tumor development. However, although injection-site sarcomas may occur in ferrets, ferrets appear less prone than cats to tumor development. In a study of vaccine reactions in ferrets, mink, and cats, cats had more lymphocytes at the injection site than either ferrets or mink after vaccination with three different rabies vaccines.11 Results of this study suggest a lower species susceptibility to vaccine-associated sarcomas in ferrets than in cats.

PARASITES Endoparasites Gastrointestinal parasitism is uncommon in domestic ferrets. Rarely, pet ferrets may become infected with parasites from other natural hosts through intermediate hosts or vectors. Protozoan parasites are occasionally seen. Therefore perform routine fecal flotations and direct fecal smears for all young ferrets at the initial examination. Coccidiosis (Isospora species) is seen infrequently, usually in young ferrets, which shed oocysts between 6 and 16 weeks of age.3 The infection is usually subclinical; occasionally, however, ferrets may have loose stool or bloody diarrhea. Treatment of ferrets with coccidiosis is similar to that of other small animals and should be continued for at least 2 weeks. Coccidiostats, such as sulfadimethoxine and amprolium, are effective and safe. The Isospora species that infect ferrets may cross-infect dogs and cats; therefore other pets in the household should be checked for coccidia and treated as needed. Giardiasis is occasionally seen in ferrets. Results of a recent study on molecular characterization of Giardia duodenalis isolates from pet ferrets show that genetic sequences from isolates in ferrets differ from isolates of humans and other animals, suggesting that Giardia isolates from ferrets may be host specific.1 Giardia species can be detected by identifying cysts or

CHAPTER 2  Basic Approach to Veterinary Care trophozoites in a fresh fecal smear or zinc sulfate flotation, or by fecal ELISA. Treat ferrets with giardiasis with metronidazole (20 mg/kg PO q12h) for 5 to 10 days. Fenbendazole (50 mg/kg PO q24h for 3 to 5 days) is used in dogs and cats, but safety and efficacy in ferrets are unknown. Cryptosporidiosis can occur in a high percentage of young ferrets.53 Infection is usually subclinical in both immunocompetent and immunosuppressed animals. Although most immunocompetent animals recover from infection within 2 to 3 weeks, infection can persist for months in immunosuppressed animals. Oocysts of Cryptosporidium are small (3 to 5 μm) and difficult to detect but can be found in samples of fresh feces examined immediately after acid-fast staining.3,53 No treatments exist for Cryptosporidium infection. Because of the zoonotic potential, ferrets may be a source of infection for human beings, especially immunocompromised individuals with acquired immunodeficiency syndrome (AIDS).53 Heartworms (Dirofilaria immitis) can cause disease in ferrets. Ferrets that are housed outdoors in heartworm-endemic areas are most susceptible to infection; however, all ferrets in endemic areas should be given preventive medicine. Oral administration of ivermectin is currently the most practical preventive measure because it is administered once per month (see Chapter 5 and Appendix).

Ectoparasites Ear mites (Otodectes cynotis) are common in ferrets, but affected animals rarely exhibit pruritus or irritation. This mite species also infects dogs and cats, and animals in households with multiple pets can transmit mites to other animals. A red-brown, thick, waxy discharge in the ear canal and pinna characterizes infection. A direct smear of the exudate reveals adult mites or eggs. Because ferrets normally have brown ear wax, the color or appearance of debris in the ear canal is not pathognomonic for mites. At the initial examination, check all ferrets for ear mites and do follow-up checks at the annual examination in ferrets kept in multiple-pet households. Several products, including selamectin, are effective in treatment (see Chapter 9). Flea infestation (Ctenocephalides species) is most common in ferrets kept in households with dogs or cats. Ferrets with chronic infestation can become severely anemic. Check all ferrets during the physical examination for signs of fleas or flea dirt. Treat infested animals with products safe for use in cats and institute flea control measures (see Chapter 9). Ticks are rarely seen in domestic ferrets, and Lyme disease in ferrets has not been reported.

HOSPITALIZATION Ferrets can be hospitalized in standard stainless steel hospital cages with some adaptations. Ferrets are agile escape artists and can squeeze through even very small openings. In many standard cages designed for veterinary hospitals, the bar spacing is too wide, allowing an easy avenue of escape. For housing ferrets, use only cages with small spacing between vertical bars or use cages with small crossbars. Alternatively, adapt standard cages for use by attaching a Plexiglas plate to the front of the cage at least half the height of the cage door or higher. The plate will prevent escape through the bars yet can be easily detached and cleaned. Commercial hospital cages with Plexiglas fronts and access ports can be used for ferrets. There is no avenue of escape, and

17

ferrets are visible at all times. Acrylic or laminate animal intensive care cages or incubators also can be used to house ferrets and are especially useful for animals that need supplemental heat or oxygen. The cage should be large enough to accommodate a sleeping area or box and an area for defecation and urination. Ferrets are very careful about not soiling their sleeping area, even when very sick. All ferrets like to burrow and should be given opportunity to do so while hospitalized. Clean towels make excellent burrowing material. Alternatively, a mound of shredded paper provides much satisfaction to hospitalized animals. If not provided with burrowing material, many ferrets will burrow underneath the cage paper. Extra-small padded pet beds and fleece pet “pockets” work well as sleeping areas. An oxygen cage should be available for use with dyspneic animals. Monitor the temperature in commercial oxygen cages closely, because ferrets can become hypothermic quickly at cool cage temperatures that are used for dogs and cats. Conversely, ferrets can overheat at temperatures used for avian patients. Provide water for hospitalized ferrets in either water bottles or small weighted bowls. Ask the owner which type of watering system the ferret is accustomed to before hospitalization. Ferrets can be finicky eaters and should be fed their regular diet while hospitalized, if possible. Otherwise, feed a very palatable ferret food or a premium-quality, high-protein cat or kitten chow. If dietary changes are needed in the regular diet, recommend that changes be made gradually after the ferret has been released from the hospital. For animals that are anorexic, force-feed a high-calorie semisolid food or supplement until the animal is eating on its own (see later discussion).

CLINICAL AND TREATMENT TECHNIQUES VENIPUNCTURE Obtaining a blood sample from a ferret is relatively easy and usually does not require anesthesia. Several venipuncture sites are readily accessible; the technique and site chosen depend on how much blood is needed and the availability of assistants for restraint. Anesthesia or tranquilization can be used if assistants are unavailable, but anesthesia may affect hematologic values (see later discussion).35 Ferrets often can be distracted during restraint for venipuncture by offering semi-solid food or a product such as FerreTone (8-in-1 Pet Products) by syringe. Avoid using supplements with corn syrup or other sugars, as this will affect blood glucose levels, and collect blood for glucose determination or other fasting samples before offering food. Most veterinary laboratories offer small mammal hematologic and biochemical panels that can be done with 1.0 mL or less of blood. In-clinic point-of-care analyzers require very small sample sizes (usually 100 μL). The blood volume of healthy ferrets is approximately 40 mL in average-sized females weighing 750 g and 60 mL in males weighing 1 kg.21 Up to 10% of the blood volume can be safely withdrawn at one time in a normal ferret, but collect only the minimum amount needed for analysis. Repeated blood drawing can contribute to anemia in sick animals hospitalized for long periods. Two sites are commonly accessed to obtain large blood volumes in ferrets. The jugular vein can be approached in the neck by using the conventional technique used in cats, with the forelegs extended over the edge of a table and the neck extended up (Fig. 2-2). Use a 25-gauge needle with a 1- to 3-mL syringe for

18

SECTION I  Ferrets

Fig. 2-2  Restraint for jugular venipuncture in a ferret. Restrain the ferret similar to a cat, with the legs pulled down and the head back. Shave the neck to improve visibility of the jugular vein in the lateral neck. After the vein is punctured, the head can be “pumped” up and down slowly to facilitate blood flow.

venipuncture in most ferrets; a 22-gauge needle can be used in large males. Shave the neck at the venipuncture site to enhance visibility of the jugular vein. The vein is relatively superficial and is located more lateral in the neck than it is in dogs or cats, and it is sometimes difficult to locate in heavy males. Once the needle is inserted, the blood should flow easily into the syringe; if the neck is overextended and the head is arched back, the blood may not flow readily from the vein. Relax the hold on the head or gently “pump” the vein by moving the head slowly up and down to enhance blood flow into the syringe. With ferrets that resist limb extension, a towel-wrap technique can be used.9 Scruff the ferret with its front legs extended caudally against the ventral thorax, and wrap the animal’s body firmly with a towel from the base of the neck down. An assistant is needed to restrain the toweled ferret in dorsal recumbency while scruffing the cranial neck. Apply pressure lateral to the thoracic inlet to visualize or palpate the jugular vein lying between the thoracic inlet and the base of the ear. However, with very fractious animals, even this technique may be difficult without tranquilization. The second venipuncture site to obtain large blood samples is the cranial vena cava. The actual site of venipuncture has been called the thoracic portion of the jugular vein12,60; however, anatomically it is more likely the right or left brachiocephalic trunk or the anterior vena cava itself, depending on the point of entry and the depth of needle penetration (Fig. 2-3, A). This technique is safe in ferrets because of the long anterior vena cava and the caudal location of the heart in the thoracic cavity, which is approximately 3 cm from the thoracic inlet. However, rare instances of hemorrhage into the anterior thoracic cavity can occur. Restrain the ferret on its back with the forelegs pulled caudally and the head and neck extended (Fig. 2-3, B). In an unanesthetized ferret, two assistants are usually needed, one for restraint of the forelegs and head and the other for restraint

of the rear just cranial to the pelvis. Insert a 25-gauge needle with an attached 1-mL or 3-mL syringe into the thoracic cavity between the first rib and the manubrium at an angle 30 to 45 degrees to the body. Direct the needle toward the opposite rear leg or most caudal rib and insert it almost to the hub. Pull back on the plunger as the needle is slowly withdrawn until blood begins to fill the syringe. If the ferret struggles, quickly withdraw the needle and wait until the ferret is quiet before making a second attempt. In very fractious or active ferrets, jugular venipuncture or use of tranquilization are safer choices to avoid lacerating the vessels. The lateral saphenous or cephalic vein can be used if only a small amount of blood is needed to measure a packed cell volume or blood glucose level. To prevent collapse of the vein during venipuncture, use an insulin syringe with an attached 27- or 28-gauge needle. The saphenous vein lies just proximal to the hock joint on the lateral surface of the leg; the cephalic vein is in the same anatomic location as in a dog. Before venipuncture, shave the fur from the area to enhance visibility of the vein. Although rarely used in pet ferrets, venipuncture of the tail artery is described to obtain blood samples.8 Venipuncture at this site can be painful; anesthetize the ferret for this technique. The artery is located 2 to 3 mm deep to the skin. Insert a syringe with a 21-gauge needle into the ventral midline of the tail directed toward the body. Once the artery is entered, slowly withdraw the plunger until blood fills the syringe. Apply pressure to the venipuncture site for 2 to 3 minutes after the needle has been withdrawn.

Reference Ranges Published reference intervals for hematologic, biochemical, and plasma electrophoresis values in ferrets are listed in Tables 2-1, 2-2, and 2-3. Other published sources of reference intervals for ferrets are available.21,22 Additionally, most clinical veterinary laboratories routinely provide reference intervals for ferret hematologic and biochemical values. Published reference intervals for white blood cell (WBC) counts in ferrets vary from 2.5 to 19.1 × 103 cells/μL21,27,61; however, WBC counts generally tend to be low in ferrets. In one study, mean WBC values were 5.7 and 5.6 × 103 cells/μL in male and female ferrets, respectively.22 High WBC counts are not seen as commonly in ferrets as in dogs and cats, perhaps in part because infectious bacterial diseases are comparatively uncommon in ferrets. Isoflurane anesthesia can cause decreases in all hematologic values beginning at induction of anesthesia and reaching maximal effects at 15 minutes after induction.35 Therefore the complete blood count values of blood samples collected while a ferret is anesthetized must be carefully interpreted. Reference intervals for blood coagulation times in ferrets have been published. In a recent study, blood samples were collected into sodium citrate in a ratio of 9:1 from 18 clinically healthy ferrets (12 males, 6 females, all neutered).6 Results showed some variation in values obtained by the method used for measurement. Mean prothombin time (PT) was 12.3 seconds (range, 11.6 to 12.7 seconds) measured by fibrometer and 10.9 seconds (range, 10.6 to 11.6 seconds) measured by an automated coagulation analyzer. Mean activated partial thromboplastin time (aPTT) was 18.7 seconds (range, 17.5 to 21.1 seconds) by fibrometer and 18.1 seconds (range, 16.5 to 20.5 seconds) by automated coagulation analyzer. Mean fibrinogen concentration was 107.4 mg/dL (range,

CHAPTER 2  Basic Approach to Veterinary Care

19

JV LBT RBT

AVC

A

B Fig. 2-3  A, Dissection of the thoracic cavity of a ferret illustrating the site for blood collection using the anterior vena cava technique. The sternum and ventral ribs are removed. The site of venipuncture is either the right brachiocephalic trunk (RBT) or left brachiocephalic trunk (LBT) or the anterior vena cava (AVC), depending on the point of entry and depth of penetration (see marker). The jugular vein (JV) is usually lateral and cranial to the venipuncture site. The base of the first two ribs are shown by arrows. B, A ferret is restrained for venipuncture of the anterior vena cava. Both forelegs are pulled back, hindlegs are restrained, and the neck is extended.

  Table 2-1    Reference Intervals for Hematologic Values in Ferrets ALBINO37

FITCH16

Value

Combined Sexa

Maleb

Female

Malec

Female

Hematocrit (%) Hemoglobin (g/dL) Red blood cells (×106/μL) Reticulocytes (%) White blood cells (×103/μL) Neutrophils (%) (cells/μL) Lymphocytes (%) (cells/μL) Monocytes (%) (cells/μL) Eosinophils (%) (cells/μL) Basophils (%) (cells/μL) Bands (cells/μL) Platelets (×103/μL) Mean corpuscular volume (fL) Mean corpuscular hemoglobin (g/dL) Mean corpuscular hemoglobin concentration (g/dL)

36-48 12.2-16.5 7.01-9.65 4.3-10.7 18-47 41-73 0-4 0-4 0-2 200-459 50-54 15-18 32-35

44-61 16.3-18.2 7.30-12.18 1-12 4.4-19.1 11-82 12-54 0-9 0-7 0-2 297-730 -

42-55 14.8-17.4 6.77-9.76 2-14 4.0-18.2 43-84 12-50 2-8 0-5 0-1 310-910 -

46-57 15.2-17.7 5.6-10.8 616-7020 1728-4704 0-432 112-768 0-112 0-972 -

47-51 15.2-17.4 2.5-8.6 725-2409 1475-5590 100-372 50-516 0-172 0-248 -

aCombined

male and female pet ferrets (n = 60). From Cray C, Avian and Wildlife Laboratory, Miller School of Medicine, University of Miami, Miami, FL. bIntact males. cCastrated males.

20

SECTION I  Ferrets   Table 2-2    Reference Intervals for Biochemical Values in Ferrets SERUM Analyte

Plasmaa

Albinob

Fitchc

Alanine aminotransferase (U/L) Albumin (g/dL) Alkaline phosphatase (U/L) Amylase (U/L) Aspartate aminotransferase (U/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Carbon dioxide (mmol/L) Cholesterol (mg/dL) Chloride (mmol/L) Creatinine (mg/dL) Creatine phosphokinase (U/L) Glucose (mg/dL) Gamma glutamine transferase (U/L) Lactate dehydrogenase (U/L) Phosphorus (mg/dL) Potassium (mmol/L) Sodium (mmol/L) Total protein (g/dL) Triglycerides (mg/dL) Uric acid (mg/dL)

65-128 2.5-4.0 25-60 26-36 70-100 0.2-0.5 18-32 8.1-9.5 22-29 119-163 0.2-0.5 55-93 80-117 8-34 200-1400 5.1-6.5 4.5-6.1 142-148 4.5-6.2 30-140 1.3-1.9

2.6-3.8 9-84 28-120 40°C) +++ ++ +a Almost 100% fatal

++ (Mucoserous) +++ +++ ++b — — — Self-limitingc

Frequency of clinical signs: +, may be present; ++, common; +++, usual presentation; —, absent. aCentral nervous system signs seen in advanced stages of disease (rarely the only signs). bPyrexia occurs early in the course of disease and may be resolved by the time of presentation. cInfluenza virus infection can be fatal in neonates.

therapy. As with any flu patient, parenteral fluids to maintain hydration and antibiotics to treat secondary bacterial infections may be indicated. To relieve nasal congestion, intranasal delivery of phenylephrine can be effective.7 The antiviral medication amantadine (6 mg/kg PO q12h) (Symmetrel; ENDO Pharmaceuticals, Chadds Ford, PA) has been experimentally effective in treating ferrets with influenza, although resistance in humans is widely reported.15 Other antiviral medications include neuraminidase inhibitors like zanamivir (12.5 mg/kg as a one-time intranasal dose) (Relenza; GlaxoSmith Kline, Research Triangle Park, NC), and oseltamivir (5 mg/kg PO q12h × 10 days) (Tamiflu; Roche, Nutley, NJ). These have been shown to prevent and treat influenza infection and either agent may be used to greater effect in combination with amantadine.4,19,28 However, resistance to oseltamivir appears to be emerging among some influenza strains.52 Because ferrets are a good model with which to study influenza infection in people, they are frequently used in experimental studies to develop new anti-influenza drugs.63 Anti-influenza drugs used in humans may therefore be used to treat pet ferrets. Antibiotics can be used to control secondary bacterial infections of the respiratory tract. In neonates, death typically results from secondary bacterial infections; antibiotic therapy may thus reduce neonate mortality.25 The use of antipyretic drugs to control fever is of questionable merit because fever is an important host defense mechanism. In one study, ferrets given aspirin had lowered body temperature, but they shed more virus and their viral levels decreased less rapidly than those of ferrets not treated with an antipyretic.26 In a recent meta-analysis of the use of antipyretics in animal models of influenza virus, risk of mortality increased with the use of antipyretics (aspirin, paracetamol, and diclofenac).17 This suggests that fever is instrumental in restricting the severity of infection.26,53

PREVENTION Controlling influenza rests mainly on preventing exposure of susceptible ferrets to infected individuals. Newborn ferrets are protected from disease by milk-derived antibodies in immunized dams.27 Experimentally, ferrets remain resistant to infection from the same influenza strain 5 weeks after primary infection.21

Vaccinating ferrets against influenza virus is not ­generally r­ ecommended for several reasons. Influenza is a relatively benign disease in ferrets, and the wide antigenic variation of the virus makes vaccination difficult. Also, vaccination seems to confer only short-term immunity.21 However, if a vaccine is being given, the use of a live or recombinant rather than an inactivated vaccine should be considered, because they may induce a greater protective effect.18,33

PNEUMONIA Pneumonia is not a common diagnosis in ferrets. Viral causes of pneumonia include CDV and influenza virus. Aleutian disease virus, a parvovirus, is associated with interstitial pneumonia in mink kits1 and should be considered as a possible cause of pneumonia and dyspnea in ferrets, especially the young.60 Respiratory syncytial virus has been shown to cause rhinitis and infection in the lungs of ferrets, but clinical signs of pneumonia have not been seen.47 Pyogranulomatous pneumonia has recently been reported in association with a systemic coronavirus infection in ferrets; it appears to produce a disease syndrome similar to the dry form of feline infectious peritonitis (see Chapter 3).23,37,45 Bacterial pneumonia (Fig. 6-2) is characterized by a suppurative inflammatory process that affects the bronchial tree, the lung lobes, or both. Reported primary bacterial pathogens that cause pneumonia in ferrets are Streptococcus zooepidemicus, other streptococcal species, and numerous mycobacterial species.12,32,50,61 Gram-negative bacteria such as Escherichia coli, Klebsiella pneumoniae, and Pseudomonas aeruginosa have been isolated from ferrets.20 Other bacteria that have been isolated from the lungs of ferrets include Bordetella bronchiseptica and Listeria monocytogenes. An acute hemorrhagic syndrome has recently been described in young ferrets (8-24 weeks of age) and may result in interstitial pneumonia. Affected ferrets have a prolonged prothrombin time (PT) and activated partial thromboplastin time (APTT) when compared with unaffected ferrets.29 Pneumocystis carinii is known to infect the lungs of ferrets. Latent infections can become active with immune suppression.3,11 Diagnosis is based on identifying the organism in a tracheal or lung wash. Treatment recommendations for P. carinii

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SECTION I  Ferrets

Fig. 6-2  Bacterial pneumonia in a ferret. The lungs are diffusely congested with dark areas of consolidation.

pneumonia, based on those for dogs, include pentamidine isethionate or trimethoprim-sulfamethoxazole. Two cases of mild endogenous lipid pneumonia have been documented on histologic examination of ferrets at necropsy at the Animal Medical Center (New York, NY) (K. Quesenberry, personal communication, 2003), and one case of mortality due to endogenous lipid pneumonia was confirmed at necropsy (D. Perpinan, personal communication, 2009). In suspected cases, lung aspiration or bronchiolar lavage may help to provide antemortem diagnosis, although biopsy is needed for definitive diagnosis. Documented cases in rodents and other mustelids appear to be idiopathic or secondary to other disease processes.6 Successful treatment of lipid pneumonia has been achieved in people with prednisolone and thus may be a therapeutic option in ferrets.8 Although exogenous lipid pneumonia has not been documented in ferrets, caution should be exercised in treating animals with a mineral oil-based preparation for gastrointestinal disease (e.g., trichobezoar). Chronic aspiration of mineral oil products has been associated with lipid pneumonia in cats and people.13,39

HISTORY AND PHYSICAL EXAMINATION Ferrets with pneumonia exhibit typical clinical signs such as labored breathing, dyspnea, cyanotic mucous membranes, increased lung sounds, nasal discharge, fever, lethargy, and anorexia. Fulminant pneumonia leading to sepsis and death has been reported.20

DIAGNOSIS The diagnosis of pneumonia should be based on the clinical signs, radiographic findings, and results of supportive diagnostic tests. Results of the CBC may reveal leukocytosis caused by a neutrophilia with a left shift. In young ferrets with evidence of interstitial pneumonia, positive results of serologic tests and high concentrations of gamma globulins may support a diagnosis of Aleutian mink disease. Early in the disease, radiographs may show an interstitial pattern that changes to an alveolar pattern (Fig. 6-3) as the pneumonia progresses. If aspiration pneumonia is present,

Fig. 6-3  Ventrodorsal radiograph demonstrating an alveolar pattern and air bronchograms in a ferret with pneumonia. Photograph courtesy of Dr. Nico Schoemaker.

dependent lung lobes are primarily involved. Marked bronchial patterns suggest primary airway disease. Microbial cultures of tracheal or lung wash samples are invaluable in establishing a diagnosis and in treating ferrets with pneumonia. Submit samples for culture (aerobic or anaerobic bacterial, fungal, mycobacterial, or other) based on cytologic analysis of the collected fluid and debris. Cytologic assessment of tracheal wash samples from a ferret with pneumonia typically reveals septic inflammation and degenerating neutrophils. Results may also suggest the severity, cause, and chronicity of disease.

CHAPTER 6  Respiratory Diseases

83

TREATMENT Treat ferrets with pneumonia with good supportive care, including fluid therapy, force-feeding, and oxygen therapy as needed as well as with antimicrobials tailored according to test results. First-line antibiotics to consider before the results of culture and sensitivity testing are known are the quinolones, trimethoprimsulfamethoxazole, chloramphenicol, or the cephalosporins. Anecdotally, azithromycin at a dose of 5 mg/kg PO q24h also appears to be effective. In a report of two ferrets with mycobacterial pneumonia, both responded successfully to clarithromycin.32 Combination antibiotic therapy may be indicated. The prognosis depends on the cause of pneumonia and response to treatment. Most ferrets with bacterial pneumonia respond to antibiotic therapy and supportive care.

PREVENTION Bordetellosis is rare in ferrets. Nonetheless there is pervasive information in the lay literature about disease prevention. The best way to prevent B. bronchiseptica infection is to avoid hospitalizing ferrets where dogs, rabbits, or other common carriers are present. Anectdotally, a killed, injectable Bordetella bacterin may be effective in preventing bordetellosis in ferrets when used in accordance with manufacturer recommendations for dogs; however, no published reports supports this claim. The canine modified live intranasal Bordetella bacterin may cause disease in ferrets and is not recommended.

PULMONARY MYCOSES Pulmonary mycoses are uncommon in pet ferrets. Because ferrets in the United States are usually indoor pets, exposure to mycotic spores, which are mainly found in the soil, is unlikely.

HISTORY AND PHYSICAL EXAMINATION Not all animals with mycoses exhibit signs consistent with pulmonary disease. If lesions develop in the lungs, animals usually cough. Other signs consistent with a mycotic infection are wasting, lethargy, anorexia, lymph node enlargement, lameness, ocular and nasal discharge, and draining tracts unresponsive to antibiotic therapy.16,59 The prognosis for ferrets with pulmonary mycoses is poor.

CRYPTOCOCCOSIS Cryptococcosis, caused by Cryptococcus bacillisporus (formerly C. neoformans var gattii) and C. neoformans var grubii, has been diagnosed in a small number of ferrets.34,35 Infection can cause rhinitis, pneumonia, and pleuritis. Additionally, regional lymph node involvement is common and may also be expected to cause dyspnea when the retropharyngeal or mediastinal lymph nodes are involved.34 Invasive cryptococcal rhinitis has been successfully treated with itraconazole and surgical debulking.35

BLASTOMYCOSIS Blastomycosis, caused by Blastomyces dermatitidis, is endemic in the southeastern United States, the Mississippi River Valley, and the Ohio River Valley.59 Experimentally, the incubation period is 5 to 12 weeks. The mycelial phase is found in the soil, and

Fig. 6-4  The heart of a ferret that presented for moderate dyspnea and coughing. Necropsy demonstrated 3 female and 7 male Dirofilaria immitis worms in the heart.

the yeast form is found in the tissues. Diagnosis is made on the basis of a history of travel to an endemic region, clinical signs consistent with disease, results of cytologic assessment, positive periodic acid-Schiff reaction, or culture of B. dermatitidis. Amphotericin B and ketoconazole or itraconazole are recommended for treatment.59 Dosages should be based on those used for cats.

COCCIDIOIDOMYCOSIS Coccidioides immitis, the causative agent of coccidioidomycosis, is endemic in the southwestern United States and parts of Latin America. Primary infection develops after a susceptible host inhales the mycelia. Once in the host, spherules form and then produce endospores.16,59 Pulmonary signs develop 1 to 3 weeks after infection. Diagnosis of this disease is based on identifying the spherules on cytologic examination; they appear as refractile double-walled bodies.54 Recommended treatment, which is based on that for cats with coccidioidomycosis, includes the use of amphotericin B and ketoconazole or itraconazole.16,59

OTHER CAUSES OF RESPIRATORY SIGNS Differential diagnoses for tachypnea, dyspnea, and respiratory distress are similar to those for other small animals. After the history and physical examination, chest and abdominal radiography is the most important tool to differentiate the causes of lower respiratory tract symptoms. Ferrets that have severe traumatic injuries, such as from a fall from a great height, can develop pneumothorax or diaphragmatic hernia. These animals should be managed as one would a dog or cat with the same injuries. Ferrets with heartworm disease often present with coughing and tachypnea as the only clinical signs, even with moderate worm burdens (see Chapter 5) (Fig. 6-4).

84

SECTION I  Ferrets

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24. Greenacre CB. Incidence of adverse events in ferrets vaccinated with distemper or rabies vaccine: 143 cases (1995-2001). J Am Vet Med Assoc. 2003;223(5):663-665. 25. Husseini RH, Collie MH, Rushton DI, et al. The role of naturally-acquired bacterial infection in influenza-related death in neonatal ferrets. Br J Exp Pathol. 1983;64:559-569. 26. Husseini RH, Sweet C, Collie MH, et al. Elevation of nasal viral levels by suppression of fever in ferrets infected with influenza viruses of differing virulence. J Infect Dis. 1982;145:520-524. 27. Husseini RH, Sweet C, Overton H, et al. Role of maternal immunity in the protection of newborn ferrets against infection with a virulent influenza virus. Immunology. 1984;52:389-394. 28. Ilyushina NA, Hoffman E, Saloman R, et  al. Amantadineoseltamivir combination therapy for H5N1 influenza virus infection in mice. Antivir Ther. 2007;12(3):363-370. 29. Johnson-Delaney CA, Reavill DR. Ferret acute hemorrhagic syndrome, In Proceedings. Assoc Avian Vets and Assoc of Exot Mam Vets, 2008; 49–50. 30. Jóżwik A, Frymus T. Comparison of the immunofluorescence assay with RT-PCR and nested PCR in the diagnosis of canine distemper. Vet Res Commun. 2005;29(4):347-359. 31. Kang ES, Lee HJ, Boulet J, et  al. Potential for hepatic and renal dysfunction during influenza B infection, convalescence, and after induction of secondary viremia. J Exp Pathol. 1992;6:133-144. 32. Lunn JA, Martin P, Zaki S, Malik R. Pneumonia due to Mycobacterium abscesses in two domestic ferrets (Mustela putorius furo). Aust Vet J. 2005;83(9):542-546. 33. Mahmood K, Bright RA, Mytle N, et  al. H5N1 VLP vaccine induced protection in ferrets against lethal challenge with highly pathogenic H5N1 influenza viruses. Vaccine. 2008;26(42):5393-5399. 34. Malik R, Alderton B, Finlaison D, et  al. Cryptococcosis in ferrets: a diverse spectrum of clinical disease. Aust Vet J. 2002;80(12):749-755. 35. Malik R, Martin P, McGill J, et al. Successful treatment of invasive nasal cryptococcosis in a ferret. Aust Vet J. 2000;78(3): 158-159. 36. Marini RP, Adkins JA, Fox JG. Proven or potential zoonotic diseases of ferrets. J Am Vet Med Assoc. 1989;195:990-994. 37. Martinez J, Ramis AJ, Reinacher M, et  al. Detection of feline infectious peritonitis virus-like antigen in ferrets. Vet Rec. 2006;158:523. 38. McLaren C, Butchko GM. Regional T- and B-cell responses in influenza-infected ferrets. Infect Immunol. 1978;22:189-194. 39. Midulla F, Strappini PM, Ascoli V, et al. Bronchoalveolar lavage cell analysis in a child with chronic lipid pneumonia. Eur Respir J. 1998;11:239-242. 40. Moore GE, Glickman NW, Ward MP, et  al. Incidence of and risk factors for adverse events associated with distemper and rabies vaccine administration in ferrets. J Am Vet Med Assoc. 2005;226(6):909-912. 41. Ochi A, Danesh A, Seneviratne C, et al. Cloning, expression and immunoassay detection of ferret IFN-gamma. Dev Comp Immunol. 2008;32(8):890-897. 42. Patterson AR, Cooper VL, Yoon KJ, et  al. Naturally occurring influenza infection in a ferret (Mustela putorius furo) colony. J Vet Diag Invest. 2009;21(4):527-530. 43. Pearson RC, Gorham JR. Viral disease models. In: Fox JG, ed. Biology and diseases of the ferret. 2nd ed. Philadelphia: Lippincott Williams & Wilkins; 1998:487-498. 44. Perpinan D, Ramis A, Tomas A, et  al. Outbreak of canine distemper in domestic ferrets (Mustela putorius furo). Vet Rec. 2008;163(8):246-250. 45. Perpinan D, Lopez C. Clinical aspects of systemic granulomatous inflammatory syndrome in ferrets (Mustela putorius furo). Vet Rec. 2008;162:180-184.

CHAPTER 6  Respiratory Diseases 46. Peltola VT, Boyd KL, McAuley JL, et al. Bacterial sinusitis and otitis media following influenza virus infection in ferrets. Infect Immunology. 2006;74(5):2562-2567. 47. Prince GA, Porter DD. The pathogenesis of respiratory syncytial virus infection in infant ferrets. Am J Pathol. 1976;82:339-352. 48. Rarey KE, DeLacure MA, Sandridge SA, et  al. Effect of upper respiratory infection on hearing in the ferret model. Am J Otolaryngol. 1987;8:161-170. 49. Ryland LM, Gorham JR. The ferret and its diseases. J Am Vet Med Assoc. 1978;173:1154-1158. 50. Saunders GK, Thomsen BV. Lymphoma and Mycobacterium avium infection in a ferret (Mustela putoris furo). J Vet Diagn Invest. 2006;18(5):513-515. 51. Shin YJ, Cho KO, Cho HS, et al. Comparison of one-step RTPCR and a nested PCR for the detection of canine distemper virus in clinical samples. Aust Vet J. 2004;82(1-2):83-86. 52. Sy CL, Lee SS-J, Liu M-T, et  al. Rapid emergence of oseltamivir resistance [letter]. Emerg Infect Dis. Apr 2010;16(4). http:// www.cdc.gov/EID/content/16/4/723.htm. Accessed 1/12/2011. 53. Smith H, Sweet C. Lessons for human influenza from pathogenicity studies with ferrets. Rev Infect Dis. 1988;10:56-75. 54. Smith W, Stuart-Harris CH. Influenza infection of man from the ferret. Lancet. 1936;228:121-123. 55. Spickler AR. Influenza. Technical Fact Sheet. The Center for Food Security and Public Health, Iowa State ­University. Available at http://www.cfsph.iastate.edu/DiseaseInfo/disease. php?name=influenza&lang=en. Accessed 1/12/2011. 56. Squires S, Belyavin G. Free contact infection in ferret groups. J Antimicrob Chemother. 1975;1:35-42.

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57. Swango LJ. Canine viral diseases. In: Ettinger SJ, ed. Textbook of veterinary internal medicine. Philadelphia: WB Saunders; 1989:298-311. 58. Swenson SL, Koster LG, Jenkins-Moore M, et al. Natural cases of 2009 pandemic H1N1 influenza A virus in pet ferrets. J Vet Diagn Invest. 2010;22(5):784-788. 59. Taboada J. Systemic mycoses. In: Ettinger SJ, Feldman EC, eds. Textbook of veterinary internal medicine. 5th ed. Philadelphia: WB Saunders; 2000:453-476. 60. Une Y, Wakimoto Y, Nakano Y, et  al. Spontaneous Aleutian disease in a ferret. J Vet Med Sci. 2000;62(5):553-555. 61. Valheim M, Djonne B, Heiene R, Caugant DA. Disseminated Mycobacterium celatum (Type 3) infection in a domestic ferret (Mustela putorius furo). Vet Pathol. 2001;38:460-463. 62. Van Riel D, Munster VJ, McAuley AL, et al. Human and avian influenza viruses target different cells in the lower respiratory tract of humans and other mammals. Am J Pathol. 2007;171(4):1215-1223. 63. Yoshimoto J, Yagi S, Ono J, et al. Development of anti-­influenza drugs: II. Improvement of oral and intranasal absorption and the anti-influenza activity of Stachyflin derivatives. J Pharm Pharmacol. 2000;52:1247-1255. 64. Welter J, Taylor J, Tartaglia J, et al. Vaccination against Canine Distemper Virus Infection in Infant Ferrets with and without Maternal Antibody Protection, Using Recombinant Attenuated Poxvirus Vaccines. J Virol. 2000;74(14):6358-6367.

CHAPTER

7

Endocrine Diseases

Karen L. Rosenthal, DVM, MS, and Nicole R. Wyre, DVM, Diplomate ABVP (Avian)

Adrenal Gland Disease History and Physical Examination Clinical Pathology and Diagnostic Testing Possible Concurrent Abnormalities Management Adrenal Histopathology Prognosis Pheochromocytomas Thyroid Disease Diabetes Mellitus History and Physical Examination Clinical Pathologic and Diagnostic Testing Treatment Prognosis Pancreatic Islet Cell Tumors Etiology Pathophysiology Clinical Features Clinical Pathologic Abnormalities Differential Diagnoses for Fasting Hypoglycemia Diagnostic Approach Diagnostic Imaging Management of Islet Cell Tumors Histopathology Prognosis

ADRENAL GLAND DISEASE Adrenocortical disease has been recognized for almost 25 years as a common malady affecting pet ferrets both in the United States and in many countries around the world.11,20,25,35,39,53, 54,58,68,72,91,93,109 It is typically seen in middle-aged to older ferrets and is most commonly characterized by hair loss in both sexes and by vulvar enlargement in females.35,58,91 Clinical signs of adrenocortical disease in ferrets differ from those of classic Cushing’s disease in dogs; moreover, plasma cortisol 86

concentrations are rarely increased in ferrets. Instead, the concentrations of estradiol, 17-hydroxyprogesterone, or one or more of the plasma androgens may be increased as a result of adrenocortical hyperplasia, adenoma, or adenocarcinoma.93 There has been much speculation regarding the underlying cause of the pathologic changes in the adrenal glands of these ferrets. Suggested causes include early neutering, genetic predisposition, light-dark cycle disruptions, and diet.12,13,42,46,85,97,98 The role of early neutering has garnered the most attention and study. Historically, gonadectomy at an early age in some strains of mice was observed to lead to adrenocortical nodular hyperplasia or neoplasia of one or both adrenal glands. These glands hypersecrete estrogens or androgens.30,70,99 Like mice that undergo gonadectomy early in life, most commercially raised ferrets in the United States undergo ovariohysterectomy or castration before they are 6 weeks of age. Another possible explanation is not necessarily the age of neutering but the time period between neutering and the onset of disease.97 To this end, gonadotrophic hormones appear to play a role in the pathogenesis of hyperadrenocorticism in ferrets.98 Specifically, this condition has been defined as a disease resulting from the expression of luteinizing hormone (LH) receptors on adrenocortical cells that produce sex steroids. The speculative pathogenesis is that, after neutering, LH and follicle stimulating hormone (FSH) persistently stimulate the adrenal cortices as a result of the loss of negative gonadal feedback on hypothalamic gonadotropin releasing hormone (GnRH), resulting in adrenocortical hyperplasia. To further support this hypothesis, LH receptors that could trigger abnormal adrenal gland growth have been found in the adrenal glands of normal ferrets.97 In a small study of adult ferrets, there was a high incidence of adrenal gland disease during a 1- to 7-year period after surgery.42 These results may be further evidence that adrenal gland disease is influenced more by lack of negative feedback than the practice of early neutering.12 As the ability to study disease at the molecular level becomes more robust, the underlying pathogenicity of adrenal gland disease in ferrets will become more evident.77 Other possible causes of adrenal gland disease in ferrets may be related to husbandry. One speculated risk factor for adrenal Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

CHAPTER 7  Endocrine Diseases gland disease is the artificial light:dark cycle to which indoor ferrets are exposed.46,69,85 The long light cycle, as would be present with ferrets living indoors, stimulates the release of GnRH and LH while simultaneously inhibiting the production of melatonin. The increased LH concentrations combined with decreased circulating melatonin (a known antigonadotropic hormone) stimulate receptors in adrenal gland tissue, leading to adrenal gland disease.85 Therefore housing ferrets indoors may be a contributing risk factor to the pathogenesis of adrenal gland disease.69,85

HISTORY AND PHYSICAL EXAMINATION Progressive alopecia is the most common historic finding. Hair loss typically begins in the late winter or early spring and may continue until the ferret is partially or completely bald. Occasionally the hair coat may regrow fully during the fall. During the next winter or spring, alopecia commonly begins again. This sequence can recur over a period of 2 to 3 years until the hair does not regrow. Spayed female ferrets with adrenocortical disease frequently have a history of vulvar enlargement with or without a mucoid discharge. Male ferrets may have a history of dysuria, urinary blockage, or increased aggressive behavior. In at least one study, most ferrets reported with adrenocortical disease were female.91 Many owners are aware that an enlarged vulva in a female ferret is a cause for concern. The vulva enlarges normally during estrus, and many owners know that prolonged estrus can result in estrogen-induced bone marrow toxicosis. Thus although the disease is reported more frequently in females than in males, this may be caused by a presentation bias rather than an actual sex predilection and may not reflect the true incidence of disease.11 In one study, the average age at which signs of adrenal disease were first observed by ferret owners was approximately 3.5 years.91 A smaller study has reported an older age distribution for this disease.72 Alopecia is the most common clinical manifestation of adrenocortical disease and develops in both male and female ferrets (Fig. 7-1). More than 90% of ferrets with adrenal gland disease have some hair loss. The hair epilates easily. Alopecia is usually symmetrical, beginning on the rump, the tail, or the flanks and

Fig. 7-1  A 4-year-old female spayed ferret with extensive alopecia as a result of adrenal gland disease. Hair was first noticed to be thinning one year earlier.

87

progressing to the lateral trunk, dorsum, and ventrum. These patterns of alopecia should be documented in the medical record for comparison as the disease progresses. In more than one-third of ferrets with adrenal gland disease, owners report the ferret as pruritic. Although pruritus usually accompanies hair loss, in some ferrets pruritus is the only clinical sign. Pruritus is most frequently observed on the dorsum between the shoulder blades. The skin is often erythematous in these areas. More than 70% of female ferrets with adrenal gland disease have an enlarged vulva (Fig. 7-2).91 The vulva can be slightly to grossly enlarged and become turgid and edematous, resembling the vulva of a jill in estrus. A seromucoid discharge may be present, and results of cytologic examination may show localized vaginitis. The perivulvar skin may appear dark and bruised. Partial or complete urinary blockage in male ferrets is occasionally associated with adrenocortical disease.20,88 Affected ferrets have dysuria or stranguria. Periurethral cysts develop in the region of the prostate (possibly originating from hormoneresponsive cells) and cause urethral narrowing. Because of the urethral narrowing, passing a urinary catheter into the bladder of these ferrets can be difficult (see Chapter 2). A ferret with a urethral blockage may have a life-threatening metabolic derangement; therefore these cases usually constitute emergencies. In some ferrets, removing the diseased adrenal tissue and draining the cysts resolves the urinary blockage within 1 or 2 days. In others, the prostatic tissue is infected and aggressive surgical treatment along with antibiotic and hormonal therapy is necessary (see Chapter 11). On physical examination, enlarged adrenal glands are sometimes palpable (Fig. 7-3). The left adrenal gland is more easily identified than the right. The left gland is usually engulfed in a large fat pad cranial to the left kidney. It may feel like a small, firm, round mass. Because the right gland has a more cranial location and is under a lobe of the liver, it is more difficult to palpate. Enlarged mesenteric lymph nodes may be palpable. The spleen may be palpably enlarged, but it usually has smooth borders and is not painful. In some instances, the texture of the spleen is irregular and knobby. In most older ferrets, an enlarged spleen is an incidental finding on physical examination.

Fig. 7-2  An enlarged vulva in a female ferret with adrenal gland disease.

88

SECTION I  Ferrets Abdominal ultrasound is also useful for detecting concurrent diseases, such as renal or hepatic disease, metastasis from a pancreatic insulinoma, or enlarged lymph nodes. The presence of prostatic enlargement or uterine disease associated with adrenal gland disease can also be found by ultrasonography.11 Although not done routinely, advanced imaging studies such as computed tomography (CT) or magnetic resonance imaging (MRI) of a ferret’s abdomen can demonstrate adrenal gland abnormalities. In ferrets with right adrenal gland tumors, CT or MRI may be useful as a presurgical screening tool in determining involvement or invasion of the vena cava.9

Ancillary Diagnostic Tests

Fig. 7-3  A female spayed ferret with confirmed adrenal gland disease. The left adrenal gland was palpable on physical examination and was prominent when the ferret was in dorsal recumbency.

CLINICAL PATHOLOGY AND DIAGNOSTIC TESTING The presumptive diagnosis of adrenal gland disease is based on history, clinical signs, and results of imaging techniques and steroid hormonal assays. Diagnosis is confirmed by histologic examination of adrenal tissue obtained during surgical biopsy or adrenalectomy. Results of the complete blood count (CBC) are usually unremarkable. Rarely adrenocortical disease is associated with nonregenerative anemia. If disease is severe or prolonged, pancytopenia may rarely be present. These changes mimic those in ferrets with estrogen-induced bone marrow toxicosis (see Chapter 4). In ferrets with anemia and pancytopenia, a packed cell volume less than 15% carries a grave prognosis. Results of the biochemical profile are also usually within reference intervals. The alanine aminotransferase concentration is occasionally high, but the association of this finding with adrenal gland disease is unknown. Because insulinomas are also common in older ferrets, hypoglycemia from a pancreatic beta cell tumor (insulinoma) may be present. A urinalysis is not helpful in diagnosing adrenal gland disease. Radiographs are generally not helpful in diagnosing this disease. An enlarged adrenal gland rarely displaces other organs or calcifies; therefore organ displacement and mineralized glands are not visible radiographically. Lung metastasis of adrenal gland tumors is very rare. However, radiographs are useful as a screening tool for other conditions, such as heart disease or splenomegaly. Abdominal ultrasound is useful for detecting abnormal adrenal glands. The size, side of enlargement, and architecture can often be determined.11,53,54,71,74 In one study in normal ferrets, the mean dimensions (length and width) of the right adrenal gland were 7.6 + 1.8 by 2.6 + 0.4 mm, and those of the left adrenal gland were 7.2 + 1.8 by 2.8 + 0.5 mm.74 In another study, the mean dimensions of 28 abnormal left adrenal glands (length and thickness) were 9.2 ± 3.2 by 6.3 ± 3.0 mm, and dimensions of 19 abnormal right adrenal glands were 8.5 ± 2.5 by 5.2 ± 3.3 mm.53 In the same study, abnormal adrenal glands appeared on ultrasound as rounded with an enlarged cranial or caudal pole or both, a heterogeneous structure, and increased echogenicity; they were either with or without mineralization.53

Several diagnostic tests used in dogs with adrenocortical disease are not useful in ferrets. The adrenocorticotropic hormone (ACTH) stimulation test and the dexamethasone suppression test cannot be used to diagnose adrenal disease in ferrets. Both normal ferrets and those with adrenal disease respond equally well to an ACTH stimulation test, probably because most ferrets with this disease do not produce abnormally high concentrations of cortisol.90 Plasma concentrations of ACTH and alpha-melanocyte-stimulating hormone in ferrets with adrenal disease are similar to those of normal ferrets, suggesting that adrenal disease in ferrets is independent of ACTH and alphamelanocyte-stimulating hormone.96 Urinary cortisol/creatinine ratios were higher in 12 ferrets with adrenocortical tumors than in 51 clinically normal ferrets.39 In dogs, the urinary cortisol/ creatinine ratio is a sensitive but not a specific indicator of hyperadrenocorticism. Further studies are needed to evaluate urinary cortisol/creatinine ratios in ferrets with diseases other than hyperadrenocorticism. The measurement of serum or plasma concentrations of steroid hormones is a reliable means of diagnosing adrenal disease in ferrets. Hormone panels that measure estradiol, androstenedione, and 17-hydroxyprogesterone in blood samples are commercially available (Clinical Endocrinology Laboratory of the Department of Comparative Medicine at the University of Tennessee [http://www.vet.utk.edu/diagnostic/endocrinology/ index]). In a normal neutered ferret, these steroids are found in minute quantities, whereas in ferrets with adrenal disease, the serum concentrations of one or more of these compounds may be high.89 In Table 7-1, reference intervals are given for androstenedione, dehydroepiandrosterone sulfate, estradiol, and 17-hydroxyprogesterone in intact female and neutered ferrets.

Differential Diagnosis A ferret with an ovarian remnant or an intact female ferret may present with an enlarged vulva and alopecia, resembling a ferret with adrenal disease. However, ferrets with ovarian remnants are uncommon and tend to exhibit an enlarged vulva at an earlier age (first onset of estrus at one year of age) compared with the usual age of onset of adrenal disease (3-4 years of age). Several diagnostic methods can be used to differentiate the two conditions. In one method, human chorionic gonadotropin (HCG) 100 IU IM is administered and then repeated in 7 to 10 days. If the ferret is intact or if an ovarian remnant is present, the vulva usually decreases in size. Alternatively, imaging methods such as abdominal ultrasound or measuring steroid hormone concentrations may help differentiate the two conditions. If the concentrations of androgens—such as androstenedione, dehydroepiandrosterone sulfate, or 17-hydroxyprogesterone— are high, an adrenal tumor is likely. If only the estradiol

CHAPTER 7  Endocrine Diseases

89

  Table 7-1    Serum Concentrations of Steroid Hormones in Intact and Neutered Normal Ferrets and Ferrets with Adrenal Disease NEUTERED FERRETSa

INTACT FEMALE FITCH FERRETS (N = 11)c

NORMAL FERRETS (N = 26) (13 MALE, 13 FEMALE) Steroid Hormone Androstenedione (nmol/L) Dehydroepiandrosterone sulfate (mmol/L) Estradiol (pmol/L) 17-Hydroxyprogesterone (nmol/L)

Adrenal Disease Mean 67 (n = 25) 0.03 (n = 27) 167 (n = 28) 3.2 (n = 20)

Mean 6.6 0.01 106 0.4

Reference Intervalb

Mean

18%, with indigestible fiber at >12.5%.27

FAT Fat provides a noncarbohydrate source of energy and improves palatability. It may also help to decrease dustiness and crumbling of pellets.11 Pet rabbits, however, are prone to obesity and hepatic lipidosis; therefore high-fat diets must be avoided. Dietary fat levels of 2.5% to 4% are considered appropriate. Some laboratory strains of rabbit are prone to the development of arteriosclerosis when placed on diets high in fat.9

VITAMINS AND MINERALS Vitamin and mineral requirements of the rabbit are summarized in Table 14-1.

DIETARY COMPONENTS GRASS

Simple sugars and starches are utilized as an energy source, but care must be taken to ensure that excessive levels are not fed. Because of the rabbit’s rapid gut transit time, starch and simple sugars may be incompletely digested in the small intestine, resulting in these compounds being directed into the cecum, where they may be used as substrates for fermentation by the cecal microorganisms. Carbohydrate overload in the cecum predisposes to enterotoxemia, especially in young animals.41 Owing to this potential for incomplete starch digestion, lowenergy grains such as oats are preferred over corn or wheat,12 and they should not be processed too finely.

Grass provides a balanced source of digestible and indigestible fiber, protein, vitamins, and minerals. In addition, grass is highly abrasive because of silicates and other materials that are present to varying degrees, depending on the species of grass and growing conditions. This abrasive factor is important for normal tooth wear.13 Generally crude fiber of fresh grass varies from 20% to 40%, while the protein content tends to be 15% to 19%.41 As a rule, the fiber content is higher in lower-protein grass and vice versa. Where practical, it is recommended that pet rabbits be given access to grass for at least several hours a day. Grass must be either grazed or fed freshly cut. Obviously this is not possible in all situations (e.g., for rabbits living in apartments in urban environments). Lawn clippings should not be used, as they ferment rapidly and may cause digestive disturbances.

FIBER

HAY

Fiber is a vital component of the pet rabbit’s diet. It is essential for the maintenance of gastrointestinal health and to promote normal dental attrition. It is also believed to prevent behavioral

Hay is considered an essential part of the pet rabbit’s diet and should be available ad libitum to rabbits that do not have free access to grass for grazing. For those that do have access to grass

CARBOHYDRATE

CHAPTER 14  Gastrointestinal Physiology and Nutrition

189

  Table 14-1    Summary of Vitamin and Mineral Requirements of the Domestic Rabbita Vitamin/Mineral

Dietary Requirement

Comments

Vitamin A

10,000-18,000 IU/kg

Vitamin B complex

Pet rabbits consuming fresh grass and vegetables are unlikely to be deficient; rabbits housed indoors and fed a diet of cereal mixtures and poor-quality hay without fresh grass or greens are potential candidates for deficiency (as cereals other than corn are a poor source); deficiency is associated with enteritis, retarded growth, weight loss, neurologic symptoms, keratitis, iridocyclitis, hypopyon, blindness, abortion, low fertility, stillbirth, neonatal death, and fetal malformations; excessive levels can lead to abortion, low fertility, and fetal malformations as well as hyperostotic polyarthropathy. Produced in the cecum; supplied by the cecotrophs.

None if animal is capable of cecotrophy 10-50 ppm recommended if Synthesized endogenously, but requirements may increase during times of stressed stress. Ideally provided by exposure to Synthesized in skin following exposure to sunshine and/or provided by diet; sunlight rather than dietary dietary sources for rabbits include sun-dried hay and supplemented supplementation; published pellets; although rabbits are able to absorb calcium without vitamin D, requirement is 800-1200 vitamin D will improve absorption if dietary calcium levels are low; excess IU/kg levels (>2300 IU/kg) have been associated with fetal mortality, depressed appetite, diarrhea, ataxia, paralysis, calcification of soft tissues, and death; deficiency has been associated with hypophosphatemia and osteomalacia. 50 mg/kg Deficiency is associated with muscular dystrophy, infertility, abortions, stillbirths, and increased susceptibility to coccidiosis. Unknown, but dietary Deficiency unlikely in this species; synthesized by cecal bacteria; most supplementation should not commercial pellets/mixes contain 1-2 mg/kg; consider supplementation be necessary if animal is in cases of coccidiosis, concurrent use of sulfa drugs, especially in capable of cecotrophy pregnant does and in situations where cecotrophy is inhibited. 0.5%-1.0% Unlike most other mammals, rabbits do not control calcium uptake from the gut; calcium uptake is not dependent on vitamin D; therefore serum levels of calcium increase directly in relation to increased dietary intake and excess calcium is excreted in the urine; availability of calcium for absorption can be related to other compounds found in food (i.e., phytates, oxalates, and acetates form complexes with calcium that reduce availability); deficiency can lead to a loss of appetite, tetany, muscle tremors, and death, especially in breeding does, and is thought to contribute to poor bone and dental health; excess calcium can result in urolithiasis, renal disease, and calcification of soft tissues (e.g., aorta and kidneys). 0.17%-0.32% — 1.0 ppm — 5-20 ppm — 0.4-2 ppm — 30-100 ppm — 0.3% Deficiency results in poor growth, alopecia, hyperexcitability, convulsions, myocardial fibrosis, and fur chewing. 8-15 ppm — 0.4%-0.8% Calcium-to-phosphorus ratio of 1:1 to 2:1 or higher is recommended; inverse Ca:P ratios will result in decreased bone density if overall phosphorus levels rise above 1%; availability is influenced by the presence of plant phytates and phytases; deficiency leads to rickets (in growing animals) and osteomalacia (in adults). 0.6% — 0.05 ppm Does not seem to have a sparing effect on vitamin E requirements and does not seem to be involved in white muscle dystrophies, as in other species. 0.2%-0.25% — 0.3%-0.6% Deficiency leads to immunosuppression, failure to gain weight, increased mortality, and increased severity of induced herpes simplex keratitis in experimental animals.

Vitamin C Vitamin D

Vitamin E Vitamin K

Calcium

Chloride Cobalt Copper Iodine Iron Magnesium Manganese Phosphorus

Potassium Selenium Sodium Zinc

aData

drawn from references 6, 7, 11, 12, 14, 17, 21, 27, 33, 36, 39, and 41.

190

SECTION II  Rabbits

on a daily basis, hay can still be used to support the diet, especially at times when grass availability is poor. Hay is produced from a variety of pasture species; its quality and availability vary markedly between regions, seasons, and even individual farms. Grass or meadow hays are preferred and may be derived from timothy, prairie, brome, ryegrass, fescues, meadow grass, and orchard grass. Timothy hay is packaged in small quantities by several feed companies (e.g., Oxbow Pet Products, Murdock, NE; Kaytee Products, Inc., Chilton, WI). The fiber content of grass hays is usually around 30% to 35% and protein ranges from 6.3% to 16.7%.40 Alfalfa (lucerne) and to a lesser extent other legumes such as clover are used widely in some regions for haymaking. The grain hays, primarily oats and barley, are also available in some areas. Leguminous hays tend to be high in protein (16.5%), calcium (1.5%), and energy and thus are very useful for growing rabbits; however, legume-rich diets are thought to predispose to obesity and urolithiasis in mature nonbreeding animals. Prolonged storage of hay (especially in warm conditions) can lead to loss of nutrients such as vitamins A and D. Good hay is sweet-smelling and should not be musty. Hay can be fed from racks or nets to increase time spent feeding and thereby reduce boredom. Straw is not recommended because, although rabbits will eat it, it is low in nutrients and will lead to deficiencies if it constitutes a major part of the diet.41 The feeding of silage is generally not practiced, although it has been investigated in some countries in farmed rabbits. The high moisture content of rye and timothy grass silage was found to restrict dry matter intake and lead to a lower rate of growth.43 There is little information available at present regarding the feeding of silage to pet rabbits.41 Anecdotal reports on the use of artificially dried grass in rabbits sound promising.41 The product is palatable and, provided that it is not stored inappropriately or for too long, the nutrient content should be superior to that of sun-dried hay except for vitamin D, which is produced via the irradiation of sterols following exposure to ultraviolet radiation in sunlight.

FRESH VEGETABLES AND EDIBLE PLANTS (“GREENS”) The provision of a variety of fresh vegetables (green leafy varieties) and edible plants is recommended for all rabbits. These may provide vital micronutrients and, because of their high water content, are unlikely to contribute significantly to caloric intake. Their fiber content alone is insufficient to meet the needs of the rabbit’s gastrointestinal tract. These feed items may be bought commercially or grown fresh/harvested from the wild. Fresh greens, especially novel items that have not been given before, should be introduced to a rabbit’s diet gradually to allow the gut microbes to adapt, thereby avoiding gastrointestinal upset. Suitable vegetables include collard, mustard, and dandelion greens; carrot, beet, and broccoli tops; alfalfa sprouts and clover; herbs such as parsley, cilantro, and basil; lettuce, broccoli, cauliflower, green peppers, chicory, chard, watercress, celery leaves, endive, raddichio, bok choy, dock, spring greens, kale, and cabbage. Wild plants that can be offered include bramble, dandelion, chickweed, plantain, sunflower, wild strawberry, dock, and yarrow.49 As a rough guide, 2 cups of varied fresh vegetables and/or edible plants daily is considered appropriate for a 2.3-kg (5-lb) rabbit. Fruit should be used sparingly if at all because of the high sugar content and potential for carbohydrate overload in the hindgut:

a small amount (e.g., up to 1 tablespoon for a 2.3-kg [5-lb] rabbit) as a treat several times a week is unlikely to be problematic. Perishable vegetables, greens, and fruits must be stored in a refrigerator. Rinsing in fresh water before feeding is advised to remove any contaminants (fertilizers, insecticides, feces, urine) from the outer surfaces of the food.

COMMERCIAL MIXES AND PELLETS Commercial rabbit food has traditionally been sold in one of two forms: pelleted diets and mixed rations. Pellets are homogenous in appearance and are usually forage and/or cereal-based with a number of nutritional additives. Mixed rations (sometimes referred to as “muesli-type” feeds) consist of a variety of ingredients such as flaked, micronized, or rolled cereals, extruded biscuits, grass-based pellets, and plant stems. These two broad types of commercially prepared diets have been used for many years for commercial production and in laboratory rabbit husbandry and more recently have been widely marketed for use in pet rabbits. Despite marketing claims, these commercial concentrate rations are not essential components of the adult pet rabbit’s diet. If ad libitum hay, grass, and a variety of greens are available and cecotroph intake is normal, the diet is essentially balanced and contains sufficient energy for maintenance requirements. Nevertheless, many owners like to feed these products for convenience and, if used correctly as a supplement to the dietary mainstays of grass/hay and greens, they can be useful in the provision of some micronutrients and extra energy when required. While they should be fed in only limited quantities to adult nonbreeding pet rabbits, concentrates may play an important role in the nutrition of growing, pregnant, lactating, and diseased rabbits. Similarly, they can be used to ensure that nutrient requirements are fulfilled in rabbits that are unable to consume significant amounts of hay and/or green vegetables or those that are unable to practice cecotrophy (e.g., rabbits with advanced acquired dental disease or debilitating spinal issues). The nutritional composition of commercial feeds varies enormously, and this is true of both pellets and mixed rations. Both are calorie-rich in comparison with grass, hay, and greens. If used as part of the diet, some attention should be paid to the fiber and protein content. Commercial feeds with high protein levels, used for maximal growth and weight gain in rabbits raised for meat production, often have a lower fiber content in order to increase palatability. These foods, especially when fed in large quantities, can be associated with the development of gastrointestinal diseases. Of the two broad types of commercial feeds, rabbits tend to be better nourished when fed pellets. This is because they are unable to feed selectively12; therefore they ingest a consistent ration. Pet rabbits offered mixed-ration diets tend to favor flaked peas and corn, which are high in starch and low in calcium and fiber.26 Because most owners tend to replenish the feed bowl regularly, discarding uneaten items, the complete mixture of ingredients is rarely consumed. Additionally, some ingredients in the mixed rations (e.g., locust beans) have been implicated in cases of intestinal obstruction.27 Owner perception is an important factor in pet nutrition and many owners become easily misguided by the appearance or even the packaging of certain concentrate foods. Pelleted feeds should be hard and durable32 and be relatively high in fiber (ideally > 18%). Extruded diets are now very popular for pet rabbits, incorporating long fiber particles without the pellet becoming friable. Pelleted diets formulated from timothy

CHAPTER 14  Gastrointestinal Physiology and Nutrition hay (e.g., Bunny Basic/T, Oxbow Pet Products; Forti-Diet Rabbit Timothy Blend Adult Maintenance Formula, Kaytee Products, Inc.) are commercially available for adult pet rabbits. Ideally the indigestible fiber particles within the pellet should be 0.5 mm in length to stimulate gut motility. Small particles ( 9). Hematuria was the most commonly reported abnormality on urine sediment, followed by mucus and lipid droplets.20 Calcium carbonate, calcium oxalate, and struvite crystals are all commonly seen on sediment examinations but, as in other small animals, crystal type(s) may not predict the mineralogy of the calculi.20 Prior to antibiotic use, culture the urine or bladder wall if high numbers of red or white blood cells, bacteria, or a combination of these are present on examination of the sediment. Collect urine via cystocentesis, as free-catch samples are commonly contaminated. Even gentle expression of the bladder can lead to hematuria. Calculi containing calcium carbonate require specific methodologies to differentiate calcium carbonate crystals from calcium oxalate monohydrate (COM) crystals; therefore the laboratory chosen for analysis must be able to perform these differentiating methods.20 Calcium carbonate calculi are not found in humans; thus human and some veterinary (those that do not commonly evaluate large/exotic animal samples) laboratories do not use techniques for differentiating calcium oxalate and calcium carbonate. Confirm the lab’s ability to perform these techniques in advance to ensure that the calculus is appropriately evaluated for the guinea pig patient.

Medical treatment of urolithiasis has been unrewarding to date, and surgical removal of the stones is most often required (see Chapter 25). In females, with the patient under deep sedation and analgesia, gently flush the bladder using a 3.5-Fr red rubber catheter to attempt to remove the calculi. Catheterizing boars is much more dangerous because of the small size of the urethra. Submit the calculi for analysis and culture and submit the bladder wall for culture and sensitivity. Postoperative management includes assisted feeding, fluids, analgesics, and antibiotics based on culture and sensitivity results. However, recurrence of the disease is common. Prevention is targeted at increasing water intake and reducing (but not eliminating) dietary calcium. We have seen metabolic bone disease (fibrous osteodystrophy; see “Musculoskeletal Diseases,” below) induced in guinea pigs with severe dietary calcium restriction for prevention of urinary calculi. For most animals, water is the cornerstone of any prevention protocol. Hydrochlorothiazide (2 mg/kg q12h) is a thiazide diuretic that reduces urinary Ca2+, K+, and citrate. It is unknown if diuresis would be of benefit to guinea pigs whose urine is already considered isosthenuric.20 Do not use hydrochlorothiazide with severe renal disease or fluid imbalances. Avoid alfalfa-based diets. Diets containing a high percentage of timothy, oat, or grass hays, a lower overall percentage of pellets, and a wide variety of vegetables and fruits decrease the risk of urolith development in pet guinea pigs.19 It is possible that dietary inhibitors of calcium are found in greater concentration in hays than in pellets. Urinary acidifiers were historically recommended based upon an assumed diagnosis of calcium oxalate calculi, but the normally alkaline urine of guinea pigs making the use of dietary acidifiers concerning. Potassium citrate (30-75 mg/kg PO q12h) binds urinary Ca2+, reduces ion activity, and alkalinizes the urine. Because guinea pigs have alkaline urine even with disease, the efficacy of this treatment is unclear. Hyperkalemia occurs, so monitor plasma K+ closely during treatment. According to anecdotal reports,

CHAPTER 23  Disease Problems of Guinea Pigs

301

potassium citrate and hydrochlorothiazide have been used together with some clinical success.

CYSTITIS AND URINARY TRACT INFECTIONS It has been reported that female guinea pigs are more vulnerable to bacterial cystitis because the urethral opening is closer to the anus, potentially allowing gastrointestinal bacteria to ascend the urethra into the urinary bladder.40 Bacterial cystitis is frequently identified concomitantly with urolithiasis, but with an equal gender distribution.20 Clinical signs mimic those of urolithiasis, with vocalizing/straining during urination, pollakiuria, dysuria, hematuria, anorexia, and depression. Base your diagnosis and treatment on urinalysis, urine (obtained via cystocentesis) culture and sensitivity results, and radiographs to rule out urolithiasis. Provide analgesics. Gently flush the sow’s bladder using a 3.5-Fr red rubber catheter under deep sedation and analgesia if warranted. Catheterization is not recommended for boars because of the small size of the urethra.

CHRONIC INTERSTITIAL NEPHRITIS AND CHRONIC RENAL FAILURE Chronic interstitial nephritis is commonly found in guinea pigs more than 3 years of age. Guinea pigs more than 1 year of age can develop some degree of renal segmental fibrosis, progressing in the aged animal to chronic renal failure. The pathogenesis of this disease is still unclear. Obstructive urolithiasis and pyelonephritis can lead to interstitial nephritis. It has been reported in guinea pigs with diabetes mellitus and with staphylococcal pododermatitis with chronic renal amyloidosis and nephritis as sequelae.38 Encephalitozoon cuniculi has been shown to cause subacute interstitial nephritis in laboratory animals, but its significance in pet guinea pigs is unknown. Contrary to previous literature, advanced cases present clinically with evidence of renal compromise such as polyuria/polydipsia (PU/PD), azotemia, chronic weight loss, cardiac compromise including dyspnea and secondary heart failure due especially to uremia, and other vague clinical signs such as anorexia or diarrhea/loose stool. Elevations in blood urea nitrogen (BUN) and creatinine, changes in electrolytes and minerals, isosthenuria, and nonregenerative anemia have all been recorded antemortem.22 Typical histopathological changes include interstitial fibrosis, glomerular ectasia, and sclerosis with variable numbers of mononuclear inflammatory cells. The renal parenchyma can be completely distorted, often with no recognizable normal tissue. Cardiac lesions such as epicarditis, myocarditis and fibrosis, pericarditis, and ventricular dilation are often identified concurrently with renal lesions. While chronic interstitial nephritis may be prevented by proper husbandry and sanitation to reduce the guinea pig’s susceptibility to pododermatitis,22,38 appropriate hydration throughout the guinea pig’s lifetime is also prudent.

OTHER UROPATHIES Renal cysts are not uncommon and are usually identified incidentally on necropsy. There is one report of renal failure in a guinea pig following ingestion of peace lily, with similar clinical signs as are seen in other mammals.24 Klossiella cobayae, a renal coccidian, lives in the epithelial cells lining the renal tubules, and sporocysts are shed in the urine. Clinical disease rarely results from infection, and treatment is generally not

Fig. 23-6  Ovarian cysts are identified in up to 75% of female guinea pigs and usually occur in both ovaries.

necessary,30 but sulfadimethoxine or trimethoprim-sulfa have been used. Segmental nephrosclerosis is reported as an incidental finding at necropsy and is often associated with autoimmune disease, high-protein diet, infections, and vascular diseases.41 Cytomegalovirus inclusion bodies have been identified in the kidneys as incidental findings at necropsy.41 Neoplasia of the urinary system is not common, but transitional cell carcinoma of the bladder, renal cell carcinoma, and renal fibrosarcoma have been identified in pet guinea pigs in the practice of one of the present authors (MGH).

REPRODUCTIVE DISEASES OVARIAN CYSTS Nonfunctional serous cysts (cystic rete ovarii) are extraordinarily common and have been identified in 66% to 75% of sows between 3 months and 5 years of age.8,46 Middle-aged (2- to 4-year-old) sows are most commonly affected. Serous cysts develop spontaneously throughout the estrous cycle. Follicular cysts also occurred in 22.4% of one study population and always coincided with serous cysts.46 They may be single or multilocular and are usually filled with clear fluid (Fig. 23-6). Cysts range in diameter from 0.5 to 7 cm and increase in size and prevalence as the animal ages.34 No significant correlation has been identified between reproductive history and the prevalence of cysts,34 but other problems reported concurrently with ovarian cysts include leiomyomas, granulosa cell tumors,8 cystic endometrial hyperplasia, and endometritis. In most cases, both ovaries are affected; however, if the cysts are unilateral, the right ovary is usually affected. The incidence of serous ovarian cysts increased following passive immunization against inhibin, suggesting that serous cysts are a normal component of the cyclic guinea pig ovary and that alterations in the inhibin-folliclestimulating hormone system may modulate the incidence of serous ovarian cysts in cavies.46 Affected animals present with abdominal distention and sometimes with anorexia, weakness, depression, and hunching in pain.4 If cysts are functional (follicular cysts), bilateral symmetric hair loss can be seen in the flank region. The most consistent sign of cysts in breeding sows is a decline in fertility after approximately 15 months of age. Diagnosis is best via ultrasound (Fig. 23-7), but abdominal radiography can identify

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SECTION III  Guinea Pigs and Chinchillas after 1 year of age may deliver normally, the majority of older primiparous sows have difficulty with parturition. Sows that are at risk must be monitored closely and may require cesarean section. Signs in full-term sows include contractions and straining that produce a bloody or green discharge but no pups. Pressure from the gravid uterus or pups may also lead to temporary paresis or paralysis of the rear legs. Palpation of the pelvic area for relaxation of the symphysis will help determine the need for a C-section. If the symphysis is separated approximately 2.5 to 3 cm, palpation of the vaginal canal may reveal a pup that must be manually removed or assisted. Sterile water-based lubricating gel placed in the vaginal canal helps ease pup removal. If the symphysis is relaxed and uterine inertia is suspected, treatment with oral or injectable calcium and/or oxytocin may facilitate contractions.

Fig. 23-7  Diagnosis of ovarian cysts is best made via ultra­ sonography.

larger cysts. The treatment of choice for ovarian cysts is ovariohysterectomy, but space-occupying cysts complicate the procedure. Postoperative care requires assisted feeding, fluids, and analgesics. If GI stasis secondary to pain is present, continue supportive care until the GI tract is functioning appropriately (see “Gastrointestinal Hypomotility or Stasis,” above). Palliative therapy with ultrasound-guided percutaneous drainage of the fluid can be performed, but cysts commonly return, sometimes within days. Gonadotropin-releasing hormone and human chorionic gonadotropin have been used to induce luteinization of ovarian follicular cysts43; but with nonfunctional serous cysts, temporary or no response to treatment is seen.

UTERINE PROLAPSE Uterine prolapse is most commonly associated with parturition.31 Acutely, the prolapse may be pink or red and smooth; however, depending on the duration of the prolapse, the tissues may be dry, darker, and covered with bedding or fecal material. The sow should be stabilized with fluids, analgesics, and antibiotics. The prolapsed tissue is rinsed thoroughly with sterile saline or dilute chlorhexidine solution. Then liberal amounts of a sterile lubricating gel are placed on the tissues to keep them moist until they have been reduced. If the prolapse is acute and the tissues are in good condition, cold hypertonic saline or sugar solutions are used to reduce swelling; then manual reduction of tissues can be performed. If prolapse occurs after reinsertion, ovariohysterectomy should be performed. Future breeding of the sow is not recommended. If the tissues are in poor condition, ovariohysterectomy is recommended after stabilization.

DYSTOCIA Guinea pigs are more predisposed to dystocia than other rodents or rabbits. This may be because of the large size of their pups, narrow pelvic canals, or fusion of the pubic symphysis.6 Other causes suggested are uterine torsion, obesity, nutritional (including vitamin C) deficiencies, and uterine inertia.5,16 Most cases occur in sows first bred after 8 to 12 months of age. Many breeders do not believe that the “fusing” of the pelvic symphysis always occurs, instead blaming dystocia of mature sows on obesity and vitamin C deficiency. Although some sows first bred

TOXEMIA OF PREGNANCY Toxemia of pregnancy is most commonly seen in pregnant sows 2 weeks prepartum to 2 weeks postpartum. There are probably two forms of pregnancy toxemia that may affect the guinea pig. The more familiar form, pregnancy ketosis, is similar to that seen in sheep and is caused by a negative energy balance of the sow due to the heavy demand of the growing fetuses. Predisposing factors include obesity, lack of exercise, large fetal loads, change in diet and/or environment, heat stress, and primiparity.6,18 The second form may be similar to that seen in pregnant women, which is caused by the gravid uterus compressing either its own vascular supply or that of the kidneys or GI tract, leading to tissue ischemia, and hypertension. In some cases, this condition may initiate disseminated intravascular coagulation (DIC).6,18 Hemorrhagic syndrome, which occurs during or shortly after parturition and usually results in fatal hemorrhage in the sow, may be related to this form of toxemia.6 Proposed etiologies for this syndrome in sows include vitamin K deficiency due to poorquality feed, hepatic dysfunction secondary to pressure from the gravid uterus, calcium deficiency affecting the clotting cascade, and other causes of DIC.6,18 Signs of pregnancy ketosis include complete or partial anorexia, lethargy, depression, uncoordinated movements, and dyspnea; this may progress to muscle spasms, paralysis, and death. Some sows die acutely and others deteriorate progressively over several days. The smell of ketones on the breath may occur with ketonemia. Laboratory findings include ketonuria, proteinuria, aciduria, hypoglycemia, acidosis, hyperlipemia, and hyperkalemia.6,18 Proteinuria can occur as a result of proximal tubular necrosis and subcapsular renal hemorrhage.41 Ultrasound imaging of the liver and necropsy findings often identify hepatic lipidosis. Treat ketosis with intravenous (IV) or intraosseous (IO) isotonic fluids with dextrose and oral glucose; calcium gluconate and magnesium sulfate have been suggested to be useful. Immediate emergency treatment of 1 to 2 mL of 50% dextrose administered in 3 to 5 mL saline IV or IO has been recommended. Nutrition is a critical component of treatment and is accomplished by syringe or gavage feeding via gastric, nasogastric, or esophagostomy tube. Herbivore critical care formulas should be used for maintenance feedings. Pregnancy-related blood pressure and ischemia issues associated with the heavily gravid uterus are significantly different from the ketosis type of pregnancy toxemia. Indirect blood pressure measurement will help determine whether the patient is

CHAPTER 23  Disease Problems of Guinea Pigs hypertensive (e.g., compression of the renal vessels) or hypotensive (e.g., shock). If the patient is hypertensive, an emergency cesarean section is required to relieve vascular compression. If the patient is hypotensive, treatment should commence with IV or IO crystalloid or colloid fluids, but it is likely that a cesarean section will be required as well. Treating pregnancy toxemia is often unsuccessful; therefore prevention is essential. Avoiding stress, obesity, and changes in the diet or environment during late pregnancy reduces the potential for pregnancy toxemia. Increase carbohydrate supplementation during the last 2 weeks of gestation and early postpartum period and make sure that food and water are readily available. Encourage exercise to minimize obesity before breeding, as the fetal load may exceed the sow’s weight and she may not be able to exercise sufficiently while pregnant. Postpartum breedings should be avoided as they are more likely to lead to large litters and thus larger fetal loads.

MASTITIS Bacteria cultured from infected mammary glands include E. coli, Pasteurella species, Klebsiella species, Staphylococcus species, Streptococcus species, and Pseudomonas species.6,38 Husbandry-related causes include dirty, wet cages, sharp objects, abrasive bedding, wire cage bottoms, and trauma from pups.6,18,38 The gland may become infected secondary to neoplasia. In acute mastitis the mammary glands are swollen, inflamed, reddened, and warm. The animal is often in pain, reluctant to move or eat, and will not nurse.6,38 In more chronic cases the glands become cool and cyanotic.18,38 There may or may not be a mucopurulent or bloody discharge from the teat or in the milk,6 and some may become abscessed and rupture. Diagnosis is based on clinical signs, culture and sensitivity, and cytology of the discharge or milk. Treat mastitis with antibiotics based on culture and sensitivity results, antiinflammatory medications, analgesics, hot-packing of the glands, and supportive care.6,38 Unresponsive cases may require surgical draining or removal of abscessed glands with histopathology to evaluate for neoplasia.18,38

PYOMETRA AND METRITIS Bacterial infections of the uterus may manifest as metritis or pyometra and are identified in breeding and nonbreeding animals.6 The most commonly isolated bacteria are B. bronchiseptica and hemolytic Streptococcus species, but other possible pathogens include E. coli, Corynebacterium pyogenes, and Staphylococcus species.6,18 Clinical signs include bloody or purulent vaginal discharge, abdominal distention, depression, anorexia, and fever.6 In more chronic cases, sows may be polydipsic and hypothermic. Diagnosis is made based on clinical signs, imaging, and culture and sensitivity results from the discharge. While survey radiographs can show uterine enlargement, ultrasound is the best modality for the diagnosis of this disease. Treatment involves stabilization with supportive care (e.g., IV, IO, SC fluids and nutrition), analgesics and/or NSAIDs, and broad-spectrum antibiotics while awaiting culture and sensitivity results.6,38 Ovariohysterectomy is the treatment of choice in nonbreeding animals.38 Long-term antibiotic therapy may be used in breeding sows, but the authors recommend ovariohysterectomy as the best treatment in all cases of uterine infection.

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VAGINITIS AND “SCROTAL PLUGS” Wet, soiled bedding combined with inguinal sebaceous secretions can become adhered to the penis, scrotum, vulva, or vagina, resulting in secondary infections or urinary/fecal obstruction. Older, intact boars may accumulate a scrotal plug of bedding, feces, or sebaceous material in the skin fold of the bilateral perineal sacs. Gently soak the affected part in a dilute chlorhexidine solution or in warm saline and carefully remove the debris; administer topical or systemic antibiotics as warranted. These conditions are easily prevented by appropriate sanitation and husbandry practices.

ORCHITIS AND EPIDIDYMITIS Infections of the male reproductive tract occur in breeding and nonbreeding boars via sexual transmission, bite wounds, and hematogenous spread.6 The most common pathogens isolated are B. bronchiseptica and Streptococcus species, but other bacterial causes occur, therefore culture any discharge identified.6 Signs include unilateral or bilateral swelling of the scrotal area, preputial discharge, anorexia, weight loss, and fever.6 Animals that recover may become carriers and therefore should not be used for breeding. Treatment involves appropriate antibiotics, supportive care, and analgesics and NSAIDs as indicated. In cases that do not resolve, castration is recommended.

NEOPLASIA There are many reports of reproductive tract tumors in the guinea pig. Leiomyomas and leiomyosarcomas of the uterus, ­ovarian adenocarcinomas, fibroadenomas, and adenocarcinomas of the mammary glands have all been reported.6,18 Mammary gland tumors are common in boars as well as sows. Approximately 50% are malignant tumors, but metastasis is not common. Hormonal abnormalities and viral etiologies have been proposed. Differentiate this condition from mastitis by cytology and/or biopsy. Clinical signs of mammary tumors include swelling of one or both glands with or without serous or bloody discharge. Ovarian and uterine neoplasms can be induced by exposure to estradiol, diethylstilbestrol, or testosterone.42,47 Clinical signs of uterine and ovarian tumors may be mistaken for pregnancy and include a hemorrhagic vaginal discharge, abdominal distention, and abdominal pain.6 One enlarged testicle with or without testicular atrophy in the opposite gonad is the most common clinical sign of a testicular tumor. The treatment of choice is surgical removal with histopathologic evaluation.6 Prevention of all reproductive neoplasms is best accomplished by ovariohysterectomy or castration of young animals.

DERMATOLOGIC DISEASES To evaluate dermatologic disorders in guinea pigs, it is important to recognize what is normal for a specific breed or for cavies in general. There are 13 recognized cavy breeds in the United States, and many mixed breeds are found in the pet trade. All guinea pigs have minimal or no hair between the nose and lips, around the lips, on the outer ear pinnae, and behind the ears. The Abyssinian and Abyssinian Satin breeds have many rosettes (central hairless areas with hair radiating in a circle) that should not be confused with alopecia or skin disease. The Teddy, Teddy Satin, and Texel breeds have a terrier-like coarse, kinky coat with

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SECTION III  Guinea Pigs and Chinchillas

curly whiskers. Occasionally hair and/or eyelashes may curl into the eyes of newborn babies, causing irritation. The Teddy and Texel also seem predisposed to dry, flaky skin. The coronet and white crested breeds have one rosette on top of the head. Peruvian and Peruvian Satin breeds normally have one rosette on the forehead (creating a frontal flow of hair) and one rosette over each hip (causing hair to be directed from the rear toward the head).

DERMATOPHYTOSIS Guinea pigs, like most mammals, are susceptible to fungal infection caused by dermatophytes. Young and immunosuppressed animals are more susceptible.38 Purebred Teddy and Teddy Satin guinea pigs appear to be the most susceptible to fungal infections. Dermatophytosis presents with scaly, patchy lesions on the face, feet, and dorsum.14 These skin lesions are usually circular areas of alopecia with inflamed and sometimes crusty edges; they are also pruritic.14,38 Although both Trichophyton mentagrophytes and Microsporum canis have been isolated, the majority of clinical cases are caused by T. mentagrophytes. Diagnosis is based on clinical signs, ultraviolet fluorescent lamp evaluation (Wood’s lamp), cytology, and fungal culture. T. mentagrophytes does not fluoresce with Wood’s lamp evaluation; thus this test is often negative.14,38 Fungal culture is required for accurate diagnosis. Treatment includes oral antifungal medications (itraconazole, 5 mg/kg q24h × 4-6 weeks; griseofulvin, 25 mg/kg q24h × 14-28 days; fluconazole, 16 mg/kg q24h × 14 days), antifungal shampoos (ketoconazole/chlorhexiderm), and topical antifungal lotions or sprays (miconazole, enilconazole, butenafine).14,38 Dermatophytosis is potentially zoonotic and the organisms may survive in the environment, leading to reinfection.38

ECTOPARASITES The most common ectoparasites found in guinea pigs include mites (Trixacarus caviae, Chirodiscoides caviae), lice (Gliricola porcelli, Gyropus ovalis), and rarely Demodex caviae.14,18,25,38 The most severe ectoparasitic dermatitis is caused by T. caviae. Infection can be caused by direct or indirect contact. Severe pruritus is a hallmark sign of infection; animals can scratch so intensely that they appear to be having a seizure; this may lead to severe self-induced trauma with secondary fungal or bacterial infections.14,18,38 Lesions include white-yellow crusty areas with inflammation and abrasions from self-induced trauma (Fig. 23-8).14,38 Louse infestation usually causes a less severe dermatitis and, because lice spend their entire life cycle on the host, requires direct contact for transmission. In heavy infestations, guinea pigs may have alopecia, crusty lesions, and an unthrifty-looking the coat. Flea (Ctenocephalides felis) infestations are also reported in guinea pigs.38 Cavies housed with rabbits, other rodents, or birds may become transiently infected with other ectoparasites (e.g., Sarcoptes muris, Notoedres muris, Mycoptes musculinus).25 Diagnose the condition by direct visualization of some ectoparasites such as lice and fleas, microscopic examination of skin scrapings, acetate tape preparations, and trichograms for specific organisms or parasite eggs.14,18,38 A microspatula with a flat-ended blade is preferable for skin scrapings, but the dull edge of a scalpel blade can also be used. Perform bacterial and fungal cultures of skin if secondary infections are suspected.

Fig. 23-8  Dermatitis (sarcoptic mange) caused by Trixacarus caviae. Severe infestation is seen, with crusty lesions covering the body.

Treatment is based upon the organism identified, but all products available are used off label in the guinea pig. Ivermectin (0.2-0.5 mg/kg PO, SC q7-10d × 3-4 treatments) has been the treatment of choice for T. caviae, but anecdotally topical selamectin treatment (6 mg/kg q14d × 3 treatments) has been successful. Treatment for lice and fleas may be accomplished with flea shampoos and powders that are safe for kittens. Ivermectin (0.2-0.5 mg/kg PO, SC q7-10d × 2 treatments) has been successful against lice. Use of monthly dog and cat topical flea products in guinea pigs is common for fleas and lice, but controversy exists as to the efficacy of some, and there are no data regarding the safety of any of these products in this species. Treat severe pruritus with antihistamines (diphenhydramine, hydroxyzine) and/or NSAIDs.14 Treat animals with secondary bacterial or fungal infections based upon culture and sensitivity. Asymptomatic animals should also be treated for ectoparasites and the environment disinfected.14 Lice are usually species-specific.25 T. caviae can be zoonotic and survive in the environment, leading to reinfection.38

CERVICAL LYMPHADENITIS Cervical lymphadenitis is usually caused by Streptococcus zooepidemicus and occasionally by Streptobacillus moniliformis, which is potentially zoonotic.14,25,38 S. zooepidemicus is considered part of the normal oropharyngeal/nasal flora of guinea pigs. Oral abrasions caused by overgrown teeth, abrasive feed, or bite wounds lead to invasion of the bacteria into deeper tissues and cervical lymph nodes, which become abscessed.14,25,38 Guinea pigs are presented with swellings in the neck region that are purulent and occasionally rupture on their own. In rare cases S. zooepidemicus can spread systemically and cause pneumonia, metritis, and septicemia.38 One of the present authors (CB) has observed an outbreak of S. zooepidemicus in a herd of approximately 500 guinea pigs. Animals were identified with typical signs of cervical lymphadenitis but also with acute hind-end paralysis, enlarged lymph nodes in the caudal region, metritis, abortion, dermatitis, and pneumonia. In all animals tested, the organism recovered was S. zooepidemicus. Diagnosis is based upon clinical signs, Gram’s stain, culture and sensitivity results from purulent material or lymph node tissue, and biopsy results of tissues.14,25,38 Treatment may require surgical removal or lancing of the abscessed lymph nodes and systemic antibiotics based on culture and sensitivity results.

CHAPTER 23  Disease Problems of Guinea Pigs

Fig. 23-9  Pododermatitis in the guinea pig. Swollen soft tissues and ulceration, as seen here, should alert the clinician to potential underlying osteomyelitis; radiographs are warranted.

PODODERMATITIS Pododermatitis is commonly seen in pet and laboratory cavies. Lesions may originally develop from irritation due to improper husbandry (e.g., wire bottom cages, abrasive or soiled bedding, sharp pieces of wood chips) and secondary bacterial invasion most often involves Staphylococcus aureus.25,38 The condition is most common in obese adult guinea pigs and results in a significant amount of pain and disability. Affected animals are reluctant to move around or eat and frequently are more vocal than normal.38 Vitamin C deficiency may be an important predisposing factor in animals with pododermatitis and supplementation is indicated as part of treatment protocols.14,25 Clinical signs vary from mild-to-severe inflammation, erythematous lesions with or without ulceration of primarily the plantar surfaces of the feet (Fig. 23-9), to granulomatous, callous-like swellings that, in severe cases, may result in bacterial invasion into the tendons, joints, and bone (e.g., osteomyelitis).14,25,38 Chronic inflammation may lead to amyloidosis of the kidney, liver, spleen, adrenal glands, and pancreas.25 Radiographs are indicated to diagnose osteomyelitis.38 In less severe cases, soak the lesions with dilute chlorhexidine or iodine solutions, maintain the animals on soft substrates, bandage, and administer systemic antibiotics based upon culture and sensitivity results. More severe cases may require long-term systemic antibiotics, surgical debridement ± antibiotic-impregnated polymethylmethacralate beads, bandaging with regular changes, and, as a salvage procedure, limb amputation.14,25,38 Analgesic and anti-inflammatory medications are an important part of treatment.14,25,38 The prognosis is poor in severe cases and prevention is essential. Proper diet, husbandry, cleanliness, and prevention of obesity are all helpful in the prevention of pododermatitis.

ALOPECIA Alopecia without inflammation is often related either to husbandry or hormonal conditions.14,25 Nutritional deficiencies as well as poor sanitation and bedding material reactions can cause alopecia in the guinea pig. Hormonal causes of alopecia include follicular ovarian cysts (bilaterally symmetric, nonpruritic; see

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Fig. 23-10  One of the most common masses seen in guinea pigs, the trichofolliculoma (arrow).

“Reproductive Diseases,” above) and hormonal changes common in late pregnancy or in lactating sows in poor condition or with large litters.14,38 The hairs are not completely epilated with barbering; close examination of the skin reveals broken hair shafts. Self-inflicted barbering may occur out of boredom or poor nutrition. The addition of hay and toys to the environment may resolve the problem. Dominant guinea pigs will barber their cage mates and may have to be separated. Examination of the barbering pattern will reveal whether it is self-inflicted (in which case the head and neck are spared).14,38

NEOPLASIA Trichofolliculomas are the most common tumors of guinea pigs. They are benign skin tumors seen predominantly in males and often arise on the dorsal rump, incorporating the scent glands (Fig. 23-10). They can be large, malodorous, exudative, and ulcerated, frequently with secondary infections and/or myiasis. Complete excision is usually curative. They are also considered incidental in most cases. Liposarcoma has a relatively high prevalence and is generally found in the skin or subcutis. Lymphoma is perhaps the most common malignancy, usually multicentric, and may appear in skin and viscera. Typically, lymphoma in cavies is a high-grade malignancy with a poor prognosis. Fibropapillomas of the ear canal are occasionally found in cavies. The etiology is undetermined; they are benign and usually resolve spontaneously.

MUSCULOSKELETAL DISEASES VITAMIN C DEFICIENCY (SCURVY) Guinea pigs are incapable of endogenous synthesis of vitamin C because they possess a mutated gene for l-gulono-g-lactone oxidase, which prevents the conversion of l-gulonolactone to l-ascorbic acid.35 Lack of dietary vitamin C results in defective type IV collagen, laminin, and elastin; this compromises blood vessel integrity and results in joint and gingival hemorrhages.29 Collagen is necessary to anchor teeth tightly; without it, teeth loosen and malocclusion occurs. In addition, vitamin C is necessary for appropriate retention of vitamin E.27 Young, growing animals are more susceptible to scurvy, and clinical disease can

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A

B Fig. 23-11  Postmortem findings of vitamin C deficiency in guinea pigs include periarticular hemorrhage (arrow), swollen joints (especially the stifle) (A), and enlargement of the costochondral junctions of the ribs (B) (arrows).

develop after as little as 2 weeks of ascorbic acid deprivation.10 Guinea pigs require 10 to 25 mg/kg per day of vitamin C added to their diet; pregnant animals require 30 mg/kg per day.17 Signs of vitamin C deficiency include rough hair coat, anorexia or difficulty prehending food, diarrhea, teeth grinding, vocalizing from pain, delayed wound healing, lameness, swollen joints (especially the stifle; Fig. 23-11A), GI stasis, and increased susceptibility to bacterial infections.30 Radiographically, long bone epiphyses and costochondral junctions of the ribs are enlarged (Fig. 23-11B). Pathologic fractures may also be evident. Postmortem examination may reveal hemorrhage into joints, skeletal muscle, gingiva, intestine, and subcutaneous tissues from abnormal collagen production. The diagnosis of vitamin C deficiency is based on history, clinical signs, and radiographic and pathologic lesions. Serum ascorbic acid levels can be used to confirm the diagnosis. Treat vitamin C deficiency with parenteral ascorbic acid at a dose of 50 to 100 mg/day IM or SC, but use caution with intramuscular medication because of the musculoskeletal pain associated with this disease. Once response is noted, administer vitamin C orally at the same dose. After recovery, ensure adequate supplementation of vitamin C daily in the diet. Fresh, good-quality guinea pig pellets provide adequate vitamin C if used within 90 days of the production date, but dietary recommendations have recently been made to reduce the percentage pellets in the diet because of possible urolithiasis.19 Fresh cabbage, kale, and oranges provide good sources of vitamin C. Vitamin C tablets (50 mg) are now available and can be freshly crushed and sprinkled over vegetables. Vitamin C added to the drinking water at a concentration of 200 to 400 mg/L daily has been advocated; but the vitamin is unstable in light and is most likely inactivated quickly.

OSTEOARTHRITIS AND OSTEOARTHROSIS Spontaneous osteoarthritis is clinically seen in guinea pigs but can also be secondary to ulcerative pododermatitis (see “Dermatologic Diseases,” above). Obesity and improper exercise and/ or substrate are predisposing factors. Treatment is palliative: soft clean bedding, pain management, increased exercise and/ or physical therapy of the limbs to increase range of motion, and prevention of obesity. Spontaneous cartilage degeneration and osteoarthrosis of the femorotibial joints of young guinea pigs have been described but no cause was identified.3

FIBROUS OSTEODYSTROPHY While there is only one report in the literature to date,44 anecdotal reports suggest that fibrous osteodystrophy is being documented in pet guinea pigs with increasing frequency. This condition is caused by primary or secondary hyperparathyroidism and results in increased osteoclastic resorption of bone and replacement by fibrous tissue. An imbalanced Ca:P ratio was found in the guinea pigs from the one report,44 and we have also seen guinea pigs presented with this disease that were purposely placed on low-calcium diets to prevent urolithiasis. Breeders in Europe and the United States also report this disease in the Satin type, suggesting an inherited factor in this line of guinea pigs. Diets with a low Ca:P ratio cause reduced growth rates, stiffness of joints, and calcification of soft tissues; they may also affect the uptake of magnesium and potassium.36 In addition, vitamin D deficiency, calcium malabsorption, or other calcium metabolism disturbances may play a role in the onset of disease. Clinical signs include anorexia or difficulty eating, lethargy, difficulty walking or unwillingness to move, or pathologic fractures. Diagnosis is based upon dietary history, physical examination findings, plasma total and ionized calcium concentrations, serum 25-hydroxyvitamin D, and parathyroid hormone concentrations. This disease must be differentiated from hypovitaminosis C, as the clinical signs can appear similar. Radiographs often show extensive changes to all bones in the body and skull, including osteopenia and pathologic fractures.44 Necropsy findings include severely thinned trabecular bone, marked osteoclastic activity, resorption of cortical bone and extensive replacement with fibrous connective tissue, and hyperplastic parathyroid glands (M. G. Hawkins, personal observation). No renal lesions have been identified to date. Treatment is aimed at normalizing Ca:P ratios via the diet. Initially treat with calcium gluconate intramuscularly, then continue with oral calcium glubionate therapy, in some cases this will be required for months before resolution is seen. Supplement vitamin D and increase exposure to sunlight empirically. Osteoporosis has also been reported in guinea pigs, so vitamin D supplementation should be used with caution.23 Pain management should include opioid medications for skeletal pain as well as anti-­inflammatory medications. Provide supportive care, including assisted

CHAPTER 23  Disease Problems of Guinea Pigs feeding and fluids as needed. Prognosis is guarded without aggressive supportive care.

METASTATIC MINERALIZATION Metastatic mineralization occurs in guinea pigs generally above 1 year of age. Often disease is subclinical, but clinical signs include unthriftiness, muscle stiffness, and renal dysfunction. The etiology is unclear and is possibly related to subclinical dietary imbalances such as an imbalanced Ca:P ratio, low magnesium and potassium, oversupplementation of dietary vitamin D3 or minerals, or dehydration.22,23,36,41 Soft tissue mineralization, including mineralization of kidneys, heart, vessels, brain, and GI tract, may be identified on radiographs.41 Lesions are irreversible once evident; prevention is by providing an appropriately balanced diet.23

IATROGENIC MUSCLE NECROSIS Ketamine, diazepam, and fentanyl/droperidol combinations have been implicated in nerve damage, self-mutilation, and muscle necrosis at injection sites.38

NUTRITIONAL MUSCULAR DYSTROPHY Myopathies have been reported to be associated with vitamin E/ selenium deficiencies. Clinical signs include lethargy, hind-limb weakness, decreased reproduction in breeding sows and boars, and conjunctivitis.41 Creatine kinase (CK) may be elevated. Severely affected animals may die within 1 week of presentation. As in other animals with this disease, the muscles may be pale and streaked at necropsy. Animals may respond to supplementation with vitamin E.

NEUROLOGIC DISEASES OTITIS MEDIA/OTITIS INTERNA While less frequent than in rabbits, otitis media occurs in pet guinea pigs and can progress to otitis interna. B. bronchiseptica, S. zooepidemicus, and S. pneumoniae have been associated with otitis media.23 It is unclear whether oral flora can become opportunistic with dental disease and ascend via the eustachian tube in guinea pigs, but otitis media and dental disease are often found together and cultures often isolate oral bacteria. Also aural polyps may occur within the tympanic bullae, with secondary infection causing clinical signs. Clinical signs include head tilt, ataxia, circling, torticollis, and facial nerve paralysis with secondary ulcerative keratitis due to exposure. Take skull radiographs to evaluate the bullae (Fig. 23-12A,B) and dentition; computed tomography can also be useful to determine the extent of the disease (Fig. 23-12C). Treat with appropriate antibiotics, anti-inflammatories, analgesics, and dental treatments, but these measures are often not curative. If needed, flush the ears with warm sterile saline under deep sedation/analgesia to help break up caseous material. Surgery (bullous osteotomy) has not been reported for guinea pigs.

MITES Severe infestations with T. caviae can cause such severe pruritus that guinea pigs are often presented for seizing (see “Dermatologic Diseases,” above).

307

RABIES Although rodents are not natural reservoirs for rabies virus, rabies has nevertheless been reported in guinea pigs.12 Clinicians should always consider rabies in guinea pigs with neurologic signs and husbandry conditions that could allow exposure.

LYMPHOCYTIC CHORIOMENINGITIS VIRUS Lymphocytic choriomeningitis virus (LCMV) is an arenavirus that can cause meningitis and hind-limb paralysis in guinea pigs, although it is more commonly reported in mice, hamsters, and chinchillas. Lesions include lymphocytic infiltrates in the choroid plexus, ependyma, and meninges.30 The virus is transmitted through inhalation, ingestion, or direct contact with contaminated urine, saliva, and feces. Biting insects can transmit LCMV, and transplacental transmission also occurs. LCMV can be transmitted to humans. Signs of LCMV infection in humans include headache, vomiting, and fever; fatalities are rare.

OPHTHALMOLOGIC DISEASES CORNEAL ULCERATION Corneal ulceration is most commonly caused by trauma and is diagnosed by fluorescein staining. Disseminated T-cell lymphosarcoma has been reported in a guinea pig that was presented with a unilateral corneal opacity.49

“PEA EYE” Adult guinea pigs, especially purebred American shorthairs, develop a protrusion from the inferior conjunctival sac in one or both eyes, termed “pea eye” by fanciers (Fig. 23-13). ­Histopathology of these tissues has demonstrated lesions in the lacrimal or zygomatic glands.26 Some lesions cause ventral ectropion, lagophthalmos, and secondary axial corneal degeneration.26 However, this condition does not appear to be painful and usually resolves without treatment.

CONJUNCTIVITIS Conjunctivitis is very common in pet guinea pigs and is generally caused by infection or hypovitaminosis C (scurvy). Chlamydophila caviae causes guinea pig inclusion conjunctivitis (GPIC) in laboratory and pet guinea pigs.28,51 Juvenile animals are most commonly affected. Some infected animals remain asymptomatic, but clinical signs range from mild to severe keratoconjunctivitis with serous to purulent ocular discharge, conjunctival chemosis, follicular hypertrophy, and uveitis.28 Diagnose GPIC by identifying intracytoplasmic inclusions in conjunctival scrapings and by performing C. caviae PCR on conjunctival swabs or scrapings.28,30,51 In a recent study, 48 of 75 symptomatic and 11 of 48 asymptomatic adult guinea pigs were positive via PCR for C. caviae. Conjunctival samples submitted from one owner and his cat and rabbit were also positive by PCR for C. caviae, and C. caviae was identified in rabbits housed with guinea pigs in another study.39 While the route of transmission is still unclear for this disease, nonetheless C. caviae GPIC could pose zoonotic

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SECTION III  Guinea Pigs and Chinchillas

A

B

C Fig. 23-12  (A) Normal (arrows) and (B) abnormal (arrows) radiographic appearances of the tympanic bullae of the guinea pig and (C) computed tomography (CT) imaging of bilateral tympanic bullae disease in the guinea pig. While survey radiography can provide evidence of sclerosis of the bullae, CT can provide evidence for osteoproliferative or osteolytic changes (arrows) as well as changes to the ear canals.

potential. It is often self-limiting within 3 to 4 weeks, but the potential zoonoses warrant treatment consideration with a topical tetracycline ophthalmic ointment. Conjunctivitis is one of the many clinical signs seen with hypovitaminosis C. Neoplasia of the guinea pig eye is uncommon38,49; lymphosarcoma affecting the conjunctival lymphoid tissue is most commonly reported.38

MISCELLANEOUS DISEASES OTOTOXICITY Guinea pigs seem quite sensitive to antibiotics; they may develop antibiotic-induced ototoxicity, especially when the agent is applied topically.1,7,32,56 Cisplatin induces ototoxicity when administered at human clinical doses,45 making guinea pigs a common model for study of this condition.

DIABETES MELLITUS Spontaneous diabetes mellitus similar to adult-onset diabetes in humans has been described in laboratory colonies of guinea pigs. Shortened life span (≤5 years), bladder hypertrophy, and voiding dysfunction have been reported.2 A pet guinea pig was diagnosed with diabetes mellitus after the animal presented with cystitis and urination of small, frequent amounts; it responded to insulin therapy.53,54 Diabetes mellitus may be transient in the guinea pig, and insulin therapy is generally not necessary. A low-fat, high-fiber diet is most important in treatment and prevention.54

HEAT STRESS Guinea pigs are susceptible to heat stress. Those housed outdoors can develop heat stress in ambient temperatures as low as 75°F (24°C). Guinea pigs will salivate profusely in an attempt

CHAPTER 23  Disease Problems of Guinea Pigs

Fig. 23-13  Adult guinea pigs, especially purebred American shorthairs, develop a protrusion from the inferior conjunctival sac in one or both eyes, termed “pea eye” by fanciers.

to thermoregulate; they will exhibit shallow, rapid respirations, pale mucous membranes, and elevated rectal temperature, which may be followed by coma and death. Treatment is supportive and includes cool water baths and parenteral fluids. Prognosis is very guarded.

LYMPHOSARCOMA/CAVIAN LEUKEMIA Lymphosarcoma is the most common malignancy of guinea pigs. Typically, lymphosarcoma in guinea pigs is a highgrade malignancy with poor prognosis. Clinical signs include anorexia, lethargy, unkept coat, and peripheral lymphadenopathy. Hepatomegaly, splenomegaly, and mediastinal masses are occasionally identified. It is sometimes seen concurrently with lymphoblastic leukemia.10 Cavian leukemia caused by a type-C retrovirus was reported in the 1960s in laboratory guinea pigs; leukemic animals may have a total white blood cell count of 25,000 to 500,000/mL.10 Diagnosis is based on the results of a complete blood count and cytologic examination of aspirates of enlarged nodes or abdominal or pleural fluids. At necropsy, lymph nodes and visceral organs may be enlarged, with infiltration by proliferating lymphoblasts. The course of the disease can be short, as little as 2 to 5 weeks. The prognosis is generally poor, although some animals have responded initially to chemotherapy. To date, there are no published reports regarding the use of chemotherapeutics in guinea pigs.

References 1. Aquino TJ, Oliveira JA, Rossato M. Ototoxicity and otoprotection in the inner ear of guinea pigs using gentamicin and amikacin: ultrastructural and functional aspects. Braz J Otorhinolaryngol. 2008;74:843-852. 2. Belis JA, Curley RM, Lang CM. Bladder dysfunction in the spontaneously diabetic male Abyssinian-Hartley guinea pig. ­Pharmacology. 1996;53:66-70.

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3. Bendele A, McComb J, Gould T, et al. Animal models of arthritis: relevance to human disease. Toxicol Pathol. 1999;27:134-142. 4. Beregi A, Zorn S, Felkai F. Ultrasonic diagnosis of ovarian cysts in ten guinea pigs. Vet Radiol Ultrasound. 1999;40:74-76. 5. Bishop C. Emergency medicine and surgery of rabbits and rodents. In: Proceedings. Ontario Vet Med Assoc. 1999:115-124. 6. Bishop CR. Reproductive medicine of rabbits and rodents. Vet Clin North Am Exot Anim Pract. 2002;5:507-535. 7. Blakley BW, Hochman J, Wellman M, et  al. Differences in ototoxicity across species. J Otolaryngol Head Neck Surg. 2008; 37:700-703. 8. Burns RP, Paul-Murphy J, Sicard GK. Granulosa cell tumor in a guinea pig. J Am Vet Med Assoc. 2001;218:726-728. 9. Chrisp CE, Suckow MA, Fayer R, et  al. Comparison of the host ranges and antigenicity of Cryptosporidium parvum and Cryptosporidium wrairi from guinea pigs. J Protozool. 1992;39:406-409. 10. Collins B. Common diseases and medical management of rodents and lagomorphs. In: Jacobson E, Kollias G, eds. Exotic animals. New York: Churchill Livingstone; 1988:261-316. 11. De Voe R. Clinical snapshot: gastric bloat with possible volvulus in a guinea pig. Compend Contin Educ Pract Vet. 2001; 23:543:571. 12. Eidson M, Matthews SD, Willsey AL, et al. Rabies virus infection in a pet guinea pig and seven pet rabbits. J Am Vet Med Assoc. 2005;227:932-935. 13. Fehr M, Rappold S. Urolithiasis in 20 guinea pigs (Cavia porcellus). Tierarztl Prax. 1997;25:543-547. 14. Flecknell P. Guinea pigs. In: Meredith A, Redrobe S, eds. BSAVA manual of exotic pets. 4th ed. Quedgeley: BSAVA; 2002:52-64. 15. Gaschen L, Ketz C, Lang J, et al. Ultrasonographic detection of adrenal gland tumor and ureterolithiasis in a guinea pig. Vet Radiol Ultrasound. 1998;39:43-46. 16. Hardesty D. The effect of obesity on fertility in the cavy. In: The American Rabbit Breeders Association. Official guide book raising better rabbits and cavies. Bloomington, IL: ARBA; 2000: 196-200. 17. Harkness JE, Turner PV, Vande Woude S, et al. Biology and husbandry-guinea pig. In: Harkness and Wagner’s biology and medicine of rabbits and rodents. Ames: Wiley-Blackwell; 2010:45-57. 18. Harkness JE, Turner PV, Vande Woude S, et al. Specific diseases and conditions. In: Harkness and Wagner’s biology and medicine of rabbits and rodents. Ames: Wiley-Blackwell; 2010:249-396. 19. Hawkins MG, Drazenovich TL, Kass PH, et al. Risk factors associated with the development of urolithiasis in pet guinea pigs (Cavia porcellus). Proceedings. Annu Assoc Avian Vet Assoc Exot Mam Vet. 2008:59. 20. Hawkins MG, Ruby AL, Drazenovich TL, et  al. Composition and characteristics of urinary calculi from guinea pigs. J Am Vet Med Assoc. 2009;234:214-220. 21. Hawkins MG, Vernau W, Drazenovich TL, et al. Results of cytologic and microbiologic analysis of bronchoalveolar lavage fluid in New Zealand white rabbits. Am J Vet Res. 2008;69:572-578. 22. Hoefer H, Latney L. Rodents: urogenital and reproductive system disorders. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: BSAVA; 2009:150-160. 23. Hollamby S. Rodents: neurological and musculoskeletal disorders. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: BSAVA; 2009:161-168. 24. Holowaychuk MK. Renal failure in a guinea pig (Cavia porcellus) following ingestion of oxalate containing plants. Can Vet J. 2006;47:787-789. 25. Huerkamp MJ, Murray KA, Orosz SE. Guinea pigs. In: LaberLaird K, Swindle MM, Flecknell P, eds. Handbook of rodent and rabbit medicine. New York: Elsevier Science Inc; 1996:91-95. 26. Kern TJ. Rabbit and rodent ophthalmology. Sem in Avian Exot Pet Med. 1997;6:138-145.

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27. Liu JF, Lee YW. Vitamin C supplementation restores the impaired vitamin E status of guinea pigs fed oxidized frying oil. J Nutr. 1998;128:116-122. 28. Lutz-Wohlgroth L, Becker A, Brugnera E, et al. Chlamydiales in guinea-pigs and their zoonotic potential. J Vet Med A Physiol Pathol Clin Med. 2006;53:185-193. 29. Mahmoodian F, Peterkofsky B. Vitamin C deficiency in guinea pigs differentially affects the expression of type IV collagen, laminin, and elastin in blood vessels. J Nutr. 1999;129:83-91. 30. Manning P, Wagner JE, Harkness J. Biology and diseases of guinea pigs. In: Fox J, Cohen B, Loew F, eds. Laboratory animal medicine. Orlando: Academic Press; 1984:149-181. 31. Mehler SJ, Bennett RA. Surgical oncology of exotic animals. Vet Clin North Am Exot Anim Pract. 2004;7:783-805. 32. Migirov L, Himmelfarb M. Methodology for studying the effects of topically applied ear drops on otoacoustic emissions in guinea pigs. J Laryngol Otol. 2003;117:696-699. 33. Mitchell EM, MacLeod A, Hawkins MG. Gastric dilatation-­ volvulus in a guinea pig (Cavia porcellus). J Am Anim Hosp Assoc. 2010;46:174-180. 34. Nielsen TD, Holt S, Ruelokke ML, et al. Ovarian cysts in guinea pigs: influence of age and reproductive status on prevalence and size. J Small Anim Pract. 2003;44:257-260. 35. Nishikimi M, Kawai T, Yagi K. Guinea pigs possess a highly mutated gene for l-gulono-gamma-lactone oxidase, the key enzyme for l-ascorbic acid biosynthesis missing in this species. J Biol Chem. 1992;267:21967-21972. 36. O’Dell BL, Morris ER, Pickett EE, et al. Diet composition and mineral balance in guinea pigs. J Nutr. 1957;63:65-77. 37. Okewole PA, Odeyemi PS, Oladunmade MA, et al. An outbreak of Streptococcus pyogenes infection associated with calcium oxalate urolithiasis in guinea pigs (Cavia porcellus). Lab Anim. 1991;25:184-186. 38. O’Rourke DP. Disease problems of guinea pigs. In: ­Quesenberry KE, Carpenter JW, eds. Ferrets, rabbits, and rodents: clinical medicine and surgery. 2nd ed. St. Louis: WB Saunders; 2004:245-254. 39. Pantchev A, Sting R, Bauerfeind R, et al. Detection of all Chlamydophila and Chlamydia spp. of veterinary interest using species-specific real-time PCR assays. Comp Immunol Microbiol Infect Dis. 2010;33:473-484. 40. Peng X, Griffith JW, Lang CM. Cystitis, urolithiasis and cystic calculi in ageing guinea pigs. Lab Anim. 1990;24:159-163. 41. Percy DH, Barthold SW. Guinea pig. In: Percy DH, Barthold SW, eds. Pathology of laboratory rodents and rabbits. 2nd ed. Ames: Blackwell Publishing; 2001:209-247.

42. Porter KB, Tsibris JC, Porter GW, et al. Use of endoscopic and ultrasound techniques in the guinea pig leiomyoma model. Lab Anim Sci. 1997;47:537-539. 43. Purswell BJ, Parker NA, Bailey TL, et al. Theriogenology question of the month. Persistent estrus caused by functional granulosa cell tumor of the left ovary. J Am Vet Med Assoc. 1999;215:193-195. 44. Schwarz T, Stork CK, Megahy IW, et al. Osteodystrophia fibrosa in two guinea pigs. J Am Vet Med Assoc. 2001;219:63-66. 45. Sepmeijer JW, Klis SF. Distribution of platinum in blood and perilymph in relation to cisplatin induced ototoxicity in the guinea pig. Hear Res. 2009;247:34-39. 46. Shi F, Petroff BK, Herath CB, et  al. Serous cysts are a benign component of the cyclic ovary in the guinea pig with an incidence dependent upon inhibin bioactivity. J Vet Med Sci. 2002;64:129-135. 47. Silva EG, Tornos C, Deavers M, et  al. Induction of epithelial neoplasms in the ovaries of guinea pigs by estrogenic stimulation. Gynecol Oncol. 1998;71:240-246. 48. Spink RR. Urolithiasis in a guinea pig (Cavia porcellus). Vet Med Small Anim Clin. 1978;73:501-502. 49. Steinberg H. Disseminated T-cell lymphoma in a guinea pig with bilateral ocular involvement. J Vet Diagn Invest. 2000; 12:459-462. 50. Stieger SM, Wenker C, Ziegler-Gohm D, et  al. Ureterolithiasis and papilloma formation in the ureter of a guinea pig. Vet Radiol Ultrasound. 2003;44:326-329. 51. Strik NI, Alleman AR, Wellehan JFX. Conjunctival swab cytology from a guinea pig: it’s elementary! Vet Clin Pathol. 2005;34:169-171. 52. Stuppy DE, Douglass PR, Douglass PJ. Urolithiasis and cystotomy in a guinea pig (Cavia porcellanus). Vet Med Small Anim Clin. 1979;74:465-467. 53. Vannevel J. Diabetes mellitus in a 3-year-old, intact, female guinea pig. Can Vet J. 1998;39:503. 54. Vannevel J. Diabetes in the guinea pig–not uncommon. Can Vet J. 1999;40:613. 55. Xia Y, Hu HZ, Liu S, et  al. Clostridium difficile toxin A excites enteric neurons and suppresses sympathetic neurotransmission in the guinea pig. Gut. 2000;46:481-486. 56. Xu M, Hu HT, Jin Z, et al. Ototoxicity on cochlear nucleus neurons following systemic application of gentamicin. Acta Otolaryngol. 2009;129:745-748.

CHAPTER

24

Disease Problems of Chinchillas

 

Christoph Mans, MedVet, and Thomas M. Donnelly, BVSc, Diplomate ACLAM

Disorders of the Digestive System Gastroenteritis and Dysbacteriosis Constipation Diarrhea and Soft Feces Tympany Rectal Tissue Prolapse and Intussusception Esophageal Disorders Dental Disorders Eye Disorders Epiphora Conjunctivitis Corneal Disorders Other Eye Disorders Ear Disorders Respiratory System Disorders Reproductive System Disorders Endometritis and Pyometra Dystocia Penile Disorders Urinary System Disorders Neurologic Disorders Seizures Heat Stroke Lead Poisoning Dermatologic Disorders Dermatophytosis Fur Chewing Fur Slip Matted Fur Foot Disorders Miscellaneous Disease Problems Cardiac Disease Hepatic Lipidosis and Ketosis Diabetes Mellitus Fractures Neoplasia Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

Infectious Diseases Viral Infections Fungal Infections Parasitic Infections Bacterial Infections

DISORDERS OF THE DIGESTIVE SYSTEM GASTROENTERITIS AND DYSBACTERIOSIS In chinchillas, any systemic disease or a painful or stressful condition may result in secondary gastrointestinal problems with nonspecific clinical signs such as anorexia, lack of fecal output, and lethargy. Obtaining a thorough clinical history and physical examination is critical for formulating a sound diagnostic and therapeutic plan. A variety of infectious and noninfectious causes of gastroenteritis and dysbacteriosis may affect chinchillas and result in a range of clinical syndromes including constipation, tympany, diarrhea, intussusception, and rectal prolapse. Gastrointestinal disorders were the major cause of morbidity and mortality in farmed chinchillas in the 1960s to 1990s. In pet chinchillas, noninfectious causes such as sudden dietary changes or inappropriate oral antibiotic therapy (e.g., cephalosporins, penicillins, clindamycin, erythromycin) are more frequent and important. However, a few secondary infectious causes such as giardiasis, coccidiosis, and Pseudomonas or Enterobacter overgrowth can be seen in pet chinchillas. Identifying the underlying cause of gastroenteritis and dysbacteriosis is important to improve the therapeutic outcome and reduce the chance of recurrence. Consider performing whole-body radiographs, fecal parasite examination, fecal cytology, and fecal culture for enteric opportunistic pathogens (e.g., Escherichia coli, Pseudomonas aeruginosa) in the initial diagnostic workup. Laboratory tests such as urinalysis, plasma biochemical analysis, and a complete blood count (CBC) can aid in diagnosing non-alimentary and concurrent metabolic disorders (e.g., hepatic lipidosis, ketosis, renal disease) that will influence 311

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the prognosis and therapy. Besides specific treatment for the primary underlying cause of gastroenteritis and dysbacteriosis, general treatment guidelines for all cases of gastrointestinal disease should include replacing fluid deficits and maintaining normovolemia by parenteral and enteral fluid therapy, nutritional and caloric support, and managing pain (buprenorphine 0.03-0.05 mg/kg SC q8h) if a painful condition is suspected. Consider systemic parenteral antimicrobial therapy (enrofloxacin 10 mg/kg SC diluted q12h) (Baytril Animal Health, Shawnee Mission, KS) for the treatment of predominately gramnegative opportunistic pathogens in chinchillas with severe dysbacteriosis when an infectious cause is suspected but unconfirmed and the animal is in a compromised general condition. Avoid oral drug administration because the absorption and effectiveness of oral drugs may be decreased when gastrointestinal function is abnormal. Parenteral administration is the preferred initial route for most drugs. Once an animal is eating and gastrointestinal function is improved, switch to the oral route.

CONSTIPATION Constipation is a common problem in chinchillas.16,61 Reduced output or absence of fecal pellets that are smaller and irregular in size is commonly seen. Sudden changes in diet, an inappropriate diet, or infectious causes can lead to dysbacteriosis, gastroenteritis, and ileus and consequently constipation. Underlying causes include anorexia, dental disease, and gastroenteritis.16,61 Abdominal palpation may reveal firm ingesta in the cecum and a tense abdomen. Animals often become anorexic and progressively lethargic. An important differential diagnosis for complete lack of fecal output in chinchillas is intestinal intussusception. General guidelines for the treatment of gastrointestinal disease apply for constipation. The aim is to rehydrate the gut by fluid therapy. Consider enteral fluid therapy (100 mL/kg PO q24h divided in 4-5 doses) in chinchillas with constipation to stimulate a gastrocecal reflex and rehydrate dehydrated ingesta.50,77 Abdominal massage and regular exercise may also be helpful.

DIARRHEA AND SOFT FECES Diarrhea and soft feces are common presentations in chinchillas. Besides infectious causes (e.g., parasites, bacteria), inappropriate feeding or sudden changes in diet with resultant dysbacteriosis commonly cause soft feces. Overfeeding of fresh green feed or items high in simple carbohydrates can result in dysbacteriosis and soft feces. Feces smeared on the cage resting board of a pet with or without matted, fecal-stained perianal fur are often the first signs the owner notices. The animal might be otherwise normal or, in chronic and severe cases, it can be anorexic, dehydrated, and depressed. Rule out infectious causes based on the history and by appropriate diagnostic testing. Intestinal secondary yeast overgrowth, caused by Cyniclomyces guttulatus (previously Saccharomycopsis guttulata), which lines the stomach, is often seen in chinchillas with soft feces.145 However, increased numbers of this yeast in chinchillas with diarrhea or soft feces is considered secondary rather than a cause, usually promoted by an underlying disease process.38 Offer well-dried, high-quality grass hay if the animal is still eating. Consider treatment with nystatin (100,000 IU/kg PO q8h for 5 days) if C. guttulatus overgrowth is very high or no response to initial treatment is seen.

TYMPANY Tympany of the stomach and intestine is less common in chinchillas compared with other rodents or rabbits. Tympany is often secondary to gastroenteritis, dysbacteriosis, ileus or luminal obstruction or, very rarely, intestinal torsion.86 The affected animals usually has a distended and tense abdomen. In severe cases, depressed and dyspneic chinchillas may lie on their sides. Signs of shock may be present. Prognosis depends on the severity and duration of tympany but is usually poor in severe or chronic cases. Institute treatment based on general guidelines for the management of gastroenteritis. Although gastric decompression is recommended for severe cases of tympany, this might result in collapse and death in a decompensated patient. Do not use motility-enhancing drugs (e.g., cisapride) if an infectious or obstructive cause cannot be ruled out.

RECTAL TISSUE PROLAPSE AND INTUSSUSCEPTION Rectal tissue prolapse and intestinal intussusception frequently occur together, secondary to dysbacteriosis, enteritis, constipation, or diarrhea (Fig. 24-1).82,128 Intussusception of the descending colon and rectum is associated with most cases of rectal prolapse; however, the small intestine can also be affected.82,94,128 Abdominal palpation might reveal a turgid cylindrical mass reflecting the intussuscepted portion of the intestine.82 The amount of intestine involved in the intussusception can be extensive and the affected portion is usually cyanotic, congested, and, in advanced cases, often nonviable (see Fig. 24-1, B).82 Besides treatment of the primary underlying cause, surgically correcting the intussusception is critical: intestinal resection and anastomosis may be necessary. Simple replacement or resection of prolapsed rectal tissue is insufficient. Assess the prolapsed tissue for viability and trauma. If an intussusception is ruled out, carefully clean and soak the edematous prolapsed rectal tissue in a concentrated sugar solution (50% dextrose). Replace the prolapsed tissue and place a perianal purse-string suture.94,128 Successful outcome after laparotomy and manual correction of more proximally located intussusceptions has been reported.82 However, the prognosis remains poor in most cases and the outcome will depend on the location and duration of the intussusception, viability of the prolapsed and intussuscepted tissue, and the underlying primary cause. Recurrence and rapid deterioration of affected animals is unfortunately common, since rectal prolapse and intussusception usually reflect an acute complication of a more chronic underlying primary problem.

ESOPHAGEAL DISORDERS Because chinchillas cannot vomit, food items such as raisins, fruit, and nuts as well as bedding material and ingested placentas in postparturient females can become stuck in the oropharynx and upper esophagus.20,38 Clinical signs are a sudden onset of anorexia, drooling, retching, and possible dyspnea. Removal of the foreign material is usually curative and the prognosis is good if the problem is dealt with early.38 Megaesophagus was diagnosed in a 2-year-old chinchilla that presented for recurrent acute episodes of dyspnea and nasal discharge.63 During hospitalization, the animal regurgitated food material episodically, followed by dyspnea, retching, gasping, and nasal discharge.

CHAPTER 24  Disease Problems of Chinchillas

313

B

A

Fig. 24-1  A, Rectal prolapse and intussusception in a 1.5-year-old male chinchilla. The animal presented with a 24-hour history of reduced fecal output and appetite. A firm cylindrical mass, identified as a small intestinal intussusception on necropsy, was palpable in the midabdomen. B, Dissected small intestine of the chinchilla in (A) showing a 4.5-cm section of intussuscepted necrotic jejunum.

Fig. 24-2  Radiograph of a 2-year-old chinchilla 15-minutes after oral administration of barium. The chinchilla presented for recurrent acute episodes of dyspnea and nasal discharge. Megaesophagus was diagnosed based on the dilated and barium-filled thoracic esophagus. Part of the bolus is seen in the stomach. (From Jopp IP, Stengel C, Kraft W. Megaesophagus in a chinchilla (Chinchilla lanigera). A case report. [German] Megaosophagus bei einem Chinchilla (Chinchilla lanigera) Ein Fallbericht. Tierarztl Prax Ausg K Klientiere Heimtiere 2004;32:96-100.)

Upper gastrointestinal contrast radiography aided the diagnosis of megaesophagus in this animal (Fig. 24-2).

DENTAL DISORDERS Dental disease is commonly diagnosed in chinchillas and can affect animals of all ages (see Chapter 32).61 In two studies, abnormalities related to subclinical dental disease were detected in 35%21 and 33%61 of apparently healthy chinchillas presented for routine physical examination. Nutritional (e.g., less abrasive

diet in captivity) and genetic causes have been proposed as the predisposing factors for the development of dental disease in chinchillas.24 Tooth elongation and its secondary complications, affecting the reserve or the clinical crown or both, is the underlying cause for most clinical signs associated with dental disease. Despite sometimes dramatic elongation of reserve and clinical crowns of the cheek teeth, animals often have no difficulty eating and maintain a good body condition until severe changes and complications, such as soft tissue trauma from sharp dental spikes or periodontal infection, have occurred. Common clinical findings associated with dental disease in chinchillas are reduced food intake, changed food preferences toward more easily chewed feed items, weight loss, reduced fecal output with smaller, irregularly shaped fecal pellets, salivastained skin and fur with crusting and alopecia of the perioral area, wetting and crusting of the chin (“slobbers”) and forefeet, epiphora, poor fur condition, and fur chewing.21,61 On clinical examination, palpable irregularities of the ventral borders of the mandible, and overgrown or irregular occlusal surfaces of the incisor teeth may be found.21,61 Facial abscesses of periodontal origin are seen infrequently but can occur.22,61 A limited examination using a pediatric laryngoscope, otoscope, or vaginal speculum can be performed in a conscious animal, but up to 50% of intraoral lesions can be missed.21 Instead, perform a thorough examination of the oral cavity under general anesthesia. Endoscopic-guided intraoral examination (stomatoscopy) provides superior visibility and increases the chance for detection of pathologic lesions (see Chapter 32). Common intraoral findings involving the cheek teeth include coronal elongation, changes of the occlusal surface, formation of sharp spikes buccally and less commonly lingually on the edges of the occlusal surfaces, and widened interproximal coronal spaces that facilitate impaction and promote the development of periodontal disease.22,61 Resorptive and caries-like lesions of the cheek teeth are common in chinchillas; loss of tooth substance or brown discoloration of occlusal and interproximal tooth surfaces is seen.21-23,61 Erosions of the buccal mucosa, gingival hyperplasia, and gingival pocketing are common intraoral findings secondary to dental disease.21,61

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SECTION III  Guinea Pigs and Chinchillas

Evaluate radiographs or computed tomography (CT) scans of the skull in any chinchilla with a history or clinical signs suggestive of underlying dental disease. Some clinicians like to use anatomic reference lines for objective interpretation of skull radiographs and staging of dental disease in chinchillas.11 The prognosis for chinchillas diagnosed with dental disease depends on the severity of dental disease, the animal’s general condition, and owner compliance. Repeated intraoral examinations and treatments under general anesthesia at varying intervals, often for the life of the animal, are usually necessary to control complications of dental disease and maintain an acceptable quality of life for the animal. The goals of therapy should include reduction of periodontal infection and soft tissue trauma, which both lead to discomfort and pain, and either recovering or maintaining the animal’s ability to eat unaided. After removing spikes and reducing elongated crowns (see Chapter 32), probe interproximal spaces and gingival pockets and remove impacted debris. Rinse cleaned gingival and periodontal pockets carefully with 2% hydrogen peroxide or diluted chlorhexidine solution.115 Instilling Doxirobe gel (Pfizer Animal Health, New York, NY) in deep (>5 mm) gingival and periodontal pockets may help delay reimpaction with debris and reduce periodontal inflammation.144 If evidence of significant periodontal infection exists, begin systemic antimicrobial therapy with antimicrobials that are effective against anerobic bacteria predominating in periodontal infections (e.g., penicillin G benzathine 50,000 IU/kg SC q5d).133 Limit extraction of cheek teeth to those that are severely diseased and mobile. Removal of cheek teeth may lead to an increased rate of coronal overgrowth of the opposing cheek teeth, owing to the lack of attrition, and can necessitate more frequent coronal crown reductions. Managing pain (e.g., buprenorphine 0.03-0.05 mg/kg SC q8h, meloxicam 0.3-0.5 mg/kg PO, SC q12-24h) and coexisting conditions such as hepatic lipidosis, ketosis, and constipation is critical. With animals in advanced stages of dental disease, offer easy-to-chew food items (soft leafy grass hay, moistened pellets, and “critical-care” formulas offered on a dish). Many animals will require short- and sometimes long-term nutritional support after a dental procedure.

EYE DISORDERS EPIPHORA Epiphora (“wet eye”) is a common condition in chinchillas, usually characterized by unilateral serous discharge, wetting of the periocular fur, and potentially periocular alopecia and dermatitis. Because of secondary bone remodeling, a common underlying cause for epiphora is elongated apical reserve crowns of both the maxillary premolar and first two molar teeth, causing complete or partial compression and consequent obstruction of the nasolacrimal duct.22,25 In contrast to rabbits, obliteration of the nasolacrimal duct at the level of the apical portion of the maxillary incisor is uncommon in chinchillas.16,25,61 Because of the very small size of the lacrimal punctae, visualizing and catheterizing the lacrimal duct for flushing is very difficult and not routinely performed.22 Treat concurrent or underlying infection and inflammation with appropriate topical antibiotics and nonsteroidal anti-inflammatory drugs (see Chapter 37). If underlying dental disease is responsible for the nasolacrimal duct obstruction, it is unlikely that permanent patency of

the nasolacrimal duct will be regained, as there is no effective treatment for apical reserve crown elongation.

CONJUNCTIVITIS Conjunctivitis is common in chinchillas. Irritation from excessive sand bathing, inadequate cage ventilation, or underlying nasolacrimal duct obstruction is often the cause. A variety of bacteria can cause primary bacterial conjunctivitis. Conjunctivitis caused by P. aeruginosa as a localized infection or as part of a systemic infection has been reported.28,138 Depression and anorexia, commonly seen in cases of pseudomonal conjunctivitis, indicates possible underlying systemic infection.28,138 There is controversy as to whether P. aeruginosa is part of the normal conjunctival flora, which consists predominately of gram-­ positive organisms including Staphylococcus, Streptococcus, and Corynebacterium species.75,85,91 Fluorescein staining is necessary to rule out damage to the corneal surface. Submit a conjunctival swab for bacterial culture and antibiotic susceptibility testing. Treatment includes thoroughly lavaging the conjunctival sac with physiologic saline and applying a topical broad-spectrum antibiotic (e.g., gentamicin) that provides coverage against gram-negative and multiresistant bacteria. Access to a sand bath should be restricted until the chinchilla is fully recovered. Early recognition, systemic antibiotic therapy, and supportive care are critical for chinchillas suffering from systemic Pseudomonas infection.28

CORNEAL DISORDERS Corneal damage and keratitis secondary to trauma are common clinical findings in chinchillas, usually associated with blepharospasm, discharge, and conjunctivitis.35 The large, prominent corneal surface possibly predisposes chinchillas to corneal trauma. Excessive sand bathing, the provision of inappropriate sand for bathing, and inappropriate housing conditions should be considered as possible underlying causes.35,135 Diagnosis is by fluorescein staining of the corneal surface. Avoid direct contact between fluorescein-impregnated test strips and the corneal surface because this can lead to false-positive results.35 Rule out possible nontraumatic underlying causes (e.g., exophthalmos, trichiasis) to avoid reoccurrence. Treatment of acute superficial lesions includes application of antibiotic ophthalmic formulations. In cases of chronic nonhealing ulcers, consider corneal debridement or grid keratomy after controlling any potential bacterial infection.

OTHER EYE DISORDERS Exophthalmos can be caused by a retrobulbar process or increased intraocular pressure. The chinchilla’s orbit is shallow, and manual retraction of the eyelids can cause iatrogenic proptosis of the globe.135 Retrobulbar periapical abscesses of the maxillary cheek teeth—causing exophthalmos—are less common in chinchillas than in guinea pigs or rabbits.21 A rare retrobulbar taenia cyst has been described.56 Primary glaucoma has not been reported in chinchillas. Secondary glaucoma is uncommon but has been reported, and reference intervals for intraocular pressure in chinchillas have been published.85,111 Uveitis and panophthalmitis are uncommon diagnoses in chinchillas and may be caused by trauma or a systemic inflammatory process.135 A rare case of human herpesvirus I infection

CHAPTER 24  Disease Problems of Chinchillas in a chinchilla, causing ulcerative keratitis, retinitis, neuritis and meningoencephalitis, has been reported.140 Lenticular changes such as cataracts, nucleosclerosis, and lens sutures can occur in chinchillas.37,81,85,111 Because diabetogenic cataracts have been reported in chinchillas, diabetes mellitus should be ruled out in animals presenting with uni- or bilateral cataracts.37 Asteroid hyalosis of the vitreous humor, in which lipid-­ calcium particles are formed as part of a degenerative process, can occur in older chinchillas.111

EAR DISORDERS Chinchillas are used as animal models for human otologic disease research, including hearing loss and otitis media.3,5 Multiple studies on the pathophysiology and antimicrobial treatment of experimentally induced otitis media in chinchillas are available.1,2,4,19,60 Clinical signs caused by otitis externa, media, and interna can range from external ear canal discharge, head shaking, and mild head tilt to facial paresis and severe neurologic deficits.138 Otitis externa is often the result of a perforated tympanic membrane secondary to otitis media; therefore discharge from the ear canal should prompt the clinician to rule out a middle ear infection. Skull radiographs in dorsoventral projection or, preferably, computed tomography (CT) scans are used for the diagnosis of otitis media (Fig. 24-3). Otoscopic examination of the external ear canal and tympanic membrane may reveal purulent exudate and inflammation of the external ear canal. A variety of bacteria may be cultured from the middle ear in chinchillas diagnosed with otitis media. Clinical signs associated with an epizootic outbreak of P. aeruginosa-induced otitis media and interna in a breeding facility were ear discharge, conjunctivitis, neurologic signs, and sudden death due to septicemia.138 Treatment of

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bacterial otitis media and interna remains challenging because of the formation of biofilms in the middle ear that can reduce the efficacy of antimicrobial drugs.3 Bacterial isolation and antimicrobial susceptibility testing are critical. Minimally invasive access to the middle ear through the dorsal roof of the tympanic bulla is a common surgical procedure in chinchillas used for human otologic disease research, and this technique can be used in a clinical setting for sterile sampling of the middle ear.15 Antibiotic selection should be based on susceptibility of the isolated organisms and the potential of the available drugs to eliminate bacteria effectively from the middle ear. Recommendations of fluoroquinolones and chloramphenicol for the treatment of otitis media in chinchillas are based on anecdotal evidence. Azithromycin (30 mg/kg PO q24h) reaches tissue levels high enough to recreate sterile conditions within the middle ear of chinchillas.4,19

RESPIRATORY SYSTEM DISORDERS Primary respiratory disease is uncommon in pet chinchillas. Historically, bacterial pneumonia was an important cause of mortality in ranched chinchillas and is still significant if husbandry is inadequate.8,14,16 Pneumonia is usually bacterial in origin. Important predisposing factors are poor husbandry, such as overcrowding, inadequate ventilation, and poor hygiene. Predominately gram-negative organisms have been isolated from chinchillas diagnosed with pneumonia.7,8,58 Clinical signs that may are tachypnea, dyspnea, and, in severe cases, open-mouth breathing. Presenting animals are often in poor body condition and have a poor hair coat. After ruling out other causes, such as congestive heart failure, initiate systemic antimicrobial therapy. Recommendations for treatment include oxygen therapy, systemic antibiotic therapy, and nebulization with antimicrobials (e.g., gentamicin, tobramycin). Once dyspnea is evident and an animal is in poor body condition, indicative of chronic disease, the prognosis is guarded to poor. Nasal discharge is relatively uncommon in chinchillas but can be associated with underlying dental disease, nasal foreign bodies, rhinitis, or lower respiratory tract disease. Base treatment on the underlying cause. Recurrent pneumonia and mucopurulent nasal discharge were associated with a megaesophagus in a 2-year-old chinchilla (see Fig. 24-2).63

REPRODUCTIVE SYSTEM DISORDERS ENDOMETRITIS AND PYOMETRA

Fig. 24-3  Dorsoventral skull radiograph of a 7-year-old chinchilla that presented for right-sided ear discharge. The changes of increased soft tissue opacity within the thickened bony walls of the right tympanic bulla are consistent with chronic otitis media.

Pet chinchillas may develop endometritis and pyometra. Affected animals can present with a history of acute onset of depression and anorexia or with mild lethargy or behavioral changes.69,136 Clinical signs vary, but an open vulva and vaginal discharge are present in most cases. Vaginal discharge can range from mucoid or mucopurulent to hemorrhagic. Anogenital fur staining may occur. Uterine enlargement or vaginal discharge may be evident on abdominal palpation. Radiographs and abdominal ultrasonography are helpful to evaluate the uterus. Vaginal cytology can be useful to differentiate metritis from physiologic vaginal discharge during estrus.9 Ovariohysterectomy has been recommended as the treatment of choice for endometritis and pyometra.136 In cases of mild endometritis with vaginal discharge, if the animal is in good condition, systemic antimicrobial therapy

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SECTION III  Guinea Pigs and Chinchillas

based on culture and susceptibility can be attempted.51 As in other species, prognosis will depend on the underlying and coexisting causes, such as hepatic lipidosis, ketosis, and possible complications in severe cases such as sepsis.

DYSTOCIA Signs of dystocia in chinchillas include restlessness, frequent crying, constant attention to the genital region, and a widened vaginal opening. Allantoic fluid or mucoid-hemorrhagic discharge from the vagina is often seen. Dystocia is usually associated with the presentation of a single oversized fetus or malpresentation of one or more kits.20 Uterine inertia has also been reported as a cause of dystocia.113 Use radiographs to assess the number, size, and position of the fetus(es). In an uncomplicated dystocia, lubrication and gentle traction of the fetus with feline obstetric forceps may correct the condition. Attempt treatment for uterine inertia with oxytocin (0.5-1 IU/kg SQ) and calcium gluconate (25-50 mg/kg SC diluted). Surgical intervention is imperative if the chinchilla is in labor for longer than 4 hours. Pet chinchillas respond well to cesarean section.113 Fetal resorption, mummification, retention, and abortion are common in chinchillas.137 Fetal death may be caused by a variety of infectious and noninfectious causes. Incompletely reabsorbed, retained, or mummified fetuses can remain in the uterus for extended periods and can predispose to sterility and endometritis. The ovarian bursa in chinchillas is underdeveloped and a case of an ectopic pregnancy has been reported.43 Several reports describe pulmonary trophoblastic emboli as incidental findings during postmortem examination in chinchillas; they have no clinical significance.57,132 While neoplasia in chinchillas is rare, several reports describe uterine leiomyosarcomas.

Fig. 24-4  Penile fur ring in an adult single-housed male chinchilla. The fur ring was an incidental finding during a routine physical examination and not associated with any clinical signs or complications.

PENILE DISORDERS Accumulation of hair (“fur ring”), smegma, or both around the base of the glans penis when it is enclosed within the prepuce is common in chinchillas (Fig. 24-4). Fur rings can be found in single-housed as well as breeding males. Male chinchillas that groom excessively, strain to urinate, frequently produce small amounts of urine, or repeatedly clean the penis may have a fur ring.59 The ring of hair can eventually stop the penis from retracting into the prepuce. In severe cases, an engorged penis is seen protruding from the prepuce, resulting in paraphimosis.59 The condition is not only painful but also may cause urethral constriction and acute urinary blockage. Treatment includes carefully removing any fur or debris from the penis. Topical ointments or gels are applied to prevent drying of the exposed penis and everted prepuce. For cases with underlying infection (Fig. 24-5), use systemic antimicrobial treatment. Complete resolution of infection and swelling is possible with topical and systemic treatment, but it may require long-term attention. Always examine the penis of male chinchillas during routine physical examination and remove any accumulated fur or secretions.

URINARY SYSTEM DISORDERS Male chinchillas can develop urinary calculi and urolithiasis.62,125 Quantitative mineral analysis of uroliths retrieved from 73 chinchillas showed that calculi were composed of calcium carbonate in approximately 90% of animals and of other

Fig. 24-5  Preputial abscess in an adult male chinchilla that presented for preputial swelling. The abscess is 2.5 cm in diameter. Despite the abscess, the animal maintained normal micturition.

material in 10%.108 Affected animals present with hematuria, stranguria, or anuria. Abdominal radiographs show radiodense calculi in the bladder and/or urethra. Treatment consists of surgical removal of the calculi if feasible. Submit swab samples of urine or bladder lining taken at surgery for bacterial culture and susceptibility testing to rule out underlying urinary tract infection. Because the underlying cause of urolith formation in chinchillas in unknown, recurrence of uroliths within a few weeks to months after surgical removal is possible. Currently there

CHAPTER 24  Disease Problems of Chinchillas are no specific recommendations for preventing reoccurrence of uroliths in affected chinchillas. Anecdotal recommendations include increased diuresis and a calcium-restricted diet. Spontaneous oxalate nephrosis in six female chinchillas has been reported.46 No access to ethylene glycol (antifreeze) had occurred and the underlying cause for the renal tubular oxalate crystal deposition could not be determined.

NEUROLOGIC DISORDERS SEIZURES Several reports of seizures in chinchillas are described.36,38,51,129,140 Encephalitis, septicemia, toxicosis, dietary deficiencies, hypocalcemia, hypoglycemia, hepatic or renal insufficiency, and heat stroke are described causing either generalized or focal convulsions. Hypersalivation, lateral recumbency, and nonresponsiveness are often seen.36,38 Although uncommon in pet chinchillas, consider infectious causes of encephalitis, such as listeriosis, human herpes simplex virus, or cerebrospinal nematodiasis (see below).119,140 Rule out extracranial causes, such as hypocalcemia, hyperthermia, hypoglycemia, organ failure, and lead poisoning.

HEAT STROKE The ambient temperature range to which chinchillas are adapted is 65°F to 80°F (18.3°C-26.7°C) in a low-humidity environment. Exposure to higher ambient temperatures, especially in the presence of high humidity and poor ventilation, can result in heat stroke. Affected animals are recumbent or ataxic, exhibit rapid breathing, and have bright red mucous membranes, prominent ear vessels, and thick, stringy saliva. Treatment includes cooling the animal and, if the animal is in shock, administering intravenous fluids. Long-term prognosis is guarded to poor and animals often deteriorate after initial improvement.51

LEAD POISONING Lead poisoning is uncommon in chinchillas and incidence varies based on geographic location. Two cases of lead toxicosis in pet chinchillas have been reported, one of which presented for seizures.53,96 Blood lead concentrations of 25 mg/dL or higher are indicative of lead poisoning. Successful treatment with calcium disodium edetate (30 mg/kg SC q12h) has been reported.53

DERMATOLOGIC DISORDERS DERMATOPHYTOSIS Dermatophytosis (ringworm) in chinchillas is most commonly caused by Trichophyton mentagrophytes, although Microsporum canis and Microsporum gypseum have been incriminated in outbreaks of spontaneously occurring dermatophytosis.29 Infected chinchillas may have small, scaly patches of alopecia on the nose, behind the ears, or on the forefeet. Lesions may appear on any part of the body; in advanced cases, a large circumscribed area of inflammation with scab formation is typical. Although most mycologic studies of chinchillas are based on animals with clinical signs, T. mentagrophytes has been

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cultured from 5% of fur-ranched chinchillas with normal skin and 30% of those with fur damage.29 Because T. mentagrophytes does not fluoresce, ultraviolet light is not useful for diagnosis. Diagnosis requires microscopic examination of hair and skin samples and dermatophyte culture. For topical therapy, 2% chlorhexidine/2% miconazole shampoo, or 0.2% enilconazole rinse is effective. Topical treatment removes spores from hair shafts, and systemic treatment acts at hair follicles. Systemic drugs that can be used are terbinafine (20-40 mg/kg PO q24h), itraconazole (5-10 mg/kg PO q24h), or ketoconazole (10-15 mg/kg PO q24h).90 Terbinafine is more effective than itraconazole against Trichophyton species in rodents.90 Continue treatment until two negative dermatophyte cultures have been obtained.

FUR CHEWING Fur chewing can be a common problem in chinchillas; up to 20% of animals in breeding facilities can be affected (see Fig. 22-8). Fur chewing is also commonly seen in chinchillas suffering from dental disease.61,131 Many theories concerning the cause of fur chewing have been proposed, including dietary deficiencies, fungal and endoparasite infections, hormonal dysregulation, environmental stress, boredom, and genetic predisposition.33,131,134 Scant scientific evidence exists for most proposed theories. Currently, the most widely accepted theory suggests that fur chewing is a maladapted displacement behavior triggered by stress and affecting predominately stress-­sensitive animals. Adrenocortical hyperplasia and histologic changes of the skin, correlating with increased cortisol secretion, support this theory.131 Clinical signs vary and affected animals may chew their own or their cagemate’s fur, resulting in a coat with a moth-eaten appearance. Fur-chewed areas occur usually along the midspinal area from the lumbar part to the tail. In animals that chew their own fur, the head and distal extremities are usually not affected and the chewed areas are usually covered with short, darker-colored fur.131 Perform a thorough history and physical examination to rule out dietary deficiencies, dental disease, and possible environmental stressors. A variety of dermatologic diagnostic tests can be considered, including dermatophyte culture and skin biopsy, although dermatophytes can be cultured from animals without fur lesions and do not necessarily represent the primary underlying cause of fur damage.33 Obtaining a definitive diagnosis and identifying the primary underlying cause of fur chewing behavior may be difficult. While fur chewing can be annoying for the pet chinchilla owner, it is not a significant threat to the animal’s health. Consider environmental and dietary improvements after ruling out infectious and organic underlying causes. This may include reducing possible stressors, such as frequent handling and light and noise disturbance. Carefully review social dynamics if housing multiple animals together. Avoid overcrowding, and separate affected animals from dominant or aggressive cage mates. Offer multiple sleeping boxes and feeding spots for animals housed in groups. Recommend environmental enrichment such as providing high-quality grass hay and branches for chewing and foraging as well as a structured enclosure and regular exercise. Treatment with antidepressants such as fluoxetine hydrochloride (5 to 10 mg/kg PO q24h) has been suggested for other rodents, but no clinical results have been reported in chinchillas.116

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SECTION III  Guinea Pigs and Chinchillas

FUR SLIP Fur slip is a predator avoidance mechanism in chinchillas. When an animal is fighting or roughly handled, it can release a large patch of fur, thus enabling it to escape. A clean, smooth area of skin is left; hair may require several months to regrow.

MATTED FUR Matted fur can develop in chinchillas, especially if they are kept in a warm (>80°F [26.7°C]), humid environment or if they are deprived of dust baths (see Chapter 22). Provide dust baths for approximately 15 minutes per day and reduce humidity levels if necessary.

FOOT DISORDERS In chinchillas, foot disorders predominately affect the hind feet. Lesions can include hyperkeratosis and erythema; less commonly, deep infections or open lesions of the plantar aspect of the feet can occur. Possible predisposing factors are obesity, inappropriate cage flooring or substrate, and poor hygiene. In severe cases, infection can involve the tendons and bones. Treatment depends on the severity of disease. In mild cases, environmental improvements and application of petroleum-based ointment are often sufficient to resolve hyperkeratosis and erythema. Advise owners to reduce body weight gradually in overweight animals. In severe cases, perform surgical debridement followed by open-wound management and bandaging until healing is complete.

MISCELLANEOUS DISEASE PROBLEMS CARDIAC DISEASE Heart murmurs ranging from mild to moderate are often auscultated, particularly in young chinchillas presented for routine examination.55 To date, reports of cardiac disease in chinchillas are scant and the significance of heart murmurs in young, clinically healthy animals remains unknown. Anecdotally, cardiomyopathy, ventricular septal defect, as well as mitral and tricuspid valve insufficiencies have been reported.55,76 Use echocardiography to differentiate innocent from pathologic murmurs. Echocardiographic reference values for chinchillas have been published.76 At present, the clinical management of heart disease in chinchillas is empiric.

HEPATIC LIPIDOSIS AND KETOSIS Hepatic lipidosis because of excessive fat mobilization and, less frequently, toxicosis is one of the most common findings in chinchillas at necropsy.32,112 Chinchillas with hepatic lipidosis often have a history of anorexia and decreased fecal output and, in advanced cases, can be depressed and dehydrated. Hyperglycemia, pronounced ketonuria, and possible glucosuria and acidosis can develop.92 The negative energy balance in hyporectic or anorectic chinchillas leads to increased mobilization of lipids and excessive beta oxidation of fatty acids in hepatocytes, promoting hepatic ketogenesis. Treatment should correct the underlying primary

cause leading to anorexia (e.g., dental disease, gastroenteritis), along with correcting fluid deficits and providing nutritional support. Measure urinary ketones repeatedly as a simple and noninvasive tool to monitor the response to treatment.

DIABETES MELLITUS Diabetes mellitus is rare in chinchillas. Treatment is difficult and the prognosis is poor. Only two cases (suggestive of type 2 diabetes) have been reported.37,81 When presented with an anorexic and hyperglycemic chinchilla, rule out common causes for anorexia (e.g., dental disease, pain). Many chinchillas will become ketoacidotic and hyperglycemic secondary to anorexia. Determine serum insulin levels to confirm a diagnosis of diabetes mellitus caused by endocrine pancreatic insufficiency. Hyperglycemia secondary to peripheral insulin resistance (type 2), as seen in cases of ketoacidosis, should be treated by focusing on the underlying primary cause of anorexia and providing nutritional support. In two cases, a 5-year-old obese female81 and a 1-year-old male37 had blood glucose levels elevated to at least four times the upper limit of the reference range, as well as severe glucosuria and ketonuria. Both animals were treated with insulin (2 IU daily and increasing to 12 IU daily81 and 4 IU daily increasing to 6 IU daily37). After initial improvement, both animals died. Postmortem examination showed atrophy of the pancreas and vacuolation of the islets of Langerhans in the obese female chinchilla and a pancreatic adenoma in the 1-year-old male. Hypoglycemia is always a great risk in treating diabetes with recombinant human insulin or porcine insulin.100,107 Consequently any attempt to treat diabetes with insulin must be conservative and dramatic increases in the insulin dose should be avoided. Ancillary treatment should be aimed at reducing obesity and feeding a diet that is high in protein, low in fat, and high in complex carbohydrates.

FRACTURES Traumatic fractures of the tibia are commonly seen. The tibia is a long straight bone with little soft tissue covering. It is longer than the femur, and the fibula is virtually nonexistent.18 Tibial fractures are usually either transverse or short spiral fractures. Tibial fracture often occurs when a chinchilla is grabbed by its hind limb or a hind limb catches in a cage bar. Like the bones of rabbits, those of chinchillas are thin and fragile; surgical repair can be difficult and complications are common (see Chapter 33). Soft, padded bandages and lateral splints usually do not provide adequate stability for tibial fractures to heal.54 External fixation and intramedullary pins, alone or in combination, have been recommended for surgical stabilization of tibial fractures in chinchillas.38 Restricted exercise in a single-level enclosure, ideally without cage bars, is necessary. The prognosis for tibial fractures is guarded and complications after surgical fixation are common. These include bone-pin loosening and infection, nonunion, necrosis of the distal limb, and automutilation.38 Consider hind-limb amputation if surgical fracture stabilization fails or is not indicated. Chinchillas usually adapt very well after amputation.26,70,130 Fractures

CHAPTER 24  Disease Problems of Chinchillas of the fore limbs distal to the elbow can be managed by external cooptation and splinting; chinchillas usually tolerate such treatment well.

NEOPLASIA Despite the long life span of chinchillas compared with other rodents, references to neoplasia in chinchillas are rare. Postmortem examinations on 1,005 fur-ranched chinchillas before 194913 and another 1,000 fur-ranched chinchillas between 1949 and 195214 ranging in age between less than 6 months and 11 years did not list neoplasia as a cause of death. However, tumors such as neuroblastoma, carcinoma, lipoma, and hemangioma were reported in fur-farmed chinchillas in the annual reports of the San Diego County Livestock Department during the 1950s.101 Individual case reports of neoplasia have included two cases of lymphosarcoma.65,101 Both cases occurred in young male animals (18 months and 4.5 years) and both animals had generalized lymphadenopathy and neoplastic cell infiltration in multiple organs. Four other reports described a nonmetastasizing cholangiohepatic carcinoma in a 3-year-old female chinchilla102; a fibrosarcoma located at the tail base of a 6-year-old male chinchilla38; a nonmetastasizing lumbar osteosarcoma in a 13-year-old female chinchilla123; and an undifferentiated carcinoma of the salivary gland in a 12-year-old female chinchilla.124 A 5-year retrospective from 1991 to 1996 of chinchillas presented to the Animal Medical Center in New York City revealed only one case of a uterine leiomyosarcoma with no associated metastases. A 35-year retrospective evaluation of chinchilla presentations to the University of Tennessee College of Veterinary Medicine showed only two histopathologically documented cases of neoplasia: one was lymphosarcoma and the other adenocarcinoma of the lung.47 A 9-year retrospective between 1990 and 1999 of chinchillas submitted for necropsy at the Institute of Pathology and Forensic Veterinary Medicine in Vienna, ­Austria, documented a low incidence of neoplasia in chinchillas (2.6%) compared with rabbits, guinea pigs, rats, and mice.39 Three of 115 chinchillas had tumors. The cases included a ­2-year-old female with uterine leiomyoma; a 3-year-old male with hemangioma of the subcutis; and a castrated 10-year-old male with an adenoma of the pituitary gland. A second 9-year retrospective between 1994 and 2003 of 325 chinchillas presented to the University of Zurich Veterinary Hospital revealed tumors in only three animals (1%). During the same period, the incidence of neoplasia was higher in rabbits and rodents compared with chinchillas (guinea pigs 7%, rats 34%, and rabbits 6%).72 The low number of reports of neoplasia up to the present may reflect the emphasis on chinchillas as fur producers or research animals, since chinchillas were bred for their fur or kept as laboratory animals before they became popular as pets. Therefore ‘‘geriatric’’ conditions, including neoplasia, may not have been represented in the older reports. Alternatively, the 5- to 35-year surveys of neoplasia in exotic pets presented to veterinary specialty centers suggest that the low incidence of neoplasia in chinchillas may reflect a true low frequency compared with the incidence in other rodents and rabbits. Further clinical surveys are needed to elucidate this question.

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INFECTIOUS DISEASES VIRAL INFECTIONS Diagnoses of viral infections in chinchillas are rare. During several epizootic disease outbreaks in ranched chinchillas, a viral etiology was suspected but diagnosis could not be established.20,38 Chinchillas are susceptible to human herpesvirus type 1 and may play a role as a temporary reservoir for human infections. A spontaneous herpes-like viral infection was reported in a female chinchilla.45 In a 1-year-old male chinchilla with a 2-week history of conjunctivitis and subsequent neurologic signs of seizures, disorientation, recumbency, and apathy, histologic examination showed a nonsuppurative meningitis and polioencephalitis with neuronal necrosis and intranuclear inclusion bodies; these data were confirmed as indicating herpesvirus type 1.140 Both eyes displayed ulcerative keratitis, uveitis, retinitis and retinal degeneration, and optical neuritis. The clinical signs, the distribution of the lesions, and the viral antigen suggested a primary ocular infection with subsequent spread to the central nervous system.

FUNGAL INFECTIONS There are two case reports of Histoplasma capsulatum infection in chinchillas. One case was in a chinchilla imported from the United States to Switzerland.17 The second case was in a female chinchilla that originated from a commercial chinchilla ranch in central Missouri.109 At necropsy, pulmonary lesions included multiple hemorrhagic foci, alveolar consolidation, and bronchopneumonia; the organism was found to be present in numerous giant cells. Multifocal pyogranulomatous splenitis and hepatitis, with H. capsulatum in giant cells, was also noted. Histoplasma capsulatum was subsequently cultured from timothy hay used for food. Cyniclomyces guttulatus is part of the physiologic intestinal flora in chinchillas.145 Animals with soft feces or diarrhea often show increased numbers of this yeast. This finding is indicative of dysbacteriosis, which leads to opportunistic overgrowth of C. guttulatus.38,51 Treatment should address the primary underlying cause of the dysbacteriosis. In severe cases of yeast overgrowth, consider treatment with nystatin (100,000 IU/kg PO q8h for 5 days). Aflatoxicosis is an acute, fatal disease resulting from toxicity from improperly stored feed contaminated with Aspergillus fungi. In one report, the death of 200 chinchillas was attributed to high concentrations of aflatoxin B-1 in the feed.112 The liver is the primary target organ of aflatoxin; in affected animals, it is enlarged, pale yellow, and friable. In the described cases, histopathologic analyses of hepatic parenchyma showed severe, diffuse cytoplasmic vacuolation of hepatocytes.112 Because the histopathologic changes caused by acute aflatoxicosis are nonspecific, the diagnosis of aflatoxicosis is usually made in combination with mycotoxicologic feed analysis.112

PARASITIC INFECTIONS Protozoal Infections Historically, group-housed chinchillas in fur ranches and research colonies had a high prevalence of giardiasis.122 However, the role of Giardia duodenalis (syn. G. lamblia) in

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causing disease in chinchillas is difficult to establish. Giardia are rarely found in fecal samples from wild chinchillas, but Giardia is found in both healthy and sick captive-bred animals.33,40 In pet chinchillas, G. duodenalis is highly prevalent. In 80 pet chinchillas from both pet owners and breeders in Belgium screened for Giardia, 53 (66%) excreted Giardia cysts. Young animals and animals participating in shows were at significantly higher risk of infection.74 A total of 22 isolates based on genetic groups (or assemblages) were characterized. Healthy chinchillas can harbor G. duodenalis organisms in low numbers in the small intestine, and experimental infection of healthy chinchillas with Giardia cysts failed to induce clinical disease.33,38 Predisposing factors, such as stress and poor husbandry, are believed to cause an increase in parasite numbers, resulting in diarrhea and potentially death. Recently weaned juvenile animals seem to be prone to developing clinical signs.38,74 Signs of giardiasis in pet chinchillas can include a cyclic sequence of appetite loss and diarrhea associated with a declining body and fur condition. Zinc sulfate flotation allows identification of cysts, and trophozoites can be identified in fresh fecal smears in acute cases of heavy infection. The use of an antigen enzyme-linked immunosorbent assay (ELISA) (Remel ProSpecT microplate assay; Thermo Fisher Scientific, Lexana, KS) for detecting G. duodenalis antigen in chinchilla feces has recently been reported.110 However, the sensitivity and specificity for this test in chinchillas has not been determined, and because clinically normal chinchillas can also shed G. duodenalis in the feces, the significance of a positive ELISA is of questionable clinical significance.33,40 Quantifying fecal cyst or trophozoite numbers in animals with diarrhea appears to be more clinically relevant to determine if an infection is related to the clinical signs. Assemblage specific polymerase chain reaction (PCR) (e.g., A-G) is clinically and epidemiologically beneficial, as mixed genetic group infections are likely to be missed using conventional PCR approaches.74 Treat chinchillas with giardiasis with metronidazole, albendazole, or fenbendazole. It is unknown if these compounds eradicate Giardia completely or only inhibit cyst production; therefore treated animals may remain a source of chronic cyst shedding. Treat all animals in contact and thoroughly disinfect the environment to prevent reinfection. Replace wooden cage interior parts, such as resting boards. Giardia cysts remain infectious for up to several weeks in a cool, humid environment. Until further research in chinchillas is done, the pathogenicity of G. duodenalis and its association with disease remain controversial. Giardiainfected chinchillas are a potential reservoir for zoonotic transmission.64,74,121 Eimeria chinchillae is strictly host-specific and can occasionally cause enteritis and diarrhea predominately in younger animals. Stress likely leads to clinical manifestation of a previous subclinical infection. This coccidium causes damage of the intestinal mucosa, with subsequent disturbance of the physiologic flora and possible secondary dysbacteriosis. Diagnosis is made by fecal flotation. Treatment with sulfonamide compounds (e.g., sulfadimethoxine) usually resolves clinical signs and cyst shedding. It is unknown if these compounds are able to eradicate E. chinchillae. As for giardiasis, treatment of contact animals and thorough environmental disinfection is mandatory.

Although toxoplasmosis was common in fur-ranched chinchillas in the past, it is now rarely seen.66 At necropsy, lesions include an enlarged spleen and mesenteric lymph nodes as well as hemorrhagic lungs. In two chinchillas with focal necrotic meningoencephalitis caused by Toxoplasma gondii,87 several lobulated Frenkelia cysts up to 0.6 mm in diameter were found in the brains, independent of and remote from the toxoplasmal inflammatory reaction. The authors considered that chinchillas might be susceptible to a Frenkelia species occurring in other free-living species. Tissue cysts of Frenkelia microti were found in the brain of a chinchilla bred in Minnesota and used for biomedical research.30 This was the first report of Frenkelia infection in chinchillas in the United States. A case of acute hepatic sarcocystosis was reported in a pet female chinchilla.114 The owners had housed and fed it with six other chinchillas that remained healthy, and the source of infection was unknown. Gastroenteritis associated with Cryptosporidium species was described in an 8-month-old pet chinchilla that originated from a pet shop.143 Meningitis and optic nerve neuritis due to Cryptococcus species was reported in a 13-year-old chinchilla.10

Helminthic Infections Recent studies report a low prevalence of nematode and cestode infections in pet chinchillas.110 Haemonchus contortus, Trichostrongylus colubriformis, Ostertagia ostertagi, and an unspecified Oxyuris species have been reported in chinchillas.110,127 Disease outbreaks of cerebral nematodiasis caused by the raccoon ascarid Baylisascaris procyonis have been reported in chinchillas from western Canada.119 Affected chinchillas showed ataxia, torticollis, paralysis, incoordination, and tumbling. Outbreaks of fatal central nervous system disease were linked to use of hay contaminated by raccoon feces. Raccoons infected with B. procyonis are more common in temperate regions of North America, especially the midwestern and northeastern United States. Cestodes. Rodentolepis nana (previously Hymenolepis nana) infections are reported in chinchillas.44,105,127 This cestode does not require an intermediate host and infection can occur by direct transmission via fecal-oral route.120 Animals infected by high numbers of R. nana can show anorexia, diarrhea, weight loss, and death,98 but subclinical infections are more common. Rodentolepis nana is zoonotic and can cause severe infections, particularly in immunocompromised humans.31,106 Demonstrate Rodentolepis eggs by fecal flotation. Treat with praziquantel (5 mg/kg PO or SQ q10d). Chinchillas can serve as intermediate hosts for cestodes, including Taenia serialis, Taenia pisiformis, Taenia multiceps, Echinococcus granulosus, and Echinococcus multilocularis.44,56,126 In the 1950s, infections were seen when chinchillas were given feed accidentally contaminated with infected dog feces. A recent report described more than 600 cysts of Taenia crassiceps found in the abdominal cavity of a pet chinchilla imported from the Netherlands into Japan.71

BACTERIAL INFECTIONS Opportunistic bacterial infections in chinchillas can cause disease, which is localized either to one organ or as septicemia. Affected animals are usually immunocompromised by age, underlying disease, nutritional status, or husbandry-related factors (poor hygiene, poor ventilation, contaminated feed).

CHAPTER 24  Disease Problems of Chinchillas Members of the family Enterobacteriaceae and P. aeruginosa have been associated with significant morbidity and mortality in chinchillas.* However, Enterobacteriaceae and P. aeruginosa can also be isolated from clinically healthy animals.12,83,91 Therefore most of these organisms are not considered primary pathogens. Pseudomonas aeruginosa infections and epizootic outbreaks in chinchillas have been reported frequently.49,79,89,138 Initially, infections are often localized to one organ and are associated with conjunctivitis, enteritis, pneumonia, otitis media and interna, metritis, and abortion. As the disease progresses, systemic spread is common. In addition, an acute generalized form with septicemia and often sudden death can occur. P. aeruginosa can be part of the normal intestinal flora in healthy chinchillas. A survey of 67 healthy pet or laboratory chinchillas isolated the organism from 42% of animals.52 Stress, intercurrent disease and/or or contaminated drinking water predispose to infection and clinical disease.28,83,91,138 Conjunctivitis is a common initial sign of Pseudomonas infection in chinchillas. Anorexia, lethargy, and decreased fecal output often follow.27,28,138 In a case of P. aeruginosa infection described in a laboratory chinchilla, the affected animal displayed a variety of clinical signs, including conjunctivitis, scrotal swelling, anorexia, weight loss, and corneal and oral ulcerations.28 Characteristic pathologic lesions are miliary necrosis in the internal parenchymal organs and a necrotizing typhlocolitis. Multidrug-resistant, reduced-antibiotic susceptibility and highly virulent strains of P. aeruginosa are widespread in chinchillas.52 Base antimicrobial drug selection on culture and susceptibility testing. Because affected animals are often in a critically compromised condition, empiric drug selection is necessary. Generally, P. aeruginosa is susceptible to fluoroquinolones, third-generation cephalosporins, and aminoglycosides.117,118 Use topical polymyxin B and gentamicin-containing formulations to treat because of the low prevalence of isolates that are resistant to these drugs. A vaccine against P. aeruginosa has been developed for attempted immunization and is used in fur-ranched chinchillas.79,84 Escherichia coli is not considered part of the physiologic intes­tinal flora of healthy captive chinchillas but has been isolated from healthy wild-caught chinchillas.12,83 It is a ubiquitous environmental organism and an opportunistic pathogen. The ingestion of large numbers of organisms, a primary underlying disease, or predisposing risk factors are prerequisites to infection and clinical disease. Pathogenicity depends on the serotype, enteropathogenicity, and endotoxin production. Clinical signs associated with E. coli-associated enteritis can very be similar to enteritis caused by P. aeruginosa, such as constipation, diarrhea, variable form and size of fecal pellets, mucoid or hemorrhagic adhesions to fecal pellets, depression, and anorexia. Sudden death due to endotoxemia or septicemia is possible.12 In a study of chinchillas experimentally infected with enteropathogenic E. coli via drinking water, animals became increasingly depressed, exhibited a stretched body posture, initially had diarrhea, and later had reduced fecal output and anorexia.12 In chinchillas that died, E. coli was cultured from all internal organs and the animals

*References 7, 8, 12, 28, 88, 138.

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had suffered from gastroenteritis. Pure growth of E. coli was seen in fecal cultures 4 to 7 days after the start of the experiment. Parenteral treatment with antibiotics caused a significant improvement of clinical signs and disappearance of E. coli from fecal cultures.12 Yersinia pseudotuberculosis (formerly Pasteurella pseudotuberculosis) and Yersinia enterocolitica occur worldwide in areas of moderate and subtropical climate, and outbreaks in chinchillas are commonly described.34,41,42,48,73 The species most frequently isolated from chinchillas is Y. enterocolitica. Yersiniosis is an enteric disease that damages epithelium of the ileum, cecum, and colon, resulting in mucosal hemorrhage and ulceration. Lymphoid infiltration results in hypertrophy of Peyer’s patches and mesenteric lymph nodes and necrotizing granulomas. Systemic spread results in granulomatous lesions in the lungs, spleen, and liver and eventual death. A “chinchillatype” strain of Y. enterocolitica (biovar 3, antigens or serovar 1, 2a, 3) appears to persist enzootically among chinchilla stock worldwide.141 Listeriosis in chinchillas was first reported by MacKay et al. in 1949.80 It was and still is common in fur-ranched chinchillas but not in laboratory or pet chinchillas.67,68,103,139 Listeria monocytogenes is a highly adaptable environmental bacterium that can exist as both an animal pathogen and a plant saprophyte; it is also part of the normal microbial flora in healthy ruminants and is found in environmental sources such as decaying vegetation. Most animal and human cases of listeriosis arise from the ingestion of contaminated food; the disease is common in animals fed on silage.78 Unlike most food-borne pathogens that cause gastrointestinal disease, L. monocytogenes causes several easily recognized invasive syndromes, such as encephalitis, abortion, and septicemia. In chinchillas, listeriosis is a cecal disease with blood-borne dissemination. The main target organ is the liver, where the bacteria multiply inside hepatocytes, followed by cell lysis, bacterial release, septicemia, and, in surviving hosts, the development of lung, brain, spleen, lymph node, and liver abscesses. Enterotoxaemia associated with Clostridium species in chinchillas has been reported.95 Enterotoxemia caused by Clostridium perfringens enterotoxin has been described as well as deaths due to C. perfringens A enterotoxemia.6,104 Salmonellosis characterized by gastroenteritis and abortion has been reported often from ranched chinchillas; it causes significant morbidity and mortality.44,93,99 There are two case reports of Salmonella infection in pet chinchillas. Salmonella arizona septicemia was reported in a chinchilla in the United Kingdom97 and septic infection with Salmonella enteritidis was described in a companion chinchilla in Japan.142

References 1. Alper CM, Doyle WJ, Seroky JT, et al. Efficacy of clarithromycin treatment of acute otitis media caused by infection with penicillin-susceptible, -intermediate, and -resistant Streptococcus pneumoniae in the chinchilla. Antimicrob Agents Chemother. 1996;40:1889-1892. 2. Antonelli PJ, Winterstein AG, Schultz GS. Topical dexamethasone and tympanic membrane perforation healing in otitis media: a short-term study. Otol Neurotol. 2010;31:519-523. 3. Apicella MA. Bacterial otitis media, the chinchilla middle ear, and biofilms. J Infect Dis. 2009;199:774-775.

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4. Babl FE, Pelton SI, Li Z. Experimental acute otitis media due to nontypeable Haemophilus influenzae: comparison of high and low azithromycin doses with placebo. Antimicrob Agents Chemother. 2002;46:2194-2199. 5. Bakaletz LO. Chinchilla as a robust, reproducible and polymicrobial model of otitis media and its prevention. Expert Rev Vaccines. 2009;8:1063-1082. 6. Bartoszce M, Nowakowska M, Roszkowski J, et al. Chinchilla deaths due to Clostridium perfringens A enterotoxin. Vet Rec. 1990;126:341-342. 7. Bartoszcze M, Matras J, Palec S, et  al. Klebsiella pneumoniae infection in chinchillas [letter]. Vet Rec. 1990;127:119. 8. Bautista E, Martino P, Manacorda A, et al. Spontaneous Proteus mirabilis and Enterobacter aerogenes infection in chinchilla (Chinchilla lanigera). Scientifur. 2007;31:27-30. 9. Bekyürek T, Liman N, Bayram G. Diagnosis of sexual cycle by means of vaginal smear method in the chinchilla (Chinchilla lanigera). Lab Anim. 2002;36:51-60. 10. Bicknese E, White A, Pessier A, et al. Cryptococcal meningitis and optic nerve neuritis in a chinchilla (Chinchilla lanigera). Proceedings. Annu Conf Assoc Exot Mammal Vet. 2010:Session #165. 11. Boehmer E, Crossley D. Objective interpretation of dental disease in rabbits, guinea pigs and chinchillas. Use of anatomical reference lines. Tierarztl Prax Ausg K Klientiere Heimtiere. 2009;37:250-260. 12. Brem M. Investigations about the diseases of the gastrointestinal tract in the Chinchilla. [German] Untersuchungen über Erkrankungen des Magen-Darmkanals beim Chinchilla. Medizinische Tierklinik. Ludwig-Maximilians Universität: Munich; 1982;125. 13. Brenon HC. Postmortem examinations of chinchillas. J Am Vet Med Assoc. 1953;123:310. 14. Brenon HC. Postmortem examinations of chinchillas. J Am Vet Med Assoc. 1955;126:222-223. 15. Brown C. Middle ear sample collection in the chinchilla. Lab Anim (NY). 2007;36:22-23. 16. Burtscher H. Pathological anatomy of chinchilla diseases. ­[German] Pathologische Anatomie der Chinchillakrankheiten. Dtsch Tierarztl Wochenschr. 1965;72:376-380. 17. Burtscher H, Otte E. Histoplasma in the chinchilla. [German] Histoplasme beim chinchilla. Dtsch Tierarztl Wochenschr. 1962;69:303-307. 18. Cevik-Demirkan A, Ozdemir V, Turkmenoglu I, et  al. Anatomy of the hind limb skeleton of the chinchilla (Chinchilla lanigera). Acta Veterinaria Brno. 2007;76:501-507. 19. Chan KH, Swarts JD, Doyle WJ, et al. Efficacy of a new macrolide (azithromycin). For acute otitis media in the chinchilla model. Arch Otolaryngol Head Neck Surg. 1988;114:1266-1269. 20. Cousens PJ. The chinchilla in veterinary practice. J Small Anim Pract. 1963;4:199-205. 21. Crossley DA. Dental disease in chinchillas in the UK. J Small Anim Pract. 2001;42:12-19. 22. Crossley DA. Dental disease in chinchillas [dissertation]. ­Manchester, UK: School of Dentistry, University of Manchester; 2003;263. 23. Crossley DA, Dubielzig RR, Benson KG. Caries and odontoclastic resorptive lesions in a chinchilla (Chinchilla lanigera). Vet Rec. 1997;141:337-339. 24. Crossley DA, Miguélez MM. Skull size and cheek-tooth length in wild-caught and captive-bred chinchillas. Arch Oral Biol. 2001;46:919-928. 25. Crossley DA, Roxburg G, Miguélez Vidales MM. Anatomy of the chinchilla (Chinchilla lanigera) lacrimal drainage system and its obstruction in dental disease. Proceedings. 8th Europ Vet Dental Soc. 1999:21-22. 26. Degasperi B, Mosing M, Kunzel F. Limb amputation as salvage procedure in small mammals. [German]. Extremitatenamputation als rettende Massnahme bei kleinen Heimtieren. Wien Tierarztl Monatsschr. 2007;94:93-98.

27. Devos A, Van Impe J, Viaene N, et al. Pseudomonas aeruginosa infection in chinchillas. [Dutch]. Vlaams Diergeneesk Tijdschr. 1966;35:222-232. 28. Doerning BJ, Brammer DW, Rush HG. Pseudomonas aeruginosa infection in a Chinchilla lanigera. Lab Anim. 1993; 27:131-133. 29. Donnelly TM, Rush EM, Lackner PA. Ringworm in small exotic pets. Sem Avian Exot Pet Med. 2000;9:82-93. 30. Dubey JP, Clark TR, Yantis D. Frenkelia microti infection in a chinchilla (Chinchilla laniger) in the United States. J Parasitol. 2000;86:1149-1150. 31. Duclos LM, Richardson DJ. Hymenolepis nana in pet store rodents. Comp Parasitol. 2000;67:197-201. 32. Egri B, Egri J, Hajnovics B. Fatty liver in young male chinchilla (Chinchilla velligera). [German] Uber Fettinfiltration der Leber beim Chinchillabockchen (Chinchilla velligera). Tierarztl Umsch. 1994;49:42, 45-47. 33. Eidmann S. Studies on the etiology and pathogenesis of fur damage in the chinchilla. [German] Untersuchungen zur Atiologie und Pathogenese von Fellscaden beim Chinchilla. Institut fur Pathologie. Hannover, Germany: Tierarztliche Hochschule; 1992;163. 34. Emirsajlow Zalewska W, Furowicz AJ, Aleksic S, et  al. Evaluation of pathogenicity determinants of Yersinia pseudotuberculosis strains isolated from chinchillas from the Western Pomerania area. Advances in Agricultural Sciences. 1996;5:19-24. 35. Eule C, Sjoberg JG. Corneal injuries in chinchillas. Exot DVM. 2007;9:7-8. 36. Ewringmann A, Gloeckner B. Neurological deficits. ­[German] Neurologische Ausfallerscheinungen. In: Ewringmann A, Gloeckner B, eds. Leitsymptome be Meerschweinchen, Chinchilla und Degu. Stuttgart: Enke; 2005:166-185. 37. Ewringmann A, Gobel T. Diabetes mellitus in rabbits, guinea pigs and chinchillas. [German] Diabetes mellitus bei Kanin­chen, Meerschweinchen und Chinchilla. Kleintierpraxis. 1998;43:337-348. 38. Fehr M. Chinchilla. In: Fehr M, Sassenburg L, Zwart P, eds. Krankheiten der Heimtiere. 6th ed. Hannover: Schluetersche; 2005:183-213. 39. Ferfschl S. Spontaneous tumors in horses, cattle and small pets: findings of the institute of pathology and forensic veterinary medicine in the years 1990-1999. [German] Spontane Tumoren bei Pferden, Rindern und kleinen Heimtieren: Untersuchungsergebnisse des Instituts für Pathologie und Gerichtliche Veterinärmedizin in den Jahren 1990-1999. Institut für Tierzucht und Genetik. Vienna: Veterinaermedizinische Universitaet Wien; 2004;117. 40. Fialho CG, Oliveira RG, Teixeira MC, et  al. Comparison of protozoan infection between chinchillas (Chinchilla lanigera) from a commercial breeding facility in southern Brazil and chinchillas from a natural reserve in Chile. [Portuguese] Comparação da infecção por protozoários em chinchila (Chinchilla lanigera) de uma criação comercial do município de ViamãoRS, Brasil, e de chinchilas no seu habitat natural, Chile. Parasitol Latinoam. 2008;63:85-87. 41. Furowicz AJ, Czernomysy-Furowicz D. Eradication of Yersinia pseudotuberculosis infection from a chinchilla farm. [Polish] Eliminacja jersiniozy (Yersinia pseudotuberculosis) z hodowli szynszyli. Magazyn Weterynaryjny. 1999;8:130-132. 42. Furowicz AJ, Czernomysy-Furowicz D, Misiura M, et  al. Yersiniosis in chinchillas caused by the enterotoxigenic strain of Yersinia pseudotuberculosis. [Polish] Enterotoksyczny szczep Yersinia pseudotuberculosis przyczyna jersiniozy szynszyli. Medycyna Weterynaryjna. 1996;52:116-118. 43. Gitlin G, Adler JH. Coexisting intrauterine and abdominal (intraperitoneal) pregnancy with possible superfoetation (superfecundation) and with adhesion of placenta to foetus in a Chinchilla (Chinchilla laniger). Acta Zool Pathol Antverp. 1969;49:65-76.

CHAPTER 24  Disease Problems of Chinchillas 44. Gorham JR, Farrell K. Diseases and parasites of chinchillas. Proceedings. 92nd Ann Meet Am Vet Med Assoc. 1955:228-234. 45. Goudas P, Giltoy JS. Spontaneous herpes-like viral infection in a chinchilla (Chinchilla laniger). J Wildl Dis. 1970;6:175-179. 46. Goudas P, Lusis P. Case report. Oxalate nephrosis in chinchilla (Chinchilla laniger). Can Vet J. 1970;11:256-257. 47. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 48. Gueraud JM. Threat of an epidemic of yersiniosis in chinchillas. [French] Vers une epizootie de yersiniose chez le chinchilla. Bull Acad Vet Fr. 1988;61:95-98. 49. Halen P, Pohl P, Thomas J. Septicaemia caused by Pseudomonas in mink and chinchillas. [French]. Septicémies à pseudomonas chez les visons et chinchillas. Ann Med Vet. 1966;110:397-406. 50. Hallowell GD. Retrospective study assessing efficacy of treatment of large colonic impactions. Equine Vet J. 2008; 40:411-413. 51. Hansen D. Chinchilla. In: Goebel T, Ewringmann A, eds. Heimtierkrankheiten. Stutgart: Verlag Eugen Ulmer; 2005:100-118. 52. Hirakawa Y, Sasaki H, Kawamoto E, et  al. Prevalence and analysis of Pseudomonas aeruginosa in chinchillas. BMC Vet Res. 2010;6:52. 53. Hoefer HL. Chinchillas. Vet Clin North Am Small Anim Pract. 1994;24:103-111. 54. Hoefer HL. Chinchillas. Proceedings. North Am Vet Conf. 1995:672-673. 55. Hoefer HL, Crossley DA. Chinchillas. In: Meredith A, Redrobe S, eds. BSAVA Manual of Exotic Pets. 4th ed. Quedgeley, Gloucester: British Small Animal Veterinary Association; 2002:65-75. 56. Holmberg BJ, Hollingsworth SR, Osofsky A, et  al. Taenia coenurus in the orbit of a chinchilla. Vet Ophthalmol. 2007;10:53-59. 57. Ilha MRdS, Bezerra Junior PS, Sanches AWD, et al. Trophoblastic pulmonary embolism in chinchillas (Chinchilla laniger). [Portuguese] Embolia pulmonar trofoblastica em chinchilas (Chinchilla laniger). Ciencia Rural. 2000;30:903-904. 58. Ioakimidis I, Tomopoulos D, Vlaikidis N. An outbreak of Bordetella bronchiseptica infection in chinchillas (Chinchilla laniger). Hellenike Kteniatrike. 1970;13:31-33. 59. Ivey ES, Hoefer HL. What’s Your Diagnosis? Pollakiuria in a chinchilla. Lab Anim (NY). 1998;27:21-22. 60. Jauris-Heipke S, Leake ER, Billy JM, et al. The effect of antibiotic treatment on the release of endotoxin during nontypable Haemophilus influenzae-induced otitis media in the chinchilla. Acta Otolaryngol. 1997;117:109-112. 61. Jekl V, Hauptman K, Knotek Z. Quantitative and qualitative assessments of intraoral lesions in 180 small herbivorous mammals. Vet Rec. 2008;162:442-449. 62. Jones RJ, Stephenson R, Fountain D, et  al. Urolithiasis in a chinchilla [letter]. Vet Rec. 1995;136:400. 63. Jopp IP, Stengel C, Kraft W. Megaesophagus in a chinchilla (Chinchilla lanigera). A case report. [German] Megaosophagus bei einem Chinchilla (Chinchilla lanigera) Ein Fallbericht. Tierarztl Prax Ausg K Klientiere Heimtiere. 2004;32:96-100. 64. Karanis P, Ey PL. Characterization of axenic isolates of Giardia intestinalis established from humans and animals in ­Germany. Parasitol Res. 1998;84:442-449. 65. Kast A. Leucosis in chinchillas. [German] Leukose beim Chinchilla. Berl Munch Tierarztl Wochenschr. 1962;75:414-415. 66. Keagy HF. Toxoplasma in the chinchilla. J Am Vet Med Assoc. 1949;94:15. 67. Kimpe A, Decostere A, Hermans K, et al. Isolation of Listeria ivanovii from a septicaemic chinchilla (Chinchilla lanigera). Vet Rec. 2004;154:791-792. 68. Kirinus JK, Krewer C, Zeni D, et al. Outbreak of systemic listeriosis in chinchillas. [Portuguese] Surto de listeriose sistêmica em chinchilas. Ciencia Rural. 2010;40:686-689.

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69. Kottwitz J. Stump pyometra in a chinchilla. Exot DVM. 2006;8:24-28. 70. Kottwitz J, Underwood JG. Midfemoral pelvic limb amputation in a chinchilla. Exot DVM. 2005;7:31-34. 71. Kugi G, Nonaka N, Ganzorig S, et al. Taenia crassiceps larvae in a chinchilla (Chinchilla brevicaudata). [Japanese]. J Vet Med. 1999;52:449-452. 72. Langenecker M. Development of species composition and health problems in exotic pets between 1994 and 2003 ­[German]. Retrospektive Untersuchung zur Entwicklung der Artenverteilung und den haufigten Krankheitsbildern bei exotischen Heimtieren im Zeitraum von 1994-2003. Division of Zoo Animals Exotic Pets and Wildlife (Departement für Kleintiere, Abteilung für Zoo-, Heim- und Wildtiere). Zurich: Vetsuisse Faculty, University of Zurich (Vetsuisse-Fakultät Universität Zürich); 2006. 73. Lazzari AM, Vargas ACd, Dutra V, et al. Infectious agents isolated from Chinchilla laniger. [Portuguese] Agentes infec­ciosos isolados de Chinchilla laniger. Ciencia Rural. 2001;31:337-340. 74. Levecke B, Meulemans L, Dalemans T, et  al. Mixed Giardia duodenalis assemblage A, B, C and E infections in pet chinchillas (Chinchilla lanigera) in Flanders (Belgium). Vet Parasitol. 2010:[Epub ahead of print]. 75. Lima L, Montiani-Ferreira F, Tramontin M, et  al. The chinchilla eye: morphologic observations, echobiometric findings and reference values for selected ophthalmic diagnostic tests. Vet Ophthalmol. 2010;13:14-25. 76. Linde A, Summerfield NJ, Johnston M, et al. Echocardiography in the chinchilla. J Vet Intern Med. 2004;18:772-774. 77. Lopes MAF, White NA, Donaldson L, et al. Effects of enteral and intravenous fluid therapy, magnesium sulfate, and sodium sulfate on colonic contents and feces in horses. Am J Vet Res. 2004;65:695-704. 78. Low JC, Donachie W. A review of Listeria monocytogenes and listeriosis. Vet J. 1997;153:9-29. 79. Lusis PI, Soltys MA. Immunization of mice and chinchillas against Pseudomonas aeruginosa. Can J Comp Med. 1971;35:60-66. 80. MacKay KA, Kennedy AH, Smith DLT, et al. Listeria monocytogenes infection in chinchillas. Annual Report of the Ontario Veterinary College. 1949:137-145. 81. Marlow C. Diabetes in a chinchilla [letter]. Vet Rec. 1995; 136:595-596. 82. Marlow CHB. An outbreak of intussusception in a herd of chinchillas. J S Afr Vet Med Assoc. 1963;34:637-642. 83. Mathieu X, Duran JC, Rivas M. Normal bacterial flora of the wild Chinchilla lanigera Silvestre. [Spanish] Estudio de la flora bacteriana normal de Chinchilla lanigera Silvestre. Rev Latinoam Microbiol. 1982;24:77-82. 84. Matthes S. Vaccination of rabbits and fur animals. [German] Schutzimpfungen bei Kaninchen und Pelztieren. Tierarztl Prax. 1985;13:107-112. 85. Mauler D, Lubke-Becker A, Eule C. Ocular findings in healthy chinchillidae. Abstracts, Annu Meet Europ Col Vet Ophthal/ Europ Soc Vet Ophthal 2009 (Abstract 31). Vet Ophthalmol. 2009;12:387. 86. McGreevy PD, Carn VM. Intestinal torsion in a chinchilla [letter]. Vet Rec. 1988;122:287. 87. Meingassner JG, Burtscher H. Double infection of the brain with Frenkelia species and Toxoplasma gondii in Chinchilla laniger. [German] Doppelinfektion des Gehirns mit Frenkelia species und Toxoplasma gondii bei Chinchilla laniger. Vet Pathol. 1977;14:146-153. 88. Menchaca ES, Martin AM, Moras EV, et al. Infectious diseases of the chinchilla. III. Proteus mirabilis and Proteus vulgaris infection. [Spanish] Enfermedades infecciosas de la chinchilla (Chinchilla lanigera). III. “Proteus mirabilis y Proteus vulgaris”. Gaceta Veterinaria. 1978;40:651-656.

324

SECTION III  Guinea Pigs and Chinchillas

89. Menchaca ES, Moras EV, Martin AM, et  al. Infectious diseases of the chinchilla. IV. Pseudomonas aeruginosa. ­[Spanish] Enfermedades infecciosas de la chinchilla (Chinchilla laniger). IV. “Pseudomona aeruginosa”. Gaceta Veterinaria. 1980;42:96-102. 90. Mieth H, Leitner I, Meingassner JG. The efficacy of orally applied terbinafine, itraconazole and fluconazole in models of experimental trichophytoses. J Med Vet Mycol. 1994;32:181-188. 91. Miller LG, Finegold SM. Normal bacterial flora of the chinchilla [abstract]. Bacteriological Proceeding. Ann Meet Am Soc Microbiol. 1967;67:66. 92. Mills R, Gilsdorf J. Middle-ear effusions following acute otitis media in the chinchilla animal model. J Laryngol Otol. 1986;100:255-261. 93. Misirlioglu D, Cetin C, Kahraman MM, et  al. Salmonella infection in a chinchilla farm. [Turkish] Bir cincila ciftliginde salmonella enfeksiyonu. Turk J Vet Anim Sci. 2002;26:151-155. 94. Misirlioglu D, Ozmen O, Cangul IT, et  al. A case of rectum prolapsus and intestinal invagination in a male chinchilla. J Turk Vet Surg. 2000;6:51-53. 95. Moore RW, Greenlee HH. Enterotoxaemia in chinchillas. Lab Anim. 1975;9:153-154. 96. Morgan RV, Pearce LK, Moore FM, et al. Demographic data and treatment of small companion animals with lead poisoning: 347 cases (1977-1986). J Am Vet Med Assoc. 1991;199:98-102. 97. Mountain A. Salmonella arizona in a chinchilla [letter]. Vet Rec. 1989;125:25. 98. Muller M, Haas H, Vogel A, et  al. Mass outbreak of Hymenolepis nana in chinchillas. [German] Massenbefall mit dem Zwergbandwurm Hymenolepis nana beim Chinchilla. Tierarztl Umsch. 2010;65:17-20. 99. Naglic T, Seol B, Bedekovic M, et al. Outbreak of Salmonella enteritidis and isolation of Salmonella sofia in chinchillas (Chinchilla laniger). Vet Rec. 2003;152:719-720. 100. Neville RWJ, Weir BJ, Lazarus NR. Hystricomorph insulins. Symposia of the Zoological Society of London. 1974;34:417-433. 101. Newberne PM, Seibold HR. Malignant lymphoma in a chinchilla. Vet Med. 1953;48:428-429. 102. Nobel TA, Neumann F. Carcinoma of the liver in a nutria (Myocaster coypus) and a chinchilla (Chinchilla laniger). Refuah Veterinarith. 1963;20:161-162. 103. Novak S, Ruttkay D, Solar I. Results of screening for bacterial diseases on large-scale chinchilla (Chinchilla laniger) farms. [Slovakian] Vysledky depistaze bakterialnych ochoreni vo vel’kochovoch cincily vlnatej (Chinchilla laniger). Slov Vet Cas. 1994;19:19-21. 104. Nowakowska M, Matras J, Bartoszcze M, et  al. Clostridium perfringens enterotoxaemia in chinchillas. [Polish] Zatrucie pokarmowe szynszyli wywolane przez enterotoksyne Clostridium perfringens. Medycyna Weterynaryjna. 1991;47:156-157. 105. Olsen OW. Natural infection of chinchillas with the mouse tapeworm, Hymenolepis nana var. fraterna. Vet Med. 1950; 45:440-442. 106. Olson PD, Yoder K, Fajardo LF, et  al. Lethal invasive cestodiasis in immunosuppressed patients. J Infect Dis. 2003; 187:1962-1966. 107. Opazo JC, Soto-Gamboa M, Bozinovic F. Blood glucose concentration in caviomorph rodents. Comp Biochem Physiol A Mol Integr Physiol. 2004;137:57-64. 108. Osborne CA, Albasan H, Lulich JP, et al. Quantitative analysis of 4468 uroliths retrieved from farm animals, exotic species, and wildlife submitted to the Minnesota Urolith Center: 1981 to 2007. Vet Clin North Am Small Anim Pract. 2009;39:65-78. 109. Owens DR, Menges RW, Sprouse RF, et al. Naturally occurring histoplasmosis in the chinchilla (Chinchilla laniger). J Clin Microbiol. 1975;1:486-488.

110. Pantchev N, Globokar-Vrhovec M, Beck W. Endoparasites from indoor kept small mammals and hedgehogs. Laboratory evaluation of fecal, serological, and urinary samples (2002-2004). [German] Endoparasitosen bei Kleinsaugern aus privater Haltung und Igeln Labordiagnostische Befunde der koprologischen, serologischen und Urinuntersuchung (2002-2004). Tierarztl Prax Ausg K Klientiere Heimtiere. 2005;33:296-306. 111. Peiffer RL, Johnson PT. Clinical ocular findings in a colony of chinchillas (Chinchilla laniger). Lab Anim. 1980;14:331-335. 112. Pereyra MLG, Carvalho EC, Tissera JL, et  al. An outbreak of acute aflatoxicosis on a chinchilla (Chinchilla lanigera) farm in Argentina. J Vet Diagn Invest. 2008;20:853-856. 113. Prior JE. Caesarian section in the chinchilla. Vet Rec. 1986; 119:408. 114. Rakich PM, Dubey JP, Contarino JK. Acute hepatic sarcocystosis in a chinchilla. J Vet Diagn Invest. 1992;4:484-486. 115. Rams TE, Slots J. Local delivery of antimicrobial agents in the periodontal pocket. Periodontol 2000. 1996;10:139-159. 116. Rossoff IS. Handbook of veterinary drugs and chemicals: a compendium for research and clinical use. 2nd ed. Taylorville, Pharmatox Publishing Company; 1994. 117. Rubin J, Walker RD, Blickenstaff K, et  al. Antimicrobial resistance and genetic characterization of fluoroquinolone resistance of Pseudomonas aeruginosa isolated from canine infections. Vet Microbiol. 2008;131:164-172. 118. Saitou K, Furuhata K, Kawakami Y, et  al. Isolation of Pseudomonas aeruginosa from cockroaches captured in hospitals in Japan, and their antibiotic susceptibility. Biocontrol Sci. 2009;14:155-159. 119. Sanford SE. Cerebrospinal nematodiasis caused by Baylisascaris procyonis in chinchillas. J Vet Diagn Invest. 1991;3:77-79. 120. Schantz PM. Tapeworms (cestodiasis). Gastroenterol Clin North Am. 1996;25:637-653. 121. Schönball U. Case report: Giardiasis in a chinchilla—possible source of infection for human being? [German] Fallbericht: Gardiaeineninfektion bei einem Chinchilla—mögliche Infektionsquelle für der Menschen? Kleintierpraxis. 1992; 37:785-788. 122. Shelton GC. Giardiasis in the chinchilla. II. Incidence of the disease and results of experimental infections. Am J Vet Res. 1954;15:75-78. 123. Simova-Curd S, Nitzl D, Pospischil A, et al. Lumbar osteosarcoma in a chinchilla (Chinchilla laniger). J Small Anim Pract. 2008;49:483-485. 124. Smith JL, Campbell-Ward M, Else RW, et al. Undifferentiated carcinoma of the salivary gland in a chinchilla (Chinchilla lanigera). J Vet Diagn Invest. 2010;22:152-155. 125. Spence S, Skae K. Urolithiasis in a chinchilla [letter]. Vet Rec. 1995;136:524. 126. Staebler S, Steinmetz H, Keller S, et  al. First description of natural Echinococcus multilocularis infections in chinchilla (Chinchilla laniger) and Prevost’s squirrel (Callosciurus prevostii borneoensis). Parasitol Res. 2007;101:1725-1727. 127. Stampa S, Hobson NK. Control of some internal parasites of chinchillas. J Am Vet Med Assoc. 1966;149:929-932. 128. Stoebe W. Rectal prolapse in chinchilla. [German] Zur Behandlung des Darmvorfalls bei der Chinchilla. Tierarztl Umsch. 1965;20:79. 129. Strake GJ, Davis LA, LaRegina M, et al. Chinchillas. In: LaberLaird K, Swindle MM, Flecknell P, eds. Handbook of rodent and rabbit medicine. Tarrytown: Elsevier; 1996:151-171. 130. Thompson L. Amputation of hindlimbs in chinchillas as a salvage procedure. Proceedings. British Vet Zoological Soc Spring Meet: Non-infectious Diseases of Mammals. 2003. Dublin University. 131. Tisljar M, Janic D, Grabarevic Z, et  al. Stress-induced Cushing’s syndrome in fur-chewing chinchillas. Acta Vet Hung. 2002;50:133-142.

CHAPTER 24  Disease Problems of Chinchillas 132. Tvedten HW, Langham RF. Trophoblastic emboli in a chinchilla. J Am Vet Med Assoc. 1974;165:828-829. 133. Tyrrell KL, Citron DM, Jenkins JR, et al. Periodontal bacteria in rabbit mandibular and maxillary abscesses. J Clin Microbiol. 2002;40:1044-1047. 134. Vanjonack WJ, Johnson HD. Relationship of thyroid and adrenal function to “fur-chewing” in the chinchilla. Comp Biochem Physiol A. 1973;45:115-120. 135. Wagner F, Fehr M. Eye diseases of chinchilla (Chinchilla lanigera): Anatomical and physiological characteristics and disorders. [German] Augenerkrankungen beim Chinchilla (Chinchilla lanigera)—anatomische, physiologische Besonderheiten, Erkrankungen. Kleintierpraxis. 2008;53:309-318. 136. Ward ML, Morrison LR, Else RW, et  al. Endometritis in the chinchilla: 3 Cases (2003-2006). In: Proceedings. British Small Anim Vet Congress Belfast. 2007. 137. Weir BJ. Aspects of reproduction in chinchillas. J Reprod Fertil. 1966;12:410-411. 138. Wideman WL. Pseudomonas aeruginosa otitis media and interna in a chinchilla ranch. Can Vet J. 2006;47:799-800.

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139. Wilkerson MJ, Melendy A, Stauber E. An outbreak of listeriosis in a breeding colony of chinchillas. J Vet Diagn Invest. 1997;9:320-323. 140. Wohlsein P, Thiele A, Fehr M, et al. Spontaneous human herpes virus type 1 infection in a chinchilla (Chinchilla lanigera f. dom.). Acta Neuropathol. 2002;104:674-678. 141. Wuthe HH, Aleksic S. Yersinia enterocolitica serovar 1,2a,3 biovar-3 in chinchillas. Zentralbl Bakteriol. 1992;277:403-405. 142. Yamagishi S, Watanabe Y, Tomura H, et  al. Septic infection of a companion chinchilla with Salmonella enteritidis. [Japanese]. Nippon Juishikai Zasshi. 1997;50:345-348. 143. Yamini B, Raju NR. Gastroenteritis associated with a Cryptosporidium sp in a chinchilla. J Am Vet Med Assoc. 1986; 189:1158-1159. 144. Zetner K, Rothmueller G. Treatment of periodontal pockets with doxycycline in beagles. Vet Ther. 2002;3:441-452. 145. Zierdt CH, Detlefson C, Muller J, et  al. Cyniclomyces guttulatus (Saccharomycopsis guttulata)—culture, ultrastructure and physiology. Antonie Van Leeuwenhoek. 1988;54:357-366.

CHAPTER

25

Soft Tissue Surgery

R. Avery Bennett, DVM, MS, Diplomate ACVS

Guinea Pigs, Chinchillas, and Degus Ovariectomy Ovariohysterectomy Pyometra Uterine Torsion Dystocia Uterine Prolapse Uterine Tumors Mammary Gland Neoplasia Orchidectomy Penile Prolapse Gastric Trichobezoars Urolithiasis Cutaneous Dermal Masses Cervical Lymphadenitis Thoracotomy Miscellaneous Procedures

Both guinea pigs and chinchillas are hystricomorph members of the order Rodentia and have similar biology, anatomy, physiology, disease susceptibility, and surgical conditions. Degus are also hystricomorph rodents. The natural history of rodents has a significant impact on their ability to survive surgical procedures. Rodents are shy and fearful prey species. With fear, pain, and stress, they lose the will to live and may die for no apparent reason. They may appear to recover from anesthesia and surgery, only to die at home 2 or 3 days later. This seems to be especially true of hystricomorph rodents. It appears that the more human contact the pet is used to receiving, the better it bears the stresses of illness and surgery; those that are frequently held, played with, and coddled are more likely to survive than those that spend all their time in a cage. It is important to bear this fact in mind in discussing options and prognosis with owners. Pet rodents that are stressed may stop eating to the point of starvation. Hystricomorph rodents ferment cellulose in the cecum; if they become anorectic, life-threatening complications 326

can occur. Some patients struggle violently, injuring themselves. Some become so frightened that their levels of circulating catecholamines become very high, which can negatively affect anesthesia, and some die acutely from these high levels of catecholamines. This concern underscores the need for appropriate pre- and postoperative analgesic, antianxiety, and tranquilizing agents. If an animal seems stressed by the hospital environment and surgery is not urgent, it may be best to hospitalize it for a day or more, allowing it to adjust to the new environment before surgery. In some cases it may be best to perform multiple short procedures rather than several procedures at once, which would require a long period of anesthesia. Because these herbivorous rodents eat frequently, a short fast of 1 to 2 hours is generally recommended, only to allow the animal to clear its mouth of food material. Herbivores are physiologically unable to vomit, so the risk of aspiration pneumonia is negligible. Because normal gastrointestinal function is vital to recovery, a long fast is not recommended. It has been shown that small mammals with a negative energy balance are at greater risk for postoperative complications.19 Parenteral fluid administration is recommended for most procedures, as it is in other animals. Vascular access can be difficult to obtain in these small patients; however, obtaining vascular access through an intravenous catheter or an intraosseous cannula allows for the administration of fluids during anesthesia and also provides a means whereby emergency drugs can be administered if necessary. If it is possible to maintain vascular access postoperatively, continue administering fluids at a maintenance level until the patient is eating and drinking normally. The reported total blood volume of small mammals is 57 mL/kg body weight.15,19 With loss of 15% to 20% of the total blood volume, most mammals experience hypovolemic shock and release high levels of catecholamines. Life-threatening consequences usually occur with loss of 20% to 30% of the total blood volume.15,19 This would be equivalent to only 4.5 to 6.8 mL of blood in a 400-g guinea pig. Crystalloid, colloid, whole blood from a conspecific, or a blood substitute can be used in patients experiencing serious blood loss. Guinea pigs, chinchillas, and degus seem to be less likely to bother surgical incisions than are other species of rodents. In selecting suture material, keep in mind the propensity of these Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

CHAPTER 25  Soft Tissue Surgery animals to develop a caseous, suppurative response to foreign materials such as sutures. Catgut is degraded by proteolysis and should not be used in rodents because of its reactive nature. Absorbable materials degraded by hydrolysis rather than proteolysis are recommended (e.g., polyglactin 910, polyglycolic acid, polydioxanone, poliglecaprone, glycomer 631, braided lactomer 9-1). Soft, absorbable, braided materials (e.g., poly­ glycolic acid, polyglactin 910, braided lactomer 9-1) are rapidly absorbed and are less irritating to subcutaneous tissues than stiffer materials such as monofilament absorbable sutures (e.g., polydioxanone, poliglecaprone, glycomer 631). Some surgeons recommend stainless steel for cutaneous sutures; however, this material is very stiff and the cut ends often cause irritation, actually stimulating rather than preventing self-mutilation. Many rodents chew out even steel skin sutures. Skin closure is best accomplished with an intradermal or subcuticular technique. This can be time-consuming if the surgeon is not adept. Skin staples are quickly applied, and most rodents will not bother them because there are no ends to poke and irritate the adjacent skin. Cyanoacrylate tissue adhesive can be used to close small incisions; however, excessive amounts of the adhesive on the skin surface will often attract the attention of the rodent and it will try to groom it off, potentially resulting in dehiscence. If the patient demonstrates a tendency toward self-mutilation, midazolam (0.5-2 mg/kg SC, IM q4-6h) may help modulate the behavior, allowing appropriate recovery and healing. This treatment may be needed for only a day or two until the patient becomes accustomed to having a surgical wound. Most patients that have proper postoperative analgesia do not bother their surgical wound initially, though they may begin to pay attention to it during later stages of healing if it becomes pruritic. Small suture size and use of a small-gauge needle are also important. Suture sizes 4-0 to 7-0 are most commonly used in these species of pet rodents. Hemostatic clips are valuable for controlling hemorrhage, especially because of the small size of these patients and because the vessels are often in locations that are difficult to access. Guinea pig skin is relatively thicker than that of chinchillas. Chinchilla skin is fragile and can easily be damaged during clipping for aseptic surgery, the fur is also very fine and epilates easily. Use a number 50 clipper blade (Oster Professional Products, McMinnville, TN) because the teeth are closer together than those of a No. 40 blade, minimizing the risk of the skin being caught between the teeth and getting nicked. Also, to help minimize the risk of tearing, flatten the skin in front of the clipper blade, move slowly over the skin, and clean and lubricate the clipper blades frequently. Tearing of the skin is not as much of a problem in guinea pigs or degus. For skin preparation, a standard surgical scrub is recommended, alternating with warm sterile saline-soaked gauze (instead of alcohol) to minimize evaporative cooling, because these species are prone to hypothermia. The soaked gauze can be warmed in a microwave before use, being careful not to overheat it. Intraoperatively, monitor the patient’s body temperature closely and provide supplemental heat with a circulating warmwater blanket, thermal pad (Thermally Controlled Surgery Pad, RICA Surgical Products, Schiller Park, IL; Hot Dog Patient Warming, Augustine Biomedical Design, Eden Prairie, MN), a forced warm-air blanket, a radiant heat lamp, or some combination of these.

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GUINEA PIGS, CHINCHILLAS, AND DEGUS OVARIECTOMY Cystic ovaries are common in guinea pigs, with an incidence reported to be as high as 100%.33,34,37 In a study of guinea pigs with reproductive problems, 76% had cystic ovaries.21 In another study of 43 guinea pigs from various owners or breeders, 53% had evidence of cystic ovaries on ultrasound examination and 36% had evidence of bilateral cystic ovaries.28 Of the guinea pigs above 2 years of age, 93% had cystic ovaries, 62% of which were bilateral. There was a direct relationship between age and the size of the ovarian cysts and there was no difference in incidence between breeding and nonbreeding sows.28 There was no difference in reproductive success in sows with or without cysts younger than 15 months of age; however, guinea pigs greater than 15 months of age with cysts had a marked decrease in reproductive success.21 While cystic ovaries are common, most sows show little evidence of clinical disease. In juvenile sows, they are small (5 μm), but they enlarge as the animal matures, reaching up to 7 cm in size.21,33 If the cysts are 2 cm, they are usually single cysts.34 The largest are often multilocular. Histologically, ovarian cysts in guinea pigs are similar to those of humans and cats, being cysts of the rete ovarii.21,33,34,37 Rete cells are from the mesonephros; they migrate into the fetal gonads and differentiate into rete testis in males and rete ovarii in females. They form a tubular, blind-ended vestigial structure within the ovary. The function of the rete cells is suspected to be phagocytosis of degenerating oocysts and production of a meiosis-inducing substance. These cells do not produce hormones. The pathogenesis of cyst formation is unknown but is suspected to be the result of a defect in ion pumps, so electrolytes are transported into the tubular structure but not out. Fluid is pulled in as the ion concentration increases, resulting in the formation of cysts. Since these cysts do not produce hormones, patients are usually asymptomatic. Clinical signs include abdominal distention as the cysts enlarge; they may cause anorexia and depression because of the effects of the space-occupying mass. Many sows with ovarian cysts also have uterine disease and may present with a hemorrhagic vaginal discharge or reported hematuria that is actually a result of uterine hemorrhage.9,21 Some guinea pigs have a nonpruritic symmetric alopecia typical of an endocrine alopecia; they become aggressive and begin mounting cage mates9; however, it is considered normal for female guinea pigs to mount cage mates during estrus. These signs are indicative of hormone increases that are difficult to explain, since the cysts do not produce hormones. The diagnosis is confirmed with ultrasound.3 Radiographically, ovarian cysts cannot be differentiated from other abdominal masses because fluid is of the same radiographic density as soft tissues. Ovariohysterectomy is the treatment of choice for guinea pigs with cystic ovaries because uterine disease secondary to these ovarian cysts is common, although a mechanism has not been established. In a study of five guinea pigs with cystic ovaries histologically confirmed to be rete ovarii cysts, all were found to have uterine disease, including endometritis, pyometra, endometrial hyperplasia, and leiomyomas.9,10 If the uterine changes are secondary to the ovarian cysts, ovariectomy at a young age would be expected to prevent both problems.37 Because ovarian rete cysts are so common and become

328

SECTION III  Guinea Pigs and Chinchillas

larger with age, routine ovariectomy or ovariohysterectomy at a young age should be recommended to owners of female guinea pigs. Drainage of ovarian cysts in guinea pigs provides temporary relief, but fluid quickly reforms. Prior to surgery, drain the cysts as well as possible, using percutaneous centesis to decrease their size. Be aware that there are often adhesions between the diseased ovaries and uterus to the body wall and other viscera. In guinea pigs with alopecia, hair regrowth is generally complete within 3 months after surgery.3,9 Rete ovarii cysts do not produce hormones and should not be affected by the administration of gonadotropin-releasing hormone or its analogs, such as human chorionic gonadotropin.5,13,37 Those hormones are effective in treating follicular cysts, causing them to luteinize. Reports of cysts other than cystic rete ovarii are rare, but a granulosa cell tumor was successfully managed by ovariohysterectomy in a guinea pig.5 Still, administration of 100 IU (1,000 USP units) of human chorionic gonadotropin administered intramuscularly in two doses given 2 weeks apart has been reported to resolve the clinical signs temporarily.5 Gonadotropin-releasing hormone has also been reported to be “very effective” in treating guinea pigs with ovarian cysts.25 Ovarian teratomas have also been reported to occur commonly in sows older than 3 years of age.8,11,13 Some teratomas may be as large as 10 cm in diameter.40 They are usually unilateral and rarely metastasize. Affected sows present for depression, weakness, or collapse due to spontaneous intra-abdominal hemorrhage from the tumor. Acute death from blood loss can occur. Ovariectomy or ovariohysterectomy is the treatment of choice for sows with these tumors. Ovariectomy of young rodents may decrease the incidence of mammary neoplasia; however, this has not been documented in hystricomorph rodents. Ovariectomy can be done through a ventral   midline ap‑­ proach or a dorsolateral approach.35 The ventral midline approach is as described for other rodents (see Chapter 28). To perform an ovariectomy using the dorsolateral approach, make a 1- to 2-cm incision on each side ventral to the erector spinae muscle and about 1 cm caudal to the last rib. Bluntly penetrate the muscle with a hemostat and enlarge the opening to about 1 cm. With pressure on the abdomen to push the ovary to the incision, reach in with forceps and grasp the ovary. Exteriorize the ovary and use a hemostatic clip or ligature to control any hemorrhage from the ovarian vessels that might occur. Also be sure to remove the entire oviduct surrounding the ovary to prevent cysts from forming. Close the opening in the body wall with 1 or 2 sutures of monofilament absorbable material and appose the skin edges with tissue adhesive or an intradermal suture. No advantage has been documented for performing ovariohysterectomy over ovariectomy unless there is concurrent uterine disease and, in fact, most complications associated with ovariohysterectomy result from removing the uterus.42 The advantage to performing ovariectomy is that the incisions are small and dorsal and the delicate gastrointestinal tract is not disturbed, resulting in less morbidity and a more rapid recovery.

OVARIOHYSTERECTOMY Indications for ovariohysterectomy in hystricomorph rodents include dystocia, uterine prolapse, pyometra, and uterine and/ or ovarian masses. The ovaries are located caudolateral to the kidneys and are approximately 8 mm in length and 5 mm in

a

b

c

Fig. 25-1  The ovary (a) in this guinea pig is located at that caudal pole of the kidney. The uterine horns (b) are long and come together to form the uterine body (c).

width (Fig. 25-1).19 The oviduct lies in close proximity to the dorsal aspect of the ovary, encircling it before joining the uterine horn.31 The uterus is bicornuate and the horns join to form a uterine body, which is divided internally by a well-developed intercornual ligament; however, there is a single cervical os. The mesovarium, mesometrium, and broad ligaments are sites of fat storage in guinea pigs and chinchillas, adding to the difficulty of the procedure. The ovarian artery and vein are branches off the renal vessels that split into an ovarian branch supplying the ovary and a uterine branch to the uterus.31 There is a single artery and vein medial to the ovaries and along the uterus to the uterine body. Once the patient has been anesthetized, clip the abdomen and prepare it for aseptic surgery. Place the patient in dorsal recumbency and drape appropriately. Make a 2- to 3-cm incision centered midway between the umbilicus and pubis. There is usually little subcutaneous tissue and the linea alba is broad, making it easy to identify. Immediately dorsal (deep) to the body wall is the thin-walled cecum, and the bladder (also thinwalled) is just caudal to the cecum. It is vital to avoid iatrogenic injury to these structures, especially the cecum. Leakage of cecal contents can cause life-threatening complications. In addition, because the cecal wall is so thin, it is difficult to achieve typhlotomy closure free of leaks. Because of the potential for damage to these organs, use of a spay hook is not recommended. Locate the uterus between the bladder and the colon. Use a blunt instrument or a finger to move the cecum and bladder to the side on which you are standing, allowing visualization of the uterine horn on the opposite side. Grasp the uterus gently with forceps, and exteriorize it. Trace it cranially to locate the ovary on that side. The ovaries are supported by the mesovarium, which that originates in the area of the caudal pole of the kidney. The mesovarium is short, making the ovaries are

CHAPTER 25  Soft Tissue Surgery more difficult to exteriorize than in carnivores. It may be necessary to extend the incision cranially to avoid accidental tearing of the friable, fat-filled ovarian ligament. The broad ligaments also contain a large amount of fat, which can make identification of the ovarian vessels difficult. A single artery and vein run medial to each ovary and uterine horn.14 Identify the vessels supplying the ovary within the mesovarium and, using gentle blunt dissection, create an opening in the mesovarium to allow placement of two hemostatic clips or two ligatures of an absorbable synthetic suture. Transect the suspensory ligament, mesovarium, and vessels distal to the ligatures. Alternatively, these vessels can be sealed and cut with a tissue-sealing device such as a CO2 laser, a Harmonic device, or a LigaSure (see Chapter 28). It is important to remove the entire oviduct encircling the ovary. Remnants of oviduct can develop into cystic masses within the abdomen.19 Repeat the procedure on the contralateral side and bluntly dissect the broad ligament on each side to the level of the uterine body. Strip the broad ligament on each side caudally to the uterine vessels and uterine body. Ligate the vessels with the uterine body unless they appear particularly large, in which case ligate them separately. The uterus may be ligated with an encircling ligature or with a transfixation ligature. It has been recommended that the uterus be ligated cranial to the cervix to prevent spillage of urine into the abdomen when the uterus is transected19; however, this is of little clinical importance. Place the ligature in the body of the uterus in a convenient location. Remove the ovaries and uterus as a unit. Close the abdomen with 4-0 monofilament absorbable suture in the linea alba in a simple continuous pattern, using 5-0 absorbable suture for the subcutaneous or subcuticular closure. If necessary, use tissue adhesive, 4-0 or 5-0 nonabsorbable suture or skin staples to appose the skin.

PYOMETRA Pyometra is infrequently reported in guinea pigs and chinchillas.4,43 Possible pathogens include Bordetella bronchiseptica, Escherichia coli, Corynebacterium pyogenes, Staphylococcus species, and Streptococcus species. Affected animals are usually presented for vaginal discharge and may be lethargic and anorectic. Some guinea pig owners report polydypsia and decreased appetite. Radiographic and abdominal ultrasound examinations are valuable in obtaining a diagnosis and in ruling out pregnancy, dystocia, and abdominal masses. Vaginal cytology, along with culture and sensitivity testing, confirms the tentative diagnosis. Stabilize the patient and perform a complete blood count and plasma biochemical panel before surgery. Vascular access is required because these patients are usually dehydrated and have other metabolic abnormalities. It also allows intravenous antibiotics to be administered. Administer an appropriate antibiotic intravenously after samples for culture have been obtained intraoperatively. Definitive treatment of pyometra is ovariohysterectomy, which is performed as soon as the patient is stable enough to undergo general anesthesia and surgery. Take care not to spill uterine contents into the abdomen during removal of the uterus. Irrigate the abdomen with warm sterile saline solution before routine closure. Continue fluid therapy and nutritional support postoperatively until the patient is eating and drinking normally.

329

UTERINE TORSION Uterine torsion is uncommon in most domestic pets but has been reported in gravid guinea pigs after 30 days of gestation and in gravid chinchillas.43 Signs are the same as those for dystocia, but usually signs of circulatory shock and acute collapse are also present. The mortality rate is high, and the diagnosis is usually made at necropsy. This is an emergency situation. Establish vascular access and obtain a minimum database before surgery. Stabilize the patient metabolically as well as possible and then perform an emergency ovariohysterectomy.

DYSTOCIA Dystocia is relatively common in guinea pigs and chinchillas because of the relatively large size of the fetuses in these animals.29,43 Degus are smaller and less well developed at birth but are still considered precocious.20 Guinea pigs should be bred before they are 6 months of age, because bony fusion of the pubic symphysis occurs between 6 and 9 months of age. If the pubic symphysis fuses before the first litter is delivered, dystocia can result.29 If a guinea pig delivers a litter before bony fusion of the pubic symphysis has occurred, cartilaginous fusion is preserved for life and future litters are possible without dystocia. Female guinea pigs are sexually mature at 28 to 35 days of age. Weaning typically occurs at 14 to 28 days of age.15 In female chinchillas, fusion of the pubic symphysis is normal and does not cause dystocia. Male and female chinchillas reach sexual maturity at 4 to 12 months of age, much later than guinea pigs.29 Chinchillas are seasonally polyestrus and age at puberty is a function of when they were born. Those born in the late summer do not reach maturity until the next fall breeding season.18 Degus generally wean between 4 and 6 weeks of age and reach sexual maturity at 3 to 4 months of age.20 Gestation is approximately 59 to 72 days (usually 63-68 days) in guinea pigs, 111 days in chinchillas, and 87 to 93 days in degus.15,19,20 Average litter size is 2 to 4 in guinea pigs, 1 to 6 in chinchillas, and 6 to 7 in degus (range, 1-10). In guinea pigs, approximately 10 days before parturition, the pubic symphysis begins to spread. Once the gap is 15 mm, parturition should occur within 48 hours15; at parturition, the symphysis is about 22 mm wide. This gap can be palpated externally; this is a sign of impending parturition.29 If the symphysis is open or the sow has had a previous litter without intervention and the sow has been in unproductive labor for longer than 30 to 60 minutes, give 0.5 to 1 U of oxytocin IM. If no young are delivered after 15 minutes, surgical intervention is likely necessary.30 If guinea pigs become pregnant for the first time after 9 months of age, many will still have normal parturition. If the breeding date is known, palpate the pubic symphysis at about day 60 to determine if it has spread. If so, it is likely that the sow will deliver naturally. If the symphysis does not separate by day 65 or if it has not separated and the sow shows signs of parturition, perform a cesarean section. If the breeding date is unknown, the author has not found a way to determine the due date. Educate owners on the signs of parturition and how to palpate for symphyseal separation. Dystocia in guinea pigs and chinchillas can be surgically treated by either cesarean section or ovariohysterectomy of the intact gravid uterus. Cesarean section is performed to obtain viable fetuses or, if the fetuses are not viable, to preserve the reproductive viability of the sow for future breeding. For either

330

SECTION III  Guinea Pigs and Chinchillas

procedure, make a routine ventral midline abdominal incision and exteriorize the gravid uterus. For a cesarean section, isolate the uterus with sponges that have been moistened with saline solution and make a longitudinal incision in the dorsal or ventral uterine body, depending on the position of the uterus after it has been exteriorized. Deliver the neonates to an assistant and close the incision with a simple continuous pattern of 4-0 or 5-0 monofilament absorbable suture material. Irrigate the abdomen with warm saline solution before closing. Ovariohysterectomy of the intact gravid uterus can be performed to retrieve the offspring and prevent future pregnancy. The technique is similar to that used for routine ovariohysterectomy. After ligating and dividing the ovarian pedicles, clamp the uterine vessels and uterine body. Transect and remove the gravid uterus, passing it to an assistant before ligating the uterine stump. The assistant opens the uterus and removes and revives the neonates while the surgeon ligates the uterine vessels and uterine stump. It may be necessary to ligate the uterine vessels separately and transfix the uterus, because these structures are enlarged during pregnancy. Clamping of the uterine vessels occludes the blood and therefore also the oxygen supply to the fetuses. Thus it is important to proceed quickly. However, the viability of the neonates is generally not affected by removing them with the uterus en bloc. If the uterus is exceptionally large or engorged with blood and the sow is anemic, the en bloc technique is not recommended. If ovariohysterectomy is indicated, perform a cesarean section first and allow the uterus to involute before removal so that most of the blood in the uterus returns to the sow’s circulation. Give oxytocin IV, IM, or injected directly into a uterine artery (to deliver the hormone directly to the uterus) to speed involution. Guinea pig and chinchilla young are precocious at birth. Their eyes are open, they ambulate well, and they can eat solid foods; however, they should be allowed to nurse as soon as the sow has recovered from anesthesia. Guinea pigs that are orphaned at less than 1 week of age have a high mortality rate, indicating that they do need to have sow’s milk.15 Degus are also precocious at birth but less developed than guinea pigs and chinchillas, having a sparse hair coat and closed eyes.20

UTERINE PROLAPSE Uterine prolapse is usually associated with parturition. In most cases a tissue mass is protruding from the vulva after delivery of young (Fig. 25-2). Most often, the owner discovers live young at the same time the prolapse is noticed. The sow may be stable at presentation or may be debilitated, depending on how long the uterus has been prolapsed. Stabilize the patient medically before administering anesthesia. Consider epidural anesthesia for patients that are not stable enough to undergo general anesthesia. Clean and assess the prolapsed tissue. The prolapsed uterus is usually contaminated by the substrate. If it appears to be viable, clean and reduce the prolapse. A concentrated sugar solution or 50% dextrose applied topically to the prolapsed tissue usually helps reduce edema and the size of the prolapsed tissue, making it easier to replace. If the reproductive viability of the sow is to be preserved, reduce the prolapsed uterine horn into its proper location within the abdomen. If the horn is not returned to its normal location, it is likely to prolapse again. Use an appropriately sized blunt probang to gently push the horn to approximately

Fig. 25-2  This guinea pig was presented for uterine prolapse the morning after farrowing.

midabdomen. After the horn is reduced, monitor the sow closely for reprolapse. If this occurs, perform an emergency ovariohysterectomy. A purse-string suture is not recommended because the prolapsed uterus might be retained in the vagina and cause urinary obstruction, which can be fatal. In most cases, ovariohysterectomy should be recommended. If the exposed uterine tissue is not viable, ovariohysterectomy must be performed after the patient has been adequately stabilized. Prognosis depends on the stability of the patient at presentation. If the patient is alert and active, the prognosis is fair to good.

UTERINE TUMORS Leiomyoma is the most common reproductive tumor of guinea pigs and is usually associated with rete ovarian cysts.10,13,15

MAMMARY GLAND NEOPLASIA Mammary gland neoplasia is uncommon in older guinea pigs and is even rarer in animals younger than 3 years of age.30 There are no published reports of mammary neoplasia in chinchillas, but of 233 specimens submitted to an exotic animal pathology service, 2 were mammary tumors with 1 being a benign adenoma and the other an adenocarcinoma. A third poorly differentiated sarcoma was reportedly collected from a mammary gland, but no glandular tissue was identified. Two specimens were diagnosed as mastitis (D. Reavill, personal communication). No mammary gland tumors were diagnosed in degus. Both male and female guinea pigs have mammary glands located inguinally as a single pair. Chinchillas have two pairs of mammary glands. Neoplasias of the glands, both benign and malignant, have been reported to occur in guinea pigs of both sexes; about 70% are benign fibroadenomas and 30% are mammary adenocarcinomas.7 Adenocarcinomas are locally invasive but rarely metastasize7,8,13,15; however, pulmonary metastasis has been reported.22 Liposarcoma, adenoma, papillary cystadenoma, and carcinosarcoma have also been reported.40 Because of the possibility of malignancy, do a preoperative biopsy or fine-needle aspirate cytology of the mass. If the mass is malignant, it is important to stage the disease before surgery. Staging includes blood work, urinalysis, thoracic radiographs, abdominal ultrasound, and evaluation of regional lymph nodes. Wider excision is recommended for malignant mammary tumors, underscoring the need for a preoperative diagnosis.

CHAPTER 25  Soft Tissue Surgery The left and right inguinal mammary glands of guinea pigs do not have a common blood or lymphatic supply.26 Information regarding the incidence of bilateral disease or of a neoplasm developing in the second gland after one is excised is lacking; however, such cases appear to be uncommon. The benefit of ovariectomy for reducing the risk of the other gland developing a tumor has not been studied. For benign mammary masses, marginal resection is all that is required, and the skin over the tumor can be preserved. The benefit of mastectomy over lumpectomy has not been studied in these rodents, but mastectomy is recommended. Excise all of the mammary tissue, including the areola on the affected side. Usually there are several blood vessels in the subcutaneous tissue supplying the tumor that may have to be ligated. Bluntly dissect around the mass, ligating or clipping vessels as they are encountered. Locate and ligate the caudal superficial epigastric artery and vein where they enter the tissue to be excised. The excision will create dead space, and drains and bandages are not well tolerated by rodents. It is best to tack the skin and subcutaneous tissues to the body wall to eliminate dead space, thus minimizing the risk of seroma formation. Submit the tissue for histologic evaluation. Mastectomy of the affected gland with 0.5 to 1.0 cm of healthy tissue surrounding the mass, including deep to it, is recommended for malignant mammary tumors. In most patients, deep to the mass, remove the body wall to obtain the needed deep margin. Plan carefully to allow for adequate closure, because tissue is not abundant in this area. There is usually adequate body wall to close primarily with simple continuous or interrupted suture. Locate and remove the inguinal lymph node. Close the subcutaneous tissues, tacking to the body wall to eliminate dead space, and close the skin routinely. Submit the tissue for histologic evaluation and request that the pathologist evaluate the margins of the submitted tissue for evidence of tumor. Also ask that the pathologist to quantitate the distance from the tumor to the cut surface if margins are clean. This is vital to determine if you have achieved local tumor control. If bilateral mammary tumors are present (benign or malignant), two unilateral mastectomies are staged 2 to 4 weeks apart, allowing one side to heal and the skin to stretch before the other gland is removed.

ORCHIDECTOMY In hystricomorphs, orchidectomy is primarily used to control reproduction. Seminal plugs have been reported to be a primary cause of urethral obstruction in guinea pigs and can be treated or prevented by orchidectomy. This is easier than performing ovariohysterectomy and is associated with less morbidity and mortality. It may also be indicated to decrease aggression and for medical reasons (e.g., testicular tumor). If a boar guinea pig is to be housed with a female that has a fused pubic symphysis, orchidectomy is recommended to prevent the sow from getting pregnant, which could result in dystocia. The testicles of most rodents are comparatively large and descend during the first week or two of life.14 The inguinal canals remain open, and a functional cremaster muscle allows the testicles to migrate into and out of the abdominal cavity.16 Hystricomorph rodents do not have a well-developed scrotum; instead, the testicles are located lateral to the penis in the inguinal region on each side (Figs. 25-3 and 25-4). Rodents have a large epididymal fat pad within the vaginal tunic that helps prevent intestinal herniation. The large seminal

331

a c

c

b d

Fig. 25-3  The genitalia of a male chinchilla. The penis (a) is directed caudally and almost touches the anus (b), and the testicles (c) are located lateral to the penis. An incision can be made over the tail of the epididymis (d) for castration.

vesicles and coagulating glands also partially occlude the internal inguinal ring, preventing herniation. Because of the anatomy, inguinal hernias are very rare in these rodents, and visceral herniation after orchidectomy has not been reported; however, herniation is frequently discussed as a major concern in performing an orchidectomy in rodents. This concern seems primarily theoretical; because these anatomic mechanisms prevent visceral herniation, it appears unlikely that the inguinal canal would have to be closed. Intratesticular injection of 2% lidocaine at 1 mg/kg per testis has been recommended for intraoperative analgesia. Once the patient is anesthetized, each testicle is injected with lidocaine to minimize the perception of pain when the testicles are manipulated in the anesthetized patient. It must be noted that this does not provide any postoperative analgesic effect but may lower the percentage of inhalant agent needed to maintain anesthesia during the procedure. Orchidectomy can be performed by a closed or open technique. A closed technique requires the least amount of exposure, and viscera cannot herniate into the scrotum because the tunic is tied off near the external inguinal ring. With the patient in dorsal recumbency, clip the fur around the scrotum, penis, and inner thighs and prepare the area for aseptic surgery. Palpate both testes, being careful not to confuse the body of the penis with a testicle. The penis can feel similar to a testicle under the skin, especially if one testicle is retracted into the abdomen. The testes are wider and rounder than the body of the penis, and the penis, which is located on the midline, cannot be pushed back into the abdomen, as the testicles can. If one testicle is in the abdomen, gentle caudoventral pressure results in its return into the scrotum. For the closed technique, holding one testicle between the thumb and forefinger, make a 1.0- to 1.5-cm incision through the scrotum parallel to and 0.5 to 1.0 cm lateral to the penis on each side near the external inguinal ring. The incision should not penetrate the tunica vaginalis. If the

SECTION III  Guinea Pigs and Chinchillas

332

b

b

a

A

B Fig. 25-4  A, The genitalia of a male guinea pig; a, penis; b, testicles; anus (arrow). B, The penis has been exteriorized.

b

a

Fig. 25-5  The testicle has been exteriorized for a closed castration. a, Testicle within tunic, b, epididymal fat within tunic. Note: inadvertently, small incisions were made in the tunic (arrow).

incision is too close to the penis, the penis can be detached from the prepuce during dissection. In a degu, such preputial damage was successfully treated by suturing the tip of the penis to the cranial edge of the prepuce with 4-0 polydioxanone suture.32 Two years later, the penis was still adhered to tip of the prepuce. Grasp the tunic and remove the testicle from the scrotum with the tunic intact (Fig. 25-5). The tunic is tightly adhered to subcutaneous tissues. Carefully and gently dissect the tunic from its attachments circumferentially. The tunic is also tightly adhered to the end of the scrotum by the ligament of the tail of the epididymis. Break down this ligament to allow the testicle to be exteriorized. Once the testicle is removed from the scrotum, apply caudal traction to it and strip the fascial attachments, using a dry gauze sponge, until the narrow portion of the cord is exposed. Remember, you need to remove only the testicle; pulling the testicle far out is of no added benefit and may damage the epididymal fat and ipsilateral ureter. Be careful to avoid tearing the vaginal tunic during this dissection. Once the testicle has

been exteriorized adequately, gently push the epididymal fat into the inguinal canal and ligate the cord, using a two-clamp technique. Crushing the tissue with the clamps before placing the ligatures is helpful with the closed technique because the tissue cord is thicker, since the vaginal tunic is incorporated. This ensures a more secure ligature, but it can also cause the tunic to tear, which will defeat the purpose of doing a closed technique (i.e., blocking the inguinal canal). The goal is only to remove the testicle; the epididymal fat must be preserved within the inguinal canal to help prevent hernias. With the closed technique, herniation into the scrotum is not possible unless the ligature fails or the tunic tears. In performing an open castration, make the incisions as described above. Extend the incision through the subcutaneous tissues and the vaginal tunic, exposing the spermatic cord. The testicles are easily exteriorized because they are not attached to the internal (parietal) surface of the vaginal tunic; however, they are attached to the tunic by the ligament of the tail of the epididymis (Fig. 25-6). Traction on the testicles will invaginate the scrotal skin toward the incision. Break down the ligament of the tail of the epididymis, allowing the skin and the vaginal tunic to return to their normal location and leaving the testicle outside the body. With the open technique, the testicle is easily exteriorized and the cord to be ligated is of a smaller diameter than with the closed technique. Be careful to preserve the epididymal fat within the inguinal canal. Double ligate the spermatic cord distal to the epididymal fat and transect it distal to the ligatures. If there is concern about visceral herniation because of trauma or removal of the epididymal fat, identify the external inguinal ring and place a single interrupted suture across it, being careful not to damage or occlude the external pudendal artery and vein, which pass through the canal. The incision in the tunic can be closed or left open. Capello describes a technique for castrating a degu that combines the benefits of an open and a closed castration technique; it is also applicable to chinchillas and guinea pigs because they have similar anatomies.6 Make the incisions as described above. Use a mosquito hemostat to bluntly dissect under and around the spermatic cord within the vaginal tunic without damaging

CHAPTER 25  Soft Tissue Surgery

333

c a b

c a

b

A

B Fig. 25-6  A, Open castration of a chinchilla. B, Open castration of a guinea pig. (a) Testis, (b) epididymis, (c) epididymal fat, vaginal tunic (arrow).

the tunic. Pass a suture ligature around the cord but do not tie it. Make an incision in the tunic and perform an open castration as described above, making sure to keep the epididymal fat cranial to the preplaced ligature. After the testicle is removed, tie the ligature around the tunic as close to the external inguinal ring as possible, thus effectively closing off the inguinal canal. Chinchillas have a particularly large tail of the epididymis. Nelson describes an approach for castration through the skin on the tail of the epididymis.27 Make a small incision over the ventral aspect of the tail of the epididymis, being careful not to cut too deeply into the vaginal tunic. Grasp the tip of the tail of the epididymis with gauze and apply traction while pushing the scrotal skin cranially. The testicle can then be completely exteriorized for either an open or a closed technique. Primary closure of the skin with intradermal suture is recommended, but tissue adhesive and second-intention healing have also been used successfully. Guinea pigs and chinchillas seem particularly prone to the development of scrotal abscesses after orchidectomy. If this occurs, it is more likely that visceral herniation will ensue, because the infection causes tissue necrosis in the area of the inguinal canal, widening the opening and allowing intestine to herniate. It is currently unknown why infection occurs more commonly in hystricomorph rodents. One theory is that the location of the incisions and the way the animal stands contaminate the incisions with feces from the substrate. Owners should be warned of this potential complication before surgery is performed and told that the substrate must be kept clean. Advise owners to use clean paper bedding and change it twice daily for 10 days after surgery. Strictly adhere to aseptic technique. Gentle tissue handling is critical, because traumatic manipulations can cause tissue necrosis and predispose to infection. Application of a layer of cyanoacrylate tissue adhesive over the incision may help to prevent bacterial invasion into the incision. Prophylactic antibiotic therapy should be considered as well.

PENILE PROLAPSE Penile prolapse seems to occur in hystricomorph rodents more commonly than in other small mammals. It is often reported anecdotally after orchidectomy and may be related to nerve trauma during the procedure. Prolapse of the penis may also occur but unassociated with any specific cause.

The penis is protected from the environment within the prepuce. When the penis is prolapsed, it is subject to trauma and contamination. With chronic exposure, the mucosa of the penis becomes thicker and more able to resist such trauma. Most owners are concerned by seeing their pet’s penis outside of its sheath. Powers reported preputial damage and lateral deviation of the penis into the subcutaneous tissues as a complication of orchidectomy in a degu.32 An attempt to maintain the penis within the prepuce by suturing the base of the penis to the base of the prepuce was not successful. Subsequently, the tip of the penis was sutured to the edge of the preputial orifice with four interrupted sutures of polydioxanone, being careful to avoid the urethra. The tissue was not scarified. This technique is simple and easy to perform and maintains the penis in its normal position within the prepuce. In that patient, the penis was still adhered to the prepuce 2 years after surgery. The author recommends this procedure for hystricomorph rodents with penile prolapse.

GASTRIC TRICHOBEZOARS Gastric trichobezoars causing clinical illness have been reported in long-haired Peruvian guinea pigs.2,24,39 The condition is not analogous to what was once called trichobezoars in rabbits. In Peruvian guinea pigs, the hairball is quite large for the animal’s body size (4-5 cm), composed of firmly compacted material consisting primarily of hair. The dense, hard nature of such a bezoar makes it unlikely that it would be broken down and passed simply by medical management; therefore surgery is indicated. In a study looking at the influence of hay on alopecia in guinea pigs, it was determined that when hay was withheld from the diet, a loss of hair density was seen within 4 weeks. This did not resolve when a high-fiber pelleted diet was given.12 The alopecia was observed to result from cage mates eating each other’s hair. Therefore a proposed etiology for trichobezoars in Peruvian guinea pigs is inadequate hay in the diet, resulting in overgrooming of the very long hair combined with inadequate exercise and also possible stressors resulting in the formation of a hard trichobezoar.39 Clinical signs of a gastric trichobezoar can be vague and insidious or acute and severe, depending on whether the bezoar is obstructing gastric outflow. Signs include anorexia,

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SECTION III  Guinea Pigs and Chinchillas

depression, weight loss, and decreased production of feces. On physical examination, a large firm mass is palpated in the left cranial abdomen, which does not indent with firm pressure. Radiographs may confirm the presence of a gastric foreign body, but only if there is gas in the stomach to serve as contrast. Orally administered barium confirmed the diagnosis in one patient and ultrasound in another.2,39 The advantage of ultrasound is that other organs can also be evaluated, including the liver, for evidence of hepatic lipidosis, which may complicate the condition and postoperative management. Gastrotomy is recommended as soon as the patient is stable enough for surgery. Pre- and postoperative fluid therapy is vital for a successful outcome. Perform a ventral midline celiotomy from the xiphoid to the pubis to allow the large object to be easily removed. Isolate the stomach with gauze moistened in saline solution and make an incision in a relatively avascular area of the stomach midway between the lesser and greater curvatures. Remove the trichobezoar and irrigate the stomach, being careful to minimize abdominal contamination. Close the stomach in two layers; a simple continuous pattern, over sewn with an inverting pattern such as a Cushing’s with a monofilament absorbable material on an atraumatic needle. Irrigate the abdomen with warm sterile saline solution and close the celiotomy routinely. Postoperative care consists of continuing fluid support, analgesia, nutritional support with a syringe-fed diet, prokinetic medications, and antibiotics if indicated. Keep the animal in the hospital until it is eating and drinking and urinating and defecating normally. Free access to fresh hay, regular grooming of the long hair and potentially cutting it, plenty of exercise, and minimizing stress are considered important to reduce the risk of recurrence. Some have advocated petrolatum-based gastrointestinal lubricants, but their efficacy for preventing trichobezoars and effect on the cecum is unknown.39

UROLITHIASIS Urolithiasis is relatively common in guinea pigs and the primary differential diagnosis for hematuria.14,16 Clinical signs include hematuria, stranguria, dysuria, incontinence, anorexia, and pain manifesting as an animal acting sick.16 Urinary tract obstruction because of calculi can occur at the renal pelvis, ureter, or urethra. In male guinea pigs, urinary tract obstruction occurs more commonly secondary to plugs of sperm and seminal fluid. As a general rule, uroliths should be removed. They cause irritation, which often results in hemorrhage and consequent anemia. Uroliths also cause serious pain and can serve as a nidus for bacterial urinary tract infections. In a patient with stones but without obstruction, the stones may to migrate to a point where they do cause obstruction. Undetected urinary tract obstruction often results in death. Calcium oxalate is the most common type of stone found in guinea pigs, and the etiopathogensis is poorly understood.17 Medical dissolution of stones has not been documented in guinea pigs and recurrence after surgical removal is common (see Chapter 23). Renal mineralization often cannot be distinguished from nephrolithiasis on plain radiographs. Ultrasound will determine the presence of stones rather than mineralization of the renal pelvis. Ureteral calculi can lodge anywhere from the renal pelvis to the entrance to the bladder, but they are most commonly located at the caudal bend of a ureter as it enters the

bladder. The urethra of female guinea pigs is quite large, and large stones often obstruct the urethral papilla because they cannot pass through it (Fig. 25-7). Stones within the bladder do not usually cause signs of obstruction but should be removed because they cause pain and cystitis, predispose to bacterial urinary tract infections, and may migrate into the urethra, causing obstruction. If a nephrolith or ureterolith is causing obstruction, the function of the kidney may be compromised. If the kidney is not functional, the best option is to perform a nephrectomy; however, that will leave the patient with only one functional kidney. It is best to determine whether there is any functional capacity in the kidney prior to surgery. The scintigraphic glomerular filtration rate is very accurate for determining whether a kidney has any remaining function, but this modality is not readily available. An excretory urogram can provide valuable information; however, its accuracy will be compromised if the patient is azotemic. Any blushing of the kidney with contrast indicates that there is still some function remaining and it is worth trying to save it. Since recurrence of urolithiasis is common in guinea pigs, it is best to save the kidney if possible. If the kidney on the affected side is not functional, infected, or severely hydronephrotic, it may be necessary to perform a nephrectomy. In other species, end-stage kidneys have been associated with systemic hypertension.38 Ultrasound is a vital part of working up any patient with urolithiasis. Ultrasound will confirm the location of the stones within the urinary system and determine whether they are causing partial or complete obstruction. Some mineral-dense objects that appear to be within the urinary tract are actually outside it. Nephrotomy and pyelotomy are used to remove nephroliths. For large stones, a pyelotomy is preferred because it causes less damage to the kidney; however, if the stones are small, the pelvis is not dilated and the procedure would be very difficult. Nephrotomy is preferred to remove small stones from the renal pelvis. It is technically easier to perform than a pyelotomy but causes more damage to the renal parenchyma, resulting in at least transient compromise of renal function. For a pyelotomy, perform a standard ventral midline celiotomy and incise the peritoneum lateral but adjacent to the kidney. Bluntly dissect the kidney free from the retroperitoneal fat and reflect it medially, exposing its the dorsal surface. Palpate the stone at the hilum of the renal pelvis and incise the pelvis with a No. 11 blade. Extend the incision to allow the stone to be removed. It is vital that the incision be made on the dorsal aspect of the renal pelvis to avoid the renal artery and vein. Once the stone or stones are removed, flush the renal pelvis with a small Teflon venous catheter and advance the catheter into the ureter to ensure its patency. Close the incision in the renal pelvis with a fine (6-0 to 8-0) monofilament suture on a small atraumatic needle. Replace the kidney to its normal position and place 2 to 4 sutures between the renal capsule and the peritoneum (nephropexy) to hold it in place and prevent renal torsion. Close routinely. To perform a nephrotomy, free the kidney from the retroperitoneal space as described above. Hold it at the hilum between the thumb and first finger. Digitally compress the renal artery and vein or occlude them with atraumatic vascular clamps or Rummel tourniquets while making an incision on the convex surface of the kidney directly through the renal parenchyma all the way down to the renal pelvis. There will be a moderate amount of hemorrhage, so work quickly. Remove the stones

CHAPTER 25  Soft Tissue Surgery

A

C

335

B

D

Fig. 25-7  A and B, A radiodense urethral calculus located at the urethral papilla is noted on abdominal radiographs of this guinea pig (arrow). C, The stone can be seen at the external orifice of the urethra (arrow). D, An incision was made over the urethra to allow the stone to be removed. The incision was allowed to heal by secondary intention. (Images courtesy of Dr. Estella Boehmer.)

and flush the renal pelvis and down the ureter to make sure all stones are removed and the ureter is patent. Once all of the stones have been removed, hold the split kidney together for 5 minutes. This allows a clot to form, and holding the two halves together and should control the hemorrhage. Place a simple continuous suture of fine monofilament material on a small needle in the renal capsule. The renal capsule is very thin and fragile, and sutures easily tear through. Replace the kidney to its normal position, perform a nephropexy, and close routinely. The ureters of hystricomorph rodents are very small, but if there is a ureteral obstruction, the lumen is usually dilated cranial to the obstruction, making surgery more feasible. The blood supply to the ureter is tenuous and must be preserved during ureterotomy. Through a standard ventral midline celiotomy, retract the viscera to the contralateral side, allowing visualization of the kidney and urinary bladder. Palpate between the kidney and bladder just lateral to the hypaxial muscles, where the ureter is located. Attempt to identify the stone by palpation in order to minimize dissection. Some distal ureteral stones can be gently manipulated into the bladder and removed through a cystotomy, which is preferable to performing a ureterotomy; however, often the stones are adhered to the wall of the ureter and cannot be manipulated into the bladder. Once the stone is located, open

the peritoneum over the stone and bluntly dissect the ureter free from the surrounding fat, being very careful not to disturb the small amount of fat directly associated with the ureter, because the ureteral vessels are within that fat. Do not attempt to clear off the fat except in the immediate vicinity of the stone. Isolate the segment with the stone and make an incision in the ureter cranial to the stone in the more dilated segment of ureter and continue the incision onto the stone. Remove the stone through as small an incision as is possible. Irrigate with a fine-gauge venous catheter in both directions. If the catheter will not advance into the caudal, smaller segment, attempt to pass a 4-0 nylon suture to confirm that it is patent caudally. If it is not patent, perform a nephrectomy. In order to minimize the risk of suturing the ureter closed, I prefer to place a stent through the ureter into the urinary bladder while closing the ureterotomy. A cystotomy is done to retrieve the stent after the ureterotomy has been closed. The stent can be a catheter or a suture placed to maintain the lumen’s patency during suture placement. It should pass into the urinary bladder so that it can be removed after the ureterotomy is closed. If the ureterotomy is small, consider closing it transversely to minimize the risk of postoperative stenosis. Once the ureter is closed, monitor the site carefully for any leakage. Urine should be flowing through the ureter and there should be no leakage.

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SECTION III  Guinea Pigs and Chinchillas

A simple continuous pattern of a very fine (8-0) monofilament material on a small needle is recommended because a continuous pattern creates a better seal than an interrupted one. Replace the ureter and close the peritoneum and cystotomy prior to routine closure of the celiotomy. Cystotomy for stone removal in rodents is routine. Make a ventral midline celiotomy incision just cranial to the pubis. Exteriorize the bladder, and examine the urinary tract carefully. Isolate the bladder with gauze sponges moistened with saline solution. If possible, place a catheter retrograde into the urethra to prevent stones from migrating into the urethra during surgery. Often, a small (24-26 gauge) Teflon intravenous catheter can be passed into the urethra of males (penis) or females (urethral opening in the clitoris). Magnification is very helpful for this procedure. Make a 2- to 3-cm cystotomy incision on the ventral aspect of the bladder, closer to the apex than the neck to avoid damaging the ureters at the trigone. Trim about 1 mm of bladder wall off one side of the incision and submit the tissue for culture and sensitivity testing. Remove the stones and irrigate the bladder to make sure that all stones are gone. Use a small intravenous catheter passed from the neck of the bladder into the urethra to confirm the patency of the urethra before closure. Submit the calculi for stone analysis and culture and sensitivity testing. Close the cystotomy in a single-layer simple interrupted or simple continuous pattern with a 4-0 or 5-0 monofilament absorbable material on a small atraumatic needle. Irrigate the abdomen with warm saline solution and close routinely. As soon as samples have been collected for culture, administer an appropriate, broad-spectrum antibiotic, which may be extended or changed based on the results of the culture and sensitivity testing. Continue to administer fluids at an appropriate rate (maintenance or higher) for 36 to 48 hours postoperatively. Hematuria may persist for several days. Stones can migrate into the urethra, which seems to be a more common occurrence in females than males, likely because the pelvic urethra in females is wider. Often urethral calculi can be manipulated or retrohydropulsed into the urinary bladder and removed through a cystotomy. In some patients, however, the calculus remains lodged, and adhered to the urethral mucosa, and cannot be moved. Perform a urethrotomy by cutting directly over the stone in the perineum, allowing it to be removed. Once the stone is removed, irrigate the site and allow it to heal by second intention. It is difficult to suture the urethra closed accurately, and it has been shown that there is no difference in healing between those sutured closed and those left to heal by second intention.44 Unsutured urethrotomies do bleed more postoperatively, but it is not enough to be of clinical concern. Not all of the factors that predispose individual rodents to the development of cystic calculi have been determined. It is difficult to cure this condition in guinea pigs, and recurrence is common.1 Cultures of the bladder wall and the calculus should be obtained during the procedure to rule out bacterial cystitis as the cause of the calculus formation. A diet low in calcium along with urine alkalinizers may be indicated; these are discussed further in Chapter 23.

CUTANEOUS DERMAL MASSES Skin and subcutaneous tumors are the second most frequently reported neoplasms in guinea pigs, representing 15% of neoplasms.8,40 Frequently they grow to a large size before the animal

is presented for treatment. Most are trichofolliculomas, which are benign tumors of basal cell origin. They are typically round, firm cutaneous masses usually in the lumbosacral region. Other cutaneous neoplasms reported include trichoepitheliomas, sebaceous adenomas, fibromas, fibrosarcomas, lipomas, fibrolipomas, undifferentiated sarcomas, and adenocarinomas.13,40 It has been suggested that the trichoepitheliomas reported in guinea pigs have actually been trichofolliculomas.13 Trichofolliculomas are typically cystic and may contain sebum, hair, and keratin debris. They can grow quite large without causing clinical problems. Aspiration of sebaceous debris from a cutaneous mass supports a diagnosis of a benign trichofolliculoma. If sebaceous and keratinaceous debris is not present cytologically, submit an aspirate or biopsy to determine the type of tumor, since the surgical and medical management of malignant dermal lesions is different. Marginal excision of a benign mass and histologic confirmation are recommended even if a benign cyst is suspected. Benign cystic masses can become infected. If surgery is delayed and the cyst becomes infected, the guinea pig will likely be ill, complicating anesthesia and surgery. Guinea pigs have a moderate amount of loose skin over the dorsum. After excising the mass, create a single-pedicle advancement flap with the pedicle or base being no less than half the length of the flap. Trim the dog ears away from the pedicle to maximize the blood supply to the flap. Use walking sutures oriented along the long axis of the flap to advance the flap and close dead space. Place simple interrupted monofilament nonabsorbable sutures in the skin. If the mass is malignant, more aggressive surgery is indicated, as described for malignant mammary tumors previously in this chapter.

CERVICAL LYMPHADENITIS Cervical lymphadenitis, a condition known as “lumps” in guinea pigs, is a streptococcal infection of the cervical lymph nodes. It has been suggested to occur secondary to oral mucous membrane trauma and usually results in lymph node abscessation.30 Isolate affected guinea pigs from other guinea pigs until the condition has resolved. The optimal treatment is complete surgical excision of any involved lymph nodes, not just lancing and draining the abscess. Provide supportive care and antibiotic therapy based on the results of culture and sensitivity testing of samples taken at surgery. If the abscesses are small, this approach may result in a cure; however, even with apparent total excision of all grossly infected tissue, the condition can recur in adjacent tissues shortly after surgery. It is important to perform surgery as early as possible, while the abscesses are small. If excision is not possible, it has been recommended that the abscess be lanced, drained, and flushed, leaving the wound open for granulation. Wound irrigation and topical as well as systemic antibiotic therapy may resolve the abscesses; however, recurrence is very common. Persistent, nonhealing abscesses can be cauterized with silver nitrate.26 The silver nitrate is caustic and kills bacteria within the cauterized tissue. Unfortunately it also causes necrosis of healthy tissue, so it must be used judiciously. More recently, antibiotic-impregnated polymethylmethacrylate (AIPMMA) beads have been used to control microscopic disease after excision of these abscesses. Excise the abscesses by removing as much infected tissue as possible so that there is no gross disease remaining. Loosely fill the dead space created by

CHAPTER 25  Soft Tissue Surgery lymph node excision with AIPMMA beads and close the subcutaneous tissues and skin over the beads routinely. The beads release antibiotic into the local tissue for an extended period and in most cases do not have to be removed.

THORACOTOMY The primary indications for thoracotomy in guinea pigs and chinchillas are pulmonary abscesses and neoplasms. Pulmonary tumors are the most common neoplasms observed in guinea pigs.13,40 Most are benign bronchogenic papillary adenomas reported to comprise 30% to 35% of all neoplasms in guinea pigs over 3 years of age,7,8,15,22 but alveolar and bronchogenic carcinomas are also reported. Most pulmonary tumors are slow growing, and clinical signs do not occur until late in the course of the disease. During a 35-year-period, only two chinchillas presented to the University of Tennessee College of Veterinary Medicine had tumors, one of which was a pulmonary adenocarcinoma.13 Thoracotomy is challenging in histricomorph rodents because endotracheal intubation is difficult. Various techniques for intubating guinea pigs have been described.23,41 If thoracotomy is indicated and the patient cannot be intubated per os, a temporary tracheostomy can be performed to establish an airway, allowing for ventilation of the patient during surgery. Make a 1.0- to 1.5-cm skin incision on the ventral cervical midline. Bluntly dissect through the subcutaneous tissues and identify the sternocephalicus muscles. Separate these muscles along the midline, being careful not to damage the thyroid vein. Identify the trachea, and bluntly dissect the peritracheal tissues off the cartilage, preserving the recurrent laryngeal nerves. Place a 3-0 nylon suture around the cartilage ring caudal to the proposed tracheotomy site and create a large loop of suture with long suture tails. Use the suture to pull the trachea to the surface, facilitating tracheostomy tube placement. Make a transverse incision in the trachea approximately one-third of the diameter of the trachea and enlarge it to 50% with hemostats to avoid cutting the recurrent laryngeal nerves. Insert a sterile endotracheal tube (1.5-2.0 mm) into the aborad segment of the trachea. After the procedure is completed and the patient is awake, remove the tracheostomy tube and allow the surgical site to heal by secondary intention. Rats maintained using a tight-fitting face mask for anesthesia and controlled ventilation had thoracotomy for lung lobectomy with very low mortality (see Chapter 28).36 This technique would likely be useful in hystricomorph rodents as well. If the mass is small enough, it can be removed through a lateral thoracotomy at the fourth-to-fifth intercostal space to gain access to the pulmonary hilum. Make a standard intercostal approach and exteriorize the lobe containing the tumor. Magnification is very helpful for this type of procedure. Identify the hilum, the pulmonary artery and vein, and the bronchus. Ideally, the pulmonary artery and vein should be isolated, ligated, and transected individually to minimize the risk of arteriovenous fistula formation. Realistically, it is best to ligate the artery, vein, and bronchus with a single ligature. Transect distal to the ligature and remove the affected lobe. Inspect the stump for hemorrhage or air leakage. Fill the chest with warm saline to observe for bubbles and allow for patient warming, and remove the solution prior to closure. Closure is routine and includes placing a thoracostomy tube to maintain negative intrapleural pressure during recovery. A 5-Fr red rubber catheter serves this

337

purpose well. Once negative pressure has been maintained for a couple of hours, remove the tube. A tube within the thoracic cavity stimulates the production of 1 to 2 mL of effusion per kilogram body weight per day. For large masses and cranial mediastinal masses, a ventral midline thoracotomy is preferred. Place the patient in dorsal recumbency and make a ventral midline incision. The approach is analogous to that used for larger animals with one exception. Because the sternebrae are very narrow, it is not feasible to split them longitudinally. Instead, cut the ribs on one side at their attachment to the sternebrae. After removal of the mass, close with figure-of-eight sutures of monofilament absorbable material encircling the sternebrae at each rib. A second layer, apposing the muscles ventrally, provides additional stability. Close subcutaneous tissues and skin routinely. Place a chest tube to allow control of the pleural space postoperatively. Pulmonary abscesses are also treated by pulmonary lobectomy; however, patients with abscesses are usually more systemically ill. Additionally, during manipulation of the lung lobe, purulent material can flow from the affected lobe into the bronchus and then into other lobes. These factors make the prognosis guarded to poor for guinea pigs, chinchillas, and degus undergoing surgical removal of pulmonary abscesses. Gently lift the affected lung lobe out of the thorax and quickly clamp the hilum to prevent pus from migrating into other lobes. Ligate or clip proximal to the clamp to control the artery, vein, and bronchus, then transect between the clamp and ligature. Check for leaks and close routinely. Long-term antibiotic therapy can be considered as an alternative to pulmonary lobectomy for abscesses; however, because of the caseous nature of the pus, medical management is often not successful in resolving the infection.

MISCELLANEOUS PROCEDURES Exploratory laparotomy for abdominal masses is indicated for hepatic cysts or neoplasms, uterine neoplasms, ovarian cysts or neoplasms, and gastrointestinal obstruction. The etiology and management of pododermatitis are discussed in Chapter 23. These fibrous granulomas of the plantar surface of the feet are usually seen in guinea pigs kept on wire floors. The prognosis for cure is guarded, and many lesions recur after treatment. It is imperative to change the husbandry conditions. Surgery should not be attempted unless the owner is willing and able to house the patient on a soft bedding and can keep the bedding clean by changing it daily. This must be done for the rest of the animal’s life. Aggressive excision of the lesions, followed by bandaging and open-wound management, allowing the surgical wounds to granulate and epithelialize, along with appropriate systemic antibiotic therapy, has met with some success. Alternatively, debridement and placement of AIPMMA beads has resulted in a cure in some cases. Debride the infected areas of purulent material and infected tissue. Place small AIPMMA beads in the defect, and suture the skin together over the beads, across the defect, to hold them in place. Pad and bandage the feet. Change the bandages every day initially and then every 2 to 3 days. The author has left beads in the feet permanently, and they have not appeared to cause lameness. Degloving of the tail, as described in Chapter 28 in gerbils, also occurs frequently in degus.1,20 Tail amputation is recommended.

338

SECTION III  Guinea Pigs and Chinchillas

References 1. Bennett RA. Rodents: soft tissue surgery. In: Keeble E, Meredith A, eds. BSAVA manual of rodents and ferrets. Gloucester: British Small Animal Veterinary Association; 2009:73-85. 2. Bennett RA, Russo EA. What is your diagnosis? Soft tissue density mass in the stomach consistent with a trichobezoar or phytobezoar. J Am Vet Med Assoc. 1985;186:812-814. 3. Beregi A, Zorn S, Felkai F. Ultrasonic diagnosis of ovarian cysts in ten guinea pigs. Vet Radiol Ultrasound. 1999;40:74-76. 4. Bodri MS, Walker LM. What is your diagnosis? Poor intra- and retroperitoneal contrast suggestive of emaciation and alimentary visceral displacement consistent with bladder and uterine mass. J Am Vet Med Assoc. 1993;202:654-655. 5. Burns RP, Paul-Murphy J, Sicard GK. Granulosa cell tumor in a guinea pig. J Am Vet Med Assoc. 2001;218:726-728. 6. Capello V. Prescrotal open technique for neutering a degu. Exot DVM. 2004;6.6:29-31. 7. Collins BR. Common disease and medical management of rodents and lagomorphs. In: Jacobson ER, Kollias GV, eds. Exotic animals. New York: Churchill Livingstone; 1988:261-316. 8. Cooper JE. Tips on tumors. In: Proceedings. North Am Vet Conf. 1994:897-898. 9. Eatwell K. Ovarian and uterine disease in guinea pigs: a review of five cases. Exot DVM. 2003;5.5:37-39. 10. Field KJ, Griffith JW, Lang CM. Spontaneous reproductive tract leiomyomas in aged guinea pigs. J Comp Pathol. 1989; 101:287-294. 11. Frisk CS, Wagner JE, Doyle RE. An ovarian teratoma in a guinea pig. Lab Anim Sci. 1978;28:199-201. 12. Gerold S, Huisinga E, Iglauer F, et al. Influence of feeding hay on the alopecia of breeding guinea pigs. Zentralbl Veterinarmed A. 1997;44:341-348. 13. Greenacre CB. Spontaneous tumors of small mammals. Vet Clin North Am Exot Anim Pract. 2004;7:627-651. 14. Harkness JE. A practitioner’s guide to domestic rodents. Denver: American Animal Hospital Association; 1993. 15. Harkness JE, Turner PV, VandeWoude S, Wheeler C. Harkness and Wagner’s biology and medicine of rabbits and rodents. 5th ed. Ames, IA: Wiley-Blackwell; 2010. 16. Hillyer EV. Common clinical maladies of pet rodents. In: Proceedings. North Am Vet Conf. 1994:909-910. 17. Hoefer HL. Guinea pig urolithiasis. Exot DVM. 2004;6.2:23-25. 18. Hoefer HL. Chinchillas. Vet Clin North Am Small Anim Pract. 1994;24:103-111. 19. Jenkins JR. Surgical sterilization in small mammals. Spay and castration. Vet Clin North Am Exot Anim Pract. 2000;3:617-627. 20. Johnson D. Exotic pet care: degus. Exot DVM. 2002;4.4:39-42. 21. Keller LS, Griffith JW, Lang CM. Reproductive failure associated with cystic rete ovarii in guinea pigs. Vet Pathol. 1987;24:335-339. 22. Kitchen DN, Carlton WW, Bickford AA. A report of fourteen spontaneous tumors of the guinea pig. Lab Anim Sci. 1975;25:92-102. 23. Kramer K, Grimbergen JA, van Iperen DJ, et al. Oral endotracheal intubation of guinea pigs. Lab Anim. 1998;32:162-164.

24. Kuenzel F, Hittmair K. Sonographische diagnosestrellung eines trichobezoars bei einem langhaarmeerschweinchen. Wein Tieraerztl Mschr. 2002;89:66-69. 25. Mayer J. The use of GnRH to treat cystic ovaries in a guinea pig. Exot DVM. 2003;5.5:36. 26. Mullen HS. Nonreproductive surgery in small mammals. Vet Clin North Am Exot Anim Pract. 2000;3:629-645. 27. Nelson WB. Technique for neutering pet chinchillas. Exot DVM. 2004;6.5:27-30. 28. Nielsen TD, Holt S, Rueløkke ML, et al. Ovarian cysts in guinea pigs: influence of age and reproductive status on prevalence and size. J Sm Anim Pract. 2003;44:257-260. 29. Peters LJ. The guinea pig: an overview. Part I. Compend Contin Educ Pract Vet. 1991;4:15-19. 30. Peters LJ. The guinea pig: an overview. Part II. Compend Contin Educ Pract Vet. 1991;5:20-27. 31. Popesko P, Rajtova V, Horak J. Atlas of the anatomy of small laboratory animals, vol. 1. Rabbit and guinea pig. London: Wolfe Publishing; 1992;148–240. 32. Powers MY, Campbell BG, Finch NP. Preputial damage and lateral penile displacement during castration in a degu. J Am Vet Med Assoc. 2008;232:1013-1015. 33. Quattropani SL. Serous cystadenoma formation in guinea pig ovaries. J Submicrosc Cytol. 1981;13:337-345. 34. Quattropani SL. Serous cysts of the aging guinea pig ovary. I. Light microscopy and origin. Anat Rec. 1977;188:351-360. 35. Redrobe S. Soft tissue surgery in rabbits and rodents. Sem Avian Exot Pet Med. 2002;11:231-245. 36. Roman CD, Hanley GA, Beauchamp RD. Operative technique for safe pulmonary lobectomy in Sprague-Dawley rats. Contemp Top Lab Anim Sci. 2002;41:28-30. 37. Rueløkke ML, McEvoy FJ, Nielsen TD, et  al. Cystic ovaries in guinea pigs. Exot DVM. 2003;5.5:33-36. 38. Syme HM, Barber PJ, Markwell PJ, et al. Prevalence of systolic hypertension in cats with chronic renal failure at initial evaluation. J Am Vet Med Assoc. 2002;220:1799-1804. 39. Theus M, Bitterli F, Foldenauer U. Successful treatment of a gastric trichobezoar in a Peruvian guinea pig (Cavia aperea porcellus). J Exot Pet Med. 2008;17:148-151. 40. Toft JD. Commonly observed spontaneous neoplasms in rabbits, rats, guinea pigs, hamsters, and gerbils. Sem Avian Exot Pet Med. 1992;1:80-92. 41. Turner MA, Thomas P, Sheridan DJ. An improved method for direct laryngeal intubation in the guinea pig. Lab Anim. 1992;26:25-28. 42. van Goethem B, Schaefers-Okkens A, Kirpensteijn J. Making a rational choice between ovariectomy and ovariohysterectomy in the dog: a discussion of the benefits of either technique. Vet Surg. 2006;35:136-143. 43. Wallach JD, Boever WJ. Diseases of exotic animals: medical and surgical management. Philadelphia: WB Saunders; 1983. 44. Weber WJ, Boothe HW, Brassard JA, et al. Comparison of the healing of prescrotal urethrotomy incisions in the dog: sutured vs nonsutured. Am J Vet Res. 1985;46:1309-1315.

SECTION FOUR

Small Rodents

CHAPTER

26

Basic Anatomy, Physiology, Husbandry, and Clinical Techniques  

Angela M. Lennox, DVM, Diplomate ABVP (Avian), and Louise Bauck, DVM, MVSc

General Characteristics Rats Mice Hamsters Gerbils Anatomic and Physiologic Characteristics General Sexing Rats Mice Hamsters Gerbils Husbandry Housing and Equipment Diet and Feeding Zoonosis Clinical Techniques Handling and Restraint Sample Collection Other Diagnostic Testing Procedures Hospitalization Therapeutics

GENERAL CHARACTERISTICS The hundreds of species belonging to the order Rodentia are grouped into three suborders based on the anatomic and functional differences of the masseter muscle. These three groups are the Caviomorpha (guinea pig-like); the Sciuromorpha Copyright © 2012 by Saunders, an imprint of Elsevier Inc.

(squirrel-like); and the Myomorpha (mouse- or rat-like).20 Although body size varies greatly, in general rodents possess a uniform body structure with various adaptations. While the medical approach to the many small rodents species commonly kept as pets is similar, unusual species are sometimes encountered in practice, and more specific information about these species can be found elsewhere.19 The suborder Myomorpha contains many species of small rodents that appear in the pet trade, including rats, mice, hamsters, and gerbils. Species in this group possess elodont incisors and anelodont cheek teeth.6 The testicles and scrotum are usually large in relation to the overall body size, and the inguinal canal is open, allowing the testicles to pass freely from the abdomen to the scrotum.20 Sciruomorph species such as squirrels and chipmunks are less commonly kep.

RATS Rats (Rattus norvegicus) are common pets and are considered one of the better rodent pets because of their larger size and calm nature. Rats are most popular in the pet market in hooded color varieties, in which the coat color is present only over the head and shoulders. Rats are generally hardy as young animals but may suffer from obesity, chronic respiratory disease, and mammary tumors when older. Rats are large enough to be easily grasped by children, and they rarely bite. Some may be excitable and run when removed from their cages; however, rats have been known to return to their cages after “escaping.” Rats are social and can live in mixed sex groups, and males may be present while females are raising litters. Introduction of strangers can be performed successfully on neutral territory.20 Rats are relatively intelligent and seem interested in humans; they can be trained to come when called for a treat. 339

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SECTION IV  Small Rodents

Giant Gambian pouched rats (Cricetomys gambianus), which weigh up to 1.5 kg, are rare in the pet trade. The U.S. Department of Agriculture (USDA) issued a temporary ban on their importation after monkey poxvirus was identified in a group of Gambian rats shipped to the United States in 2003.23 The ban was lifted in 2008. In 2004, the Gambian pouched rat was discovered to have established a breeding population in the Florida Keys. Efforts to control their populations are under way.28

MICE Standard laboratory mice (Mus musculus) in different color and coat varieties are common as pets. An adult mouse weighs approximately 30 g. Mice make good pets for older children (10 years of age and up) because they rarely bite, but they may move quickly, so younger children may not be able to handle them. Mice are largely nocturnal but are easily roused. Female mice or castrated male mice are recommended over intact males owing to the strong odor of the latter.20 In general, mice are more solitary than other common rodent pets. Female mice usually do well together, but intact males cannot be kept together, as aggression, injury, and death can result. Pet mice are hardy and rarely suffer from infectious disease; however, mite infestations are common and are difficult to treat. Pneumonia and mammary tumors are also frequently seen. Other less common species, including a variety of African spiny mice (Heteromys species) are kept as pets, but their smaller size makes them more difficult for children to handle. Characterized by their dorsal, inflexible spine-like hairs, spiny mice are likely more closely related to the gerbil.9 In the wild, spiny mice are omnivorous.9

HAMSTERS Hamsters include several groups frequently encountered as pets. Hamsters are the least hardy of the small rodents when newly purchased, and stress-related diseases such as proliferative ileitis are common. Hamsters are nocturnal, but if they are scooped up gently with two hands, they usually awaken without attempting to bite. Touching a hamster’s back with a finger in an attempt to rouse it is likely to provoke a startle or threat response. Excited hamsters often jump from hands or tables, so use appropriate caution in handling them. All hamsters are well known for their ability to escape. Chewed or damaged cage parts should be replaced immediately; if a hamster is cared for by a child, an adult should regularly check the pet’s cage for signs of wear. Hamsters may be stressed by hot, humid environments; therefore an effort should be made to keep them in a cool area of the house during the summer months. The most common species is the golden or Syrian hamster (Mesocricetus auratus), which comes in a variety of coats, including the long-haired or Teddy Bear breed.20 Dwarf hamsters include the closely related Campbell’s ­Russian dwarf (Phodopus campbelli), the Russian dwarf or Djungarian hamster (Phodopus sungorus), and the Roborovskii or desert hamster (Phodopus roborovskii). They are small (average 25-50 g) and have furred short tails, white underparts, and a grayish to tan dorsal surface with or without a dorsal stripe, depending on the species. They are excitable and more difficult to restrain than golden hamsters, and they may bite when restrained. Dwarf

hamsters are more social than golden hamsters and are more likely to live in family groups. Chinese hamsters (Cricetulus griseus) are similar in size to the dwarf hamsters but are not as social and are best kept separately.

GERBILS In nature, gerbils (Meriones unguiculatus) are desert dwellers with efficient kidneys for conserving water.20 Pet gerbils are available in white, black, buff, gray, and spotted varieties. They are extremely active and may be difficult for smaller children to handle; they can slough their tail skin if the tail is caught or grasped too firmly.20 While gerbils are social in nature and live in family groups, they are territorial and often will not tolerate introduction of strangers (cannibalism can result from attempting to keep incompatible pairs or males together). Gerbils are more disease-resistant than hamsters, although older gerbils may develop a variety of neoplastic and degenerative conditions. Epilepsy has been reported in gerbils but is uncommon in many pet strains.

ANATOMIC AND PHYSIOLOGIC CHARACTERISTICS GENERAL The word rodent is derived from the Latin verb rodere, which means “to gnaw.” Small rodents of the Myomorpha suborder possess a common dental formula: 2(I1/1, C 0/0, M 3/3). The four prominent, incisors are elodont, or continuously growing throughout life, while cheek teeth are anelodont, and do not grow after eruption. The enamel of most common rodents is white; however, some species may have enamel that is orange to yellow in color. The crowns of the mandibular incisors are longer than the maxillary incisors and may be mistakenly assumed to be overgrown. In general, the crown/length ratio for the upper to lower incisors is approximately 1:3. Many small rodents have bulging eyes and may appear exophthlamic, especially when scruffed. The harderian gland, which lies behind the eyeball, produces lipid- and porphyrin-containing secretions that aid ocular lubrication and play a role in pheromone-mediated behavior. These secretions impart a red tinge to the tears and fluoresce under ultraviolet light. Normally, the lacrimal secretions are spread over the pelage during daily grooming. However, in stressful situations and in certain disease conditions, there may be an overflow of tears; this can be inaccurately diagnosed as bleeding from the eyes and nose. Rodents are monogastric, with many species having a forestomach that is separated from the glandular stomach by a limiting ridge. They have a relatively large cecum and an elongated colon. Most rodents practice some degree of coprophagy. The ingested fecal pellets presumably provide nutrients, such as B vitamins, produced by the colonic bacteria. Most common rodent species do not vomit, in part because of the limiting ridge in the stomach; but other factors play a role as well, such as the pressure and strength of the esophageal sphincter and crural sling and the innervation of the diaphragm.27,29 For this reason and because these small mammals have such a high metabolic rate, preoperative fasting is not required or recommended. The urinary and reproductive tracts terminate in separate urethral and vaginal orifices in the female. Small rodents are spontaneous ovulators and are polyestrous. Many breed prolifically

CHAPTER 26  Basic Anatomy, Physiology, Husbandry, and Clinical Techniques in captivity. Stages of the estrous cycle can be determined with vaginal cytology. Mammary tissue can be extensive in rodents and ranges from over the shoulders to the perianal region. Mammary tumors can develop anywhere along this tract. Most female rats and hamsters have 6 pairs of nipples, while gerbils have 4 and mice have 5; however, variations in numbers  can be seen. Hamsters, rats, and mice possess four front toes and five hind toes, which is opposite in gerbils. All rodents have tails; they are longer than the animal’s body in gerbils, rats, and mice. Golden hamsters have very short tails, while dwarf hamsters have relatively longer tails. Rodents do not pant and have no sweat glands; therefore their ability to withstand high temperatures is limited. Heat dissipation occurs through the ears and tails. Some smaller rodents may salivate in response to warm temperatures.

341

Normal physiologic, reproductive, and growth reference ­values are presented in Tables 26-1 and 26-2.

SEXING Determining the sex of most rodents is easy in mature animals but can be more challenging in very young ones. In general, the distance between the anus and the genital papilla is a reliable method of determining the sex of young animals. The anogenital distance is greater in males than in females, and the genital papilla is usually more prominent and has a round opening in the male. Examining multiple young animals to make a comparison is helpful. The testes of mature males are well developed, especially in rats. Holding the rodent vertically or applying gentle pressure directed caudally on the abdomen allows the testes to pass from the abdomen through the inguinal canal

  Table 26-1     Normal Physiologic Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value

Gerbil

Hamster

Mouse

Rat

Average life span (months) Maximum reported life span (months) Average adult weight (g), male Average adult weight (g), female Heart rate (beats per minute) Respiratory rate (breaths per minute) Tidal volume (mL) Minute volume (mL) Rectal temperature (°C) Approximate daily diet consumption of adult (g) Approximate daily water consumption of adult (mL) Approximate daily fecal production (g) Recommended environmental temperature (°C) Recommended environmental relative humidity (%) Total blood volume (mL/kg)

24-39 60 46-131 50-55 260-600 85-160 — — 38.2 5-7 4 1.5-2.5 18-22 45-55 60-85

18-36 36 87-130 95-130 310-471 38-110 0.8 64 37.6 10-15 9-12 2-2.5 21-24 40-60 65-80

12-36 48 20-40 22-63 427-697 91-216 0.15 24 37.1 3-5 5-8 1-1.5 24-25 45-55 70-80

26-40 56 267-500 225-325 313-493 71-146 0.6-1.2 220 37.7 15-20 22-33 9-15 21-24 45-55 50-65

aAverage

reference values from data given in references 3, 12, 13, 15, 17, 18, and 22. Note that the ranges should be considered as guides; values are likely to vary between groups of animals according to such variables as strain, age, sex, fasted, and methodology.

  Table 26-2     Normal Reproduction and Growth Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value

Gerbil

Hamster

Mouse

Rat

Estrogen cycle length (days) Estrus (heat) duration (hours) Length of gestation (days) Pups per litter Weight at birth (g) Eyes open (days) Ears open (days) Hair coat starts (days) Start to eat dry food (days) Optimal weaning age (days) Age of maturation of male (weeks) Age of maturation of female (weeks) Recommended minimum breeding age (weeks) Chromosome number (diploid)

4-7 12-18 23-26 3-8 2.5-3.5 16-21 5 6 16 21-28 9-18 9-12 10-14 44

4-5 8-26 15-18 5-10 1.5-3 12-14 4-5 9 7-10 19-21 8 6 8 44

4-5 9-20 19-21 7-11 1-1.5 12-14 10 10 12 18-21 6 6 8 40

4-5 9-20 21-23 6-13 4-6 12-15 2.5-3.5 7-10 14 21 4-5 4-5 9 42

aAverage

reference values from data given in references 1, 3, 4, 12-14, 21, and 22. Note that the reference values may not represent the mean or range for certain populations or strains of animals; as a result, the values should be interpreted as approximations.

342

SECTION IV  Small Rodents

HAMSTERS

Fig. 26-1  Anatomy of a female hamster, dorsal recumbency. Note the separate urinary (white arrow) and vaginal (yellow arrow) orifices ventral to the anus (green arrow). (Modified from Capello V. Pet hamsters: selected anatomy and physiology. Exot DVM 3.2, Zoological Education Network, 2001.)

into the scrotum, aiding identification. Female rodents have separate vaginal and urethral orifices, the vaginal orifice being between the urethral orifice and the anus (Fig. 26-1). However, it is difficult to identify the vaginal orifice in immature and very small animals. Grossly observable nipples are seen only on the females of these species. Nipples are observable at 10 days of age in female mice and rat pups.

RATS Because of the popularity of rats in the biomedical research community, an extensive amount of information is available on rat anatomy, physiology, behavior, and diseases. In general, rats are typical rodents, many of the pertinent features of which have already been described. Several other points are useful to remember. Albino strain rats, compared with their pigmented peers, have poor eyesight and rely heavily on their vibrissae for spatial orientation. Rats do not have gallbladders. The white hair coat of rats often yellows with age, and the tail becomes more dry and scaly. Aged male rats develop brown, granular sebaceous secretions at the base of their hair shafts, which some owners may mistake for ectoparasitism.

MICE Male mice are typically twice the size of female mice. Like most other male rodents, they have open inguinal canals, an os penis, and a complex urogenital system that contains several prominent accessory glands. Intermale aggression is a common problem, particularly if the males were not raised together or if they are housed in a confined space with mature females. Male mice produce a characteristic musty odor. Pheromones play an important role in mouse behavior and are mediated through tissues such as the vomeronasal (Jacobson’s) organ, which is located in the floor of the nasal cavity. Estrus is suppressed in female mice housed in large groups (the Whitten effect). Recently bred mice that are exposed to a strange male may have impaired implantation (the Bruce effect).

Hamsters are short-tailed, stocky rodents known for their abundance of loose skin. They have large, potentially reversible cheek pouches; these are paired muscular sacs extending as far back as the scapula. The pouches are evaginations of the oral mucosa and are used for transporting food, bedding material, and occasionally young. Hamsters have a distinct forestomach that, like a rumen, has a high pH and contains microorganisms. Golden hamsters have distinctive hip or flank glands that should not be mistaken for skin tumors. These dark brown patches are found bilaterally along the lumbar area. They are poorly developed in the female, but in the mature male they are prominent and become wet and matted during sexual excitement. Dwarf hamsters (Phodopus species) have a midventral sebaceous gland. Secretions from this gland play a role in territorial marking and mating behavior. Female golden hamsters are typically larger than the males and produce a copious vaginal discharge, normally just after ovulation (day 2 of the estrous cycle). These secretions should not be misinterpreted as indicating a bacterial infection of the genital tract. These females also have paired vaginal pouches that collect exfoliated cells and leukocytes. Russian hamster males are larger than females, and the females do not normally produce a vaginal discharge after ovulation. Hamsters are permissive hibernators. Low environmental temperatures stimulate them to gather food. At temperatures of about 41°F (5°C), they may curl up and enter a deep sleep.

GERBILS Gerbils are adapted to a desert environment. They require very little water and produce only a small volume of concentrated urine. In their natural habitat, they can obtain most or all of their water requirements from metabolic processes and from any available fruit or vegetable matter. Despite this remarkable natural ability, pet gerbils should always have access to fresh water. The gerbil’s red blood cell has a life span of approximately 10 days. The rapid turnover of red blood cells is reflected on a stained blood smear as a pronounced basophilic stippling in a high percentage of these cells. Gerbils of both sexes have a distinct orange-tan oval area of alopecia on the midventral region referred to as the ventral marking gland or pad. This structure is composed of large sebaceous glands that are under the control of gonadal hormones. In the pubescent male, the gland starts to enlarge and produces an oily musk-scented secretion. Gerbils can often be seen rubbing their abdomens on objects; this is thought to be a form of territorial marking.

HUSBANDRY HOUSING AND EQUIPMENT Suitable enclosures for rodents should be escape-proof and easy to clean. While colorful and interesting, multilevel cages with tubes, wheels, and hide boxes may be so difficult to disassemble that basic cleaning is neglected. Manufacturers recommend that the entire cage be disassembled and washed thoroughly, which in reality is rarely done. Newer multilevel

CHAPTER 26  Basic Anatomy, Physiology, Husbandry, and Clinical Techniques

Fig. 26-2  Ideal habitat for pet rats. The cage portion can be separated from the base for ease of cleaning. Note the cloth and plastic “dens,” exercise wheel, and cardboard box filled with paper for enrichment.

wire cages sit on a plastic base and can be separated to facilitate cleaning (Fig. 26-2). This type of cage provides the additional benefit of separating living space up and away from urine and feces. Features of optimal enclosures are slide-out or easy-toremove bottoms for ease of cleaning; bottoms with high sides to contain bedding; adequate ventilation (aquariums are not ideal owing to poor ventilation); large doors for easy access to the pet; and a secure locking mechanisms for each cage opening. Frequent cleaning of the cage is critical in the care of pet rodents. Failure to clean the cage results in the buildup of ammonia and contributes to stress and illness. In mice and rats, Mycoplasma pulmonis organisms multiply more rapidly in the presence of ammonia levels of 50 to 100 ppm.31 The frequency of cage cleaning depends on the cage size and number of animals housed. Advise owners to notice the odor of the bedding; anything other than the scent of clean litter indicates that the cage should be cleaned. Provide food in heavy crocks or food dispensers so that the containers will not be tipped over. Bedding choices include recycled paper products, corncob products, shredded paper, and shavings of woods such as pine and various hardwoods. Much debate exists on the use hardwood and aromatic shavings such as cedar; anecdotally their use is linked to skin and respiratory disease. Paper bedding is generally preferable, although these products are more expensive than wood shavings. In a study of the endotoxin, dust, and coliform content of 20 types of rodent bedding, endotoxin and coliform levels were lowest in paper bedding, and these products were recommended to reduce the risk of respiratory disease and immune suppression in laboratory rodents.35 In rats, the rate of sneezing and incidence of lung pathology was higher in animals housed on aspen shavings than in those housed on paper bedding.5 However, results of one study in laboratory mice found no difference

343

in growth, food intake, oxygen consumption, IgE antibody concentrations, or general appearance and behavior in male CD-1 mice kept on CareFRESH (Absorption Corp., Ferndale, WA, USA) original bedding, cedar shavings, or pine shavings over a 4-month period.2 In a study evaluating the dermal toxic effects of cedar and juniper on mice and rabbits, concentrations normally found in wood shavings did not elicit any hypersensitivity reaction; reactions were seen only at much higher concentrations (50% or more).10 The housing of gerbils on shavings is indirectly involved in the development of facial lesions. Sand or dust normally present in the natural environment is absent in cages with shavings, and dust bathing is thought to be part of gerbils’ normal grooming procedure. Sandboxes can be provided for gerbils in much the same way that they are for chinchillas. Enrichment refers to providing mental stimulation and is appropriate for all captive animals. Enrichment for rodents is provided in the form of exercise wheels, hide boxes, materials to shred, treats wrapped in paper or hidden in toys, and time spent outside the enclosure interacting with the owner. Hide boxes can be commercially purchased or constructed from readily available materials such as polyvinyl chloride (PVC) pipe or cardboard boxes. Cardboard provides the additional advantage of a material that can be shredded, destroyed, and discarded when soiled. Smaller cat treat balls have been used successfully with larger rodents. Many small rodents, especially hamsters, enjoy exercise wheels. Plastic exercise wheels that are almost noise-free are available, although some hamsters may chew on them if other materials, such as soft wood blocks, are not available.

DIET AND FEEDING Rodents naturally hoard food items, and exactly how much food is actually being consumed is difficult to gauge by the rate of disappearance from the feeder. Water can be provided in bowls, but good-quality water bottles will prevent bedding from getting into the water and are generally preferred. Water bottles can malfunction over time, resulting in blockage or leakage. Educate owners to change water and test water bottles daily. Most rodents are omnivorous, often eating grasses, seeds, grain, and occasionally invertebrates in the wild.20 Dietary requirements of species in laboratory settings are well established. In the pet environment, needs are best met with a formulated diet supplemented with small amounts of fresh foods and seeds for variety and interest. Although seed mixes are popular choices for rodents, these diets often lead to selective feeding. Animals usually consume high-calorie seeds (sunflower) and ignore formulated pellets, resulting in dietary imbalance. While many rodents have such a short life span that dietary deficiency is rarely recognized, the most common adverse outcome in rats is obesity. Also of interest are various studies demonstrating increased longevity and reduction of certain diseases in rats maintained on a calorie-restricted diet.26 Gerbils have been used extensively in nutritional studies of dietary fat because of their sensitivity to high fat, high cholesterol diets with resultant changes in blood cholesterol levels. Protein requirements for rodents vary from 14% to 17% in hamsters, from 14% to 16% in rats and mice, and up to 22% in gerbils.20 Formulated diets for hamsters, rats, mice, and gerbils should reflect these requirements. Diets for reproduction in rodents should contain higher levels of protein.3

344

SECTION IV  Small Rodents

ZOONOSIS Significant public attention has focused on the zoonotic risk of rodents, in particular lymphocytic choriomeningitic (LCM) virus and hantavirus. While the natural reservoirs of these diseases are various species of wild rodents, there are documented cases of human infection with LCM from exposure to pet rodents.8 Pet hamsters, in particular, carry a wide variety of potentially infectious agents. In general, use standard precautions in handling pet rodents, including thorough handwashing. Because many rodent species are popular pets for children, discuss potential anthropozoonoses with clients, especially if pets are kept in a school setting (see Chapter 40).

CLINICAL TECHNIQUES HANDLING AND RESTRAINT The handling and restraint of small rodents can be challenging. The key is to maximize diagnostic information while maintaining safety for both the examiner and the patient. If at any time the animal appears to be in distress, release it immediately and plan an alternative technique. Many owners bring pet rodents to the clinic in their normal enclosures, which is helpful in terms of observing husbandry conditions and examining fecal output. However, attempting to retrieve a rodent from colorful caging tubes is time-consuming and best avoided. Therefore request that owners confine the pet to a separate carrier or a small travel cage within the cage to facilitate capture and restraint. Much information can be gained by examining the pet as it moves about on the examination table, including respiratory rate and effort, overall demeanor, and gait. Take care to prevent the rodent from leaping from the table. Actual

A

restraint technique varies as to species, patient size, temperament, and overall condition. Watching how owners handle their pets will provide information on how the pets will react to restraint. Most small pet rodents do not object to being held loosely in a cupped hand. Exceptions include rodents that have not been sufficiently acclimated to handling and animals in discomfort or distress. As an example, a hamster with enteritis and intestinal gas will appear to be in great discomfort and will roll onto its back, bite, and resist all attempts at handling. The same animal is often calm and will accept handling once the discomfort has subsided. In general, restraint can be accomplished by three methods: (1) examining the animal as it is loose on the table or in the hand; (2) restraining the animal with small towels or cloths; or (3) restraining the animal by some form of scruffing. In some cases, combinations of the above methods are used. For rats, holding firmly at the base of the tail allows the examiner to slow the animal as it walks freely on the table. Do not use this technique for rodents with furred tails, as this presents risk of a degloving injury of the skin (“tail slip”). Scruffing is effective for many species (mice, hamsters, and gerbils) with the exception of rats, which do not tolerate this method. To scruff, grasp generous amounts of loose skin from the lateral aspects of the neck and over the shoulders with thumb and forefinger (Fig. 26-3). Failure to incorporate enough skin allows the animal to rotate and bite the restrainer’s fingers. Calm, large rodents can be restrained by grasping them about the shoulders and thorax (Fig. 26-4). Extremely fractious or distressed rodents benefit from mild sedation for examination or diagnostic testing. Take care, however, with sick or debilitated patients. In general, if the animal is alert enough to defend itself vigorously, it will tolerate sedation. A combination of midazolam (0.25-0.5 mg/kg) with an opioid

B

Fig. 26-3  Restraint methods for rodents. Hamsters, gerbils, and mice can be restrained by scruffing, being careful to include a generous amount of skin to prevent the patient from turning and biting (A). Larger, calm rodents can often be grasped gently around the thorax (B). (Courtesy of Angela Lennox, DVM.)

CHAPTER 26  Basic Anatomy, Physiology, Husbandry, and Clinical Techniques such as butorphanol (0.1-0.4 mg/kg) or hydromorphone (0.1 mg/kg) is generally safe and effective for light sedation (see Chapter 31 and Appendix). Restrain the rodent gently with a cloth and, with a hind limb exposed, inject quickly with a 27-g needle and small syringe (Fig. 26-5) and return the animal to its enclosure. If additional sedation is needed, administer ketamine or consider brief general anesthesia. To induce anesthesia, transfer the small rodent directly into a small to medium-size face mask; afterward, replace the mask with a rodent-sized anesthetic cone or modified syringe case for maintenance (Fig. 26-6). Very small induction chambers can also be used, but avoid using a large induction chamber because of the greater volume of escaped anesthetic gas into the clinic environment and resultant staff exposure.

345

Intubation has been described in large rodents and can been performed with the aid of endoscopy or specialized equipment used in laboratory animals—in particular, a small pediatric laryngoscope and positioning device for the rat.30 Success requires considerable practice. In clinical medicine, rodents are not commonly intubated during anesthesia.

SAMPLE COLLECTION Diagnostic testing can provide the same benefits in rodents as it does in other species. Improved techniques and the ability to acquire meaningful results from small sample sizes make sample collection possible and practical in rodents. Reference intervals for selected species for hematologic and biochemical testing, serum protein electrophoresis, and urinalyses are presented in Tables 26-3 through 26-9. Reference values are often derived from laboratory populations and will vary with animal age, sex, strain, and husbandry as well as with the method of sample collection and laboratory methodology used to derive the values. Many published reference ranges are derived from very small sample numbers and therefore must be interpreted and used with caution (Carolyn Cray, personal communication).

Blood Sample Collection

Fig. 26-4  Most tame rats can be calmed using this technique: the rat is suspended with the rear feet dangling while the head and ears are gently rubbed. (Courtesy of Angela Lennox, DVM.)

Fig. 26-5  Intramuscular injection can be accomplished in most rodents with towel restraint and exposure of the rear leg.

Venipuncture should be attempted only in normovolemic, normothermic patients; therefore blood collection is often delayed until the animal’s condition is stabilized. Before collecting a sample, the clinician must know the sample volume required for the test in question and the maximum blood volume that can be safely acquired from the patient. The blood volume of most rodents is approximately 6% to 7% of total body weight34; removing a maximum of 10% of the blood volume is generally safe (see Table 26-1). For example, a 100-g gerbil is assumed to have approximately 6 to 7 mL total blood volume; of that, no more than 0.7 mL should be collected. Make modifications to these guidelines according to overall patient condition and, in all cases, collect only the blood volume required for testing. If submitting to a reference laboratory, consult the laboratory as to minimum sample requirements and if serum or plasma samples are preferred. Some laboratories require serum for biochemical analysis, whereas others routinely run analyses on plasma samples to maximize sample volume. The volume of blood required for both a biochemical analysis and an automated complete blood count (CBC) may be impractical to collect in small rodent patients. However, the required volume can be reduced if the laboratory is willing to perform a manual count or estimate the CBC from a blood film. In-house analysis of blood samples is convenient for two specific reasons: the speed of results, which is often important for sick rodent patients, and the ability to use very small sample volumes. The Abaxis VetScan (Abaxis, Union City, CA) will perform a preanesthetic or diagnostic biochemical panel on 0.13 mL of whole blood, putting testing within reach for even very small rodent patients. A single drop of blood collected into a hematocrit tube can provide a blood film for an estimated white blood cell (WBC) count, differential, hematocrit, and estimation of total serum solids. Using these techniques, a biochemical panel and modified CBC as described above can be obtained in a 25-g mouse (6% of body weight = 1.5 mL total blood volume; 10% × 1.5 mL = 0.15 mL).

SECTION IV  Small Rodents

346

B

A

C Fig. 26-6  Induction of general anesthesia in a sedated rat by using a standard small-animal mask as an induction chamber (A). Once anesthetized, anesthesia is maintained with a modified syringe case (B) or a commercial rodent face mask (C). (Courtesy of Angela Lennox, DVM.)

  Table 26-3     Hematologic Data for Micea Value Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x106/μL) Nucleated RBCs (per 100 WBCs) White blood cells (x103/μL) Segmented neutrophils (%) Band neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Platelets (103/μL)

44 (40-48) 12.6 (9.9-15.3) 7.2 (6.9-7.5) 0 (0-2)

39 (33-34) 13.4 (11.4-15.4) 5.4 (4.5-6.3) 0 (0-2)

34-50 12.8-16.1 7.5-9.7 —

12.9 (5.5-20.3) 27 (10-45) 0 (0-2) 70 (50-85) 2 (0-6) 1 (0-3) 0 (0-2) 1,200 (1,000-1,400) 79 (69-89) 27 (24-30)

8.7 (4.1-13.3) 29 (15-43) 0 (0-2) 66 (49-84) 3 (0-6) 2 (0-4) 0 (0-2) 1,200 (1,000-1,400) 73 (65-81) 25 (22-28)

4.5-9.1 21-57 — 49-82 2-8 0-3 0-3 421-733

34 (32-36)

34 (31-37)

—

Mean corpuscular volume (fL) Mean corpuscular hemoglobin (pg) Mean corpuscular hemoglobin concentration (g/dL) aValues

(n = 20)

Femaleb

Mixed sexc (n = 50)

Maleb

(n = 20)

— —

are given as mean, mean ± 2 SD, or range. River outbred mice, CD-1 (ICR)BR, 32 to 34 weeks, raised under optimal laboratory conditions. (Available at www.criver.com/sitecollectiondocuments/rm_rm_r_hematology_sex_age_outbred_ mice.pdf) cData courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Automated analyzer for cell count, hemoglobin, and hematocrit. Differential based on 100-cell count by using blood smear made at the time of sample acquisition. bCharles

CHAPTER 26  Basic Anatomy, Physiology, Husbandry, and Clinical Techniques

347

  Table 26-4      Biochemical Data for Micea Analyte

Male (n = 26)b

Female (n = 26)b

Mixed sex   (n = 50)c

Albumin (g/dl) Alkaline phosphatase (IU/L) Alanine aminotransferase (IU/L) Aspartate aminotransferase (IU/L) Bilirubin, total (mg/dl) Blood urea nitrogen (mg/dl) Calcium (mg/dl) Chloride (mEq/L) Cholesterol (mg/dl) Creatinine (mg/dl) Glucose (mg/dl) Gamma-glutamyl transpeptidase (IU/L) Phosphorus (mg/dl) Potassium (mEq/L) Sodium (mEq/L) Total protein (g/dl) Triglycerides (mg/dl)

3.3 ± 0.3 173 ± 45 48 ± 18 101 ± 33 0.22 ± 0.08 16 ± 3 11.3 ± 0.4 106 ± 2 160 ± 20 0.12 ± 0.04 207 ± 18 0.2 ± 0.5

3.6 ± 0.2 191 ± 47 41 ± 19 92 ± 38 0.18 ± 0.04 15 ± 3 10.6 ± 0.5 108 ± 3 133 ± 30 0.12 ± 0.04 225 ± 28 0

2.5-4.8 51-285 29-191 — 0.1-0.9 18-29 8.7-10.1 — — 0.1-0.4 90-193 —

11.9 ± 1.2 10.0 ± 0.2 153.0 ± 1.6 5.1 ± 0.6 100 ± 22

11.2 ± 0.4 8.8 ± 0.9 152.0 ± 2.8 5.2 ± 0.1 225 ± 21

5.4-9.3 — — 4.6-6.9 —

aValues

are given as mean ± SD or range. River, U.S., and Canadian colonies, CD-1(ICR) mice, 56 to 70 days of age, raised under optimal laboratory conditions. (Hitachi 717 Olympus AU 640e). Values rounded. (Available at http:// www.criver.com/sitecollectiondocuments/rm_rm_r_cd1_mouse_biochemistry_jun_dec05.pdf) cSerum samples courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak:Ektachem) 700XR. bCharles

The collection technique depends on patient size and temperament and clinician familiarity. The “correct” technique is that which provides consistent diagnostic-quality samples with optimal patient safety. Samples taken after repeated venipuncture attempts and the use of excessive negative pressure often result in hemolysis and poor sample quality, which can have a negative effect on results. Depending on patient size, samples can be collected with small-gauge needles (25-27 g) with attached small syringes (1 mL or smaller). In very small patients or with vessels where negative pressure causes collapse of the vessel, consider puncturing the vessel with a clear-hubbed needle only and collecting blood with a heparinized hematocrit tube as it flows into the needle hub. Most rodents do not tolerate the level of restraint required to safely collect an adequate volume of blood for analysis. Techniques for restraint of laboratory rodents for venipuncture may not be appropriate for pet rodents. Simple sedation as described above, with the addition of brief general anesthesia if needed, greatly facilitates blood collection in many animals. With thoughtful planning, a single sedation procedure can facilitate complete examination, diagnostic testing, and some therapeutic procedures as well. The cranial vena cava is the largest easily accessible vessel in rodent species. In general, it is located just within the thoracic cavity, dorsal to the cranialmost portion of the manubrium; however, species vary in the lateral excursion and depth of the vessel. Accessing this vessel is usually successful and, with few exceptions, is the first choice in rodent species.25 However, venipuncture of the cranial vena cava requires sedation or general anesthesia of the animal. To collect a blood sample from the vena cava, place the animal in dorsal recumbency, and prepare the area over the

manubrium for venipuncture. Using a 27- to 25-g needle with a 1-mL or smaller syringe, approach the vessel from the right or left of the manubrium, angling slightly medially and dorsally (Fig. 26-7). The presence of a well-developed clavicle in some species (gerbils, hamsters) may complicate needle position. As you advance the needle, apply gentle negative pressure. Entry of the cranial vena cava will result in a flash of blood. If blood does not flow freely, the needle may have passed through or oblique to the vessel or the bevel may be contacting the vessel wall. Rotate or redirect slightly. The vessel is just under the manubrium; therefore deep needle penetration is not needed or recommended. Potential complications include vessel rupture and exsanguination, but this appears to occur rarely and is likely associated with excessive patient movement. Other site options for collecting blood samples are the lateral saphenous or lateral tail vein in the rat, gerbil, or mouse; the ventral tail artery of the rat; and the tarsal veins of larger rodents.3,17,34 The lateral tail veins are located on both sides of the tail and are superficial; they can be seen easily in young animals and albino rats. Ideally, warm the tail gently to increase blood flow and occlude the veins by placing a tourniquet around the base of the tail; a rubber band and mosquito hemostat are suitable for this purpose. With a needle of appropriate gauge for the species, enter the skin at a shallow angle at a point approximately one-third down the length of the tail. If the initial attempt at collection is unsuccessful, try again at a site closer to the base of the tail. To avoid collapsing the vessel, use a smallvolume syringe to withdraw the sample or collect the blood into a microhematocrit tube as it flows freely from the needle hub. A modified butterfly catheter (i.e., with all but the proximal 5 mm of tubing removed) may also be used.

348

SECTION IV  Small Rodents   Table 26-5     Hematologic and Serum Biochemical Data for Ratsa Value/Analyte

Maleb (n > 150)

Femaleb (n > 150)

Mixed sexc (n = 50)

HEMATOLOGIC TESTING Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x106/μL) White blood cells (x103/μL) Segmented neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Reticulocytes (%) Platelets (x103/μL)

44.2 (38.5-52.0) 15.5 (13.6-17.4) 8.69 (7.62-9.99) 4.28 (1.98-11.06) 22.2 (9.0-49.3) 73.3 (44.7-87.1) 2.0 (1.0-3.6) 1.7 (0.4-4.0) 0.3 (0-0.6) 1.9 (1.4-2.8) 846 (574-1,253)

43.9 (38.5-49.2) 15.4 (13.7-17.2) 8.20 (7.16-9.24) 2.67 (0.96-7.88) 19.3 (8.8-43.8) 75.8 (48.9-88.1) 2.0 (1.0-3.6) 1.9 (0.3-4.7) 0.2 (0-0.7) 2.3 (1.4-3.9) 836 (599-1,144)

33.0-47.0 11.2-15.9 6.4-8.2 4.7-9.4 7.0-32.0 57.0-91.0 2.0-5.0 0-4.0 0-3.0 — 411-626

BIOCHEMICAL ANALYSIS Albumin (g/dL) Alkaline phosphatase (IU/L) Alanine aminotransferase (IU/L) Aspartate aminotransferase (IU/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Chloride (meq/L) Cholesterol (mg/dL) Creatinine (mg/dL) Glucose (mg/dL) Phosphorus (mg/dL) Potassium (mEq/L) Protein, total (g/dL) Sodium (mEq/L) Triglycerides (mg/dL)

4.1 (3.6-4.7) 66 (36-131) 30 (19-48)

4.6 (3.7-5.8) 30 (18-62) 30 (14-64)

2.8-5.3 87-381 36-80

96 (63-175)

101 (64-222)

—

0.10 (0.04-0.20) 16 (11-20) 10.3 (9.1-11.9) 103 (98-106) 59 (37-95) 0.4 (0.3-0.5) 141(106-184) 6.2 (3.6-8.4) 4.6 (3.9-6.1) 6.3 (5.6-7.6) 143 (137-147) 62 (27-160)

0.13 (0.07-0.21) 18 (12-25) 10.6 (9.5-12.1) 102 (97-106) 50 (23-97) 0.4 (0.3-0.6) 119 (89-163) 6.7 (4.5-9.5) 4.1 (3.4-5.1) 6.6 (5.7-8.3) 141 (135-146) 42 (16-175)

0.20-0.70 11-23 5.7-12.4 — — 0.3-0.6 50-135 6.5-12.2 — 5.6-7.4 — —

aValues

are given as means and/or reference intervals. River Crl:WI(Han) rats, 17 weeks of age or older, raised under optimal laboratory conditions. Bayer ADVIA 120 analyzer, HitachiP 800 analyzer. (Available at http://www.criver.com/sitecollection documents/rm_rm_r_wistar_han_clin_lab_parameters_08.pdf) cData courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak:Ektachem) 700XR; Automated analyzer for cell count, hemoglobin, and hematocrit. Differential based on 100-cell count by using blood smear made at the time of sample acquisition. bCharles

The ventral tail artery in the rat is another option.3 The artery courses along the ventromedial aspect of the tail, although it is not as superficial as the lateral tail veins. Place the sedated or anesthetized animal in dorsal recumbency. Use a 22-gauge needle and a 3-mL syringe from which the plunger has been removed. Alternatively, some practitioners prefer to use a 23-gauge butterfly needle with short tubing connected to a 3-mL syringe. Make the first puncture attempt at a point one-third down the tail’s length. Enter the skin at a 20- to 30-degree angle to the tail, with the bevel of the needle facing upward. A perceptible “pop” usually indicates that the artery has been entered; this is quickly followed by filling of the syringe (or butterfly tubing) with blood. The high blood pressure in this vessel negates the need for the negative pressure produced by withdrawal of the plunger. Indeed, the presence of the plunger within the syringe case impairs recognition of correct penetration of the

vessel. After the required volume of blood has been collected, withdraw the needle and apply pressure to the puncture site. Note that more time is required to stop the bleeding from the tail artery than from the tail vein. The lateral saphenous and tarsal veins are often visible coursing across the surface of the lateral leg or tarsus of larger, lighter-colored rodents. These vessels are very small and are best punctured with a needle only and samples collected directly into heparinized hematocrit tubes. A technique for saphenous venipuncture of laboratory rodents has been described that is minimally invasive, does not require anesthesia, can be performed by a single person when combined with the appropriate restraint, and can be repeated at the same location multiple times.17 The procedure does require skill and is used when relatively small quantities of blood are required. For use in a mouse, place the mouse head first in a 50-mL syringe case that has been modified

CHAPTER 26  Basic Anatomy, Physiology, Husbandry, and Clinical Techniques   Table 26-6     Hematologic Data for Hamsters and Gerbilsa Value

Hamsterb

Gerbilc

Hematocrit (%) Hemoglobin (g/dL) Red blood cells (x 106/μL White blood cells (x 103/μL) Neutrophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Platelets (x 103/μL) Mean corpuscular volume (fL) Mean corpuscular hemoglobin (g/dL) Mean corpuscular hemoglobin concentration (g/dL)

45-52 15.2-17.4 6.5-7.5 6.3-8.9 16-26 65-80 0-4 0-2 0-2 300-570c 68-74 19-24 30-34

41-52 12.1-16.9 7.0-10.0 4.3-21.6 5-34 60-95 0-3 0-4 0-1 400-600 — — —

aValues

are given as mean, mean ± 2 SD, or range. courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Automated analyzer for cell counts, hemoglobin, and hematocrit. Differential WBC count based on 100-cell count using blood smear made at the time of sample acquisition. cAverage values from data given in references 12 and 16. Note that reference values may not represent the range for certain populations or strains of animals; for this reason, the values should be interpreted as approximations. bData

  Table 26-7     Biochemical Data for Hamsters and Gerbilsa Analyte Albumin (g/dL) Alanine aminotransferase (IU/L) Alkaline phosphatase (IU/L) Bilirubin, total (mg/dL) Blood urea nitrogen (mg/dL) Calcium (mg/dL) Creatinine (mg/dL) Creatine kinase (CK) Glucose (U/L) Phosphorous (mg/dL) Protein, total (g/dL)

Hamstersb (n = 50)

Gerbilsc (n = 30)

3.5-4.9 22-128 99-186 0.1-0.7 12-26 5.3-12.0 0.4-1.0 — 37-198 3.0-9.9 5.2-7.0

2.5 (2.1-2.9) 91 (56-165) 118 (70-182) — 18 (11-32) — 0.3 (0.2-0.7) 363 (93-752) 91 (24-117) — 5.6 (4.6-6.3)

aValues

are given as reference intervals and means. Samples are mixed plasma and serum. courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. Ortho (Kodak Ektachem) 700 XR. cOrtho Vitros 250 Analyzer. bData

  Table 26-8     Reference Intervals for Serum Protein Electrophoresis for Selected Rodent Speciesa Analyte

Mouse

Rat

Albumin (g/dL) Alpha-1 globulin (g/dL) Alpha-2 globulin (g/dL) Beta globulin (g/dL) Gamma globulin (g/dL) A/G ratio

2.13-3.35 0.25-0.46 0.75-1.10 1.42-1.73 0.16-0.32 0.7-1.1

3.35-4.09 0.40-0.59 0.20-0.57 1.33-1.69 0.12-0.51 1.1-1.6

Data courtesy of Carolyn Cray, Ph.D., University of Miami Miller School of Medicine. aAnalytes measured by Beckman Paragon SPEP II gels in adult animals.

349

350

SECTION IV  Small Rodents   Table 26-9     Urinalysis Reference Values for Gerbils, Hamsters, Mice, and Ratsa Value

Gerbil

Hamster

Mouse

Rat

Urine volume (mL/24 hours) Specific gravity Average pH Protein (mg/dL)

A few drops–4

5.1-8.4

0.5-2.5

13-23

— — —

1.060 8.5 —

1.034 5.0 Males proteinuric

1.022-1.050 5-7 40, increase the flow rate to 300 mL/kg/min).

VENTILATORS Ventilators appropriate for small mammals must cope with the range of tidal volumes in the different species. In very small patients, this may be less than 1 mL; but some giant-breed rabbits might need to be as high as 500 mL. In general the three things that must be controlled by a ventilator are tidal volume, the rate at which the tidal volume is delivered, and the number of breaths per minute. The way in which each of these is controlled varies from ventilator to ventilator.45

CLINICAL TECHNIQUES VASCULAR ACCESS Vascular access is necessary for replacement fluids and to deliver anesthesia and emergency medications. Sites for intravenous catheterization include the cephalic, lateral saphenous, and

CHAPTER 31  Anesthesia, Analgesia, and Sedation of Small Mammals

A

431

B Fig. 31-1  Catheterization of the cephalic vein is the most common intravenous site used (A), but the lateral saphenous, jugular, or femoral veins and the superficial lateral tail veins in the rat can also be used. Catheters can be taped and bandaged for security; additional stability can be provided with tongue depressors for very small limbs. In cases where intravenous catheterization is not possible, an intraosseous catheter may be placed in the tibia (B) or femur for short-term use.

femoral veins in larger mammals and the superficial lateral tail veins in the rat (Fig. 31-1). Sedation may be necessary for intravenous catheter placement. A surgical cut-down procedure under anesthesia is generally necessary for jugular catheterization. Small-bore over-the-needle catheters (≤24 gauge) are used. The catheter site should be aseptically prepared; catheters are then secured with tape and/or sutured in place and bandaged for extra security. Jugular catheters, if left indwelling, require 24-hour monitoring, as fatal hemorrhage can occur if the patient pulls or chews on the catheter and damages the vessel. Many small rodents, even when severely compromised, are intolerant of bandaging material and indwelling catheters and will attempt to remove them. Intraosseous (IO) cannulation can be useful in smaller patients or during cardiovascular collapse (see Fig. 31-1).89 Products that can be used as IO cannulas include 18- to 24-gauge 1- to 1½-in. spinal needles or 18- to 25-gauge 1-in. hypodermic needles, depending upon the animal’s size. The cannula should be long enough to extend through one-third to one-half the length of the medullary cavity. A wire stylet reduces the potential for a bone core. Prepare several hypodermic needles (25- to 18-gauge) with wire stylets (stainless steel suture) and sterilize them for IO catheterization. Common sites for IO cannula placement include the trochanteric fossa of the femur and the tibial crest (see Fig. 31-1, B). Placement is similar to that of a normograde intramedullary pin and requires strict aseptic technique during placement and maintenance. Once the cortex is penetrated, the cannula should advance easily, with little resistance. Flush the cannula with heparinized saline immediately to prevent clotting. Cover the insertion site with an antibiotic ointment and secure the cannula with tape, suture, and a bandage. IO cannulas can remain patent for 72 hours without flushing but should be flushed with heparinized saline twice daily if fluid therapy is not continuous. Complications associated with IO catheterization include penetration of both cortices, failure to properly enter the medullary cavity, and extravasation of fluids with associated pain.89 IO catheters are used primarily for short-term vascular volume expansion until an IV catheter site can be obtained. Many animals become uncomfortable on limbs supporting IO catheters even after short-term placement. IO catheterization is contraindicated in septic patients and those with metabolic bone disease. Osteomyelitis may occur owing to duration or placement of the

IO catheter; administration of alkaline or hypertonic solutions can contribute to osteomyelitis and cause pain.89 Dilute these solutions before administration and flush the catheter with heparinized saline after any drug injection. Arterial catheterization for the evaluation of blood pressure, blood gas tension, and pH can be performed in many species. The most common sites include the pedal arteries in most species, the central auricular artery in the rabbit, and the ventral tail artery in the rat and ferret. The catheters are usually placed percutaneously, but a cut-down procedure is sometimes used.

INTUBATION Endotracheal intubation provides a patent airway, reduces dead space, and facilitates positive-pressure ventilation. Disadvantages include tracheal mucosal trauma, increased airway resistance, and airway occlusion due to mechanical forces or secretions. Increased resistance is of greater importance in very small patients because it is inversely related to the fourth power of the tube radius. For example, decreasing the tube radius from 5 to 3 mm increases the resistance 7-fold, whereas a decrease from 3 to 1 mm increases it more than 80-fold. Increased resistance can be overcome by positive pressure ventilation. Specialized endotracheal tubes and light sources are available to aid intubation of small mammals. The smallest commercial uncuffed tubes have an internal diameter (ID) of 1 mm. However, tubes of less than 2-mm ID are often highly flexible and kink easily. The smallest-diameter cuffed tube is 3-mm ID. Uncuffed tubes do not provide a sealed airway, so clean the oral cavity prior to intubation, elevate the head, and monitor during the procedure. If you are using a cuffed tube, the cuff is carefully inflated with just enough air to prevent leakage when 10 to 15-cm H2O pressure is applied. Very small mammals may be intubated with Teflon IV, red rubber, or urinary catheters (Fig. 31-2, A). Care is taken to ensure that no sharp edges are present at the end. This is achieved by using a small piece of silicone tubing over the end of the catheter (see Fig. 31-2, B,C). Endotracheal tube obstruction is detected by monitoring for a prolonged expiratory phase. Anticholinergics reduce the production of mucus and mucous plug formation but also increase mucous viscosity, making it harder to clear secretions. The use of an endotracheal tube with a Murphy eye decreases the

432

SECTION VI  General Topics

A

B

C Fig. 31-2  A, Commercial endotracheal tubes are useful for larger mammals, but Teflon intravenous, red rubber, and urinary catheters are often necessary to intubate very small mammals. A small piece of silicone tubing can be glued over the end of the intravenous catheter to minimize sharp edges (B, C [inset]).

likelihood of mucous occlusion. Humidification of the gases reduces mucous plug formation. Commercial endotracheal tube humidifiers are available (Humidi-vent Mini Agibeck Product, Hudson RCI, Temecula, CA). Disadvantages of their use include increased dead space and plugging of the filter with secretions. Care must be taken to minimize head and neck movement in intubated patients, as movement of the tube and changes in positioning may induce mucosal trauma. Sublaryngeal tracheal injury, ulceration, and postintubation tracheal strictures have been described in rabbits from several institutions that were intubated with both cuffed and uncuffed endotracheal tubes.38,92 Other factors that predispose to mucosal injury include ventilation technique and endotracheal tube disinfection protocols. Ferret intubation is straightforward and should be performed routinely as in the cat. All pet rabbits over 1-kg body weight can be intubated, but this is more difficult. Rabbits have large incisors, long narrow oral cavities, and thick tongues, making laryngeal visualization more difficult, and laryngospasm is easily induced. There are a number of tracheal intubation techniques that have been advocated for use in rabbits: direct visualization of the larynx with a laryngoscope and intubation, blind intubation with the neck in extension, endoscope-guided intubation, and nasotracheal intubation. Apply topical local anesthetic on the larynx 60 seconds before intubation to decrease laryngospasm. The authors use the blind technique routinely. Guide the tube to the larynx and listen for louder breath sounds, place one drop of 2% lidocaine via tomcat catheter through the endotracheal tube directly on to the larynx, then use the same approach

for intubation (Fig. 31-3). Placement of the tube is confirmed by visualizing the movement of water vapor in the tube, by the rabbit coughing, by auscultation of breath sounds in the lungs, and definitively by attaching a capnograph and seeing a characteristic capnographic trace. Rabbits are obligate nasal breathers and the patency of both nares must be carefully assessed postextubation, especially if the animal has been in dorsal recumbency. Reported complications include postextubation obstruction, respiratory arrest, and tracheal mucosal injury.38,92 Intranasal intubation or catheterization can be used in the rabbit if endotracheal intubation cannot be performed or for complicated oral procedures, such as extensive dental procedures. The ventral meatus of the rabbit’s nasal passage is surprisingly large. A blind intubation technique with a 2.5- to 3-mm ID endotracheal tube is used, but these are often stiff for smaller rabbits; standard IV tubing has a thinner wall and so is more flexible. IV tubing can be cut to the appropriate length and a bevel fashioned on the distal end; the proximal end can be fitted with a 2- to 2.5-mm endotracheal tube adapter. Administer anticholinergics with nasal intubation, as vagally mediated bradycardia can be associated with this procedure. The tube is passed through the ventral meatus with the head held in a normal or slightly extended position. Because of the difficulties of intubating rabbits, newer techniques such as laryngeal mask airways are being evaluated.7,66,111 Rodents also have a long narrow oral cavity, but equipment and techniques for intubating small rodents have been developed.87,117 In guinea pigs and chinchillas, orotracheal intubation is complicated by the fusion of the soft palate to the base of the tongue, creating the palatal ostium, which is highly vascular and easily traumatized (Fig. 31-4, A). Endotracheal tubes of 1.0- to 2.5-mm ID are most often needed for small rodents. Very small rodents may be intubated with catheters or IV tubing as in the rabbit, but occlusion with mucous plugs, due to the small internal diameter of these tubes, occurs frequently. An otoscopic cone, modified pediatric blade, commercially available rodent work stands and intubation packs, or endoscopy can help facilitate intubation. Otoscopic cones that have been modified by removing a section laterally can facilitate visualization of the epiglottis and direct placement of the endotracheal tube. A stylet placed first may help facilitate endotracheal tube placement (see Fig. 31-4, B,C). Endoscopy provides the best visualization of the epiglottis and minimizes trauma during tube placement. Nasal intubation/catheterization has also been performed for guinea pigs and other small rodents with similar issues arising as for rabbits.

PREANESTHETIC MEDICATIONS Parasympatholytics are most commonly used to minimize salivary and bronchial secretions and vagally induced bradyarrhythmias, but they can increase secretion viscosity. Many rabbits have high circulating concentrations of atropine esterases, thus reducing the efficacy of atropine and prompting the use of much higher atropine doses, with redosing as often as every  10 to 15 minutes during bradycardia.44 Large doses of parasympatholytics can alter GI motility in hind-gut fermenters; therefore the lowest doses necessary are used.48 Glycopyrrolate has a longer duration of activity than atropine (Table 31-1). Glycopyrrolate increased heart rates in rats for 240 minutes, versus 30 minutes with atropine.86 The same study demonstrated that 0.1 mg/kg glycopyrrolate administered to rabbits increased heart rate for an equal duration to the rats, but atropine doses ≤2 mg/ kg did not increase heart rate for any duration.86

CHAPTER 31  Anesthesia, Analgesia, and Sedation of Small Mammals

A

C

B

Fig. 31-3  Intubation of the rabbit using the blind technique. Provide fresh oxygen and anesthetic gas through a nasal mask during the procedure. Guide the tube to the larynx, listen for louder breaths and/or visualize water vapor in the tube (A), place one drop of 2% lidocaine via tomcat catheter through the endotracheal tube directly onto the larynx (B), wait 60 seconds, then use the same approach for intubation (C). Placement of the tube is confirmed by visualizing movement of water vapor in the tube, by the rabbit coughing, by auscultation of breath sounds in the lungs, and definitively by attaching a capnograph and seeing a characteristic capnographic trace.

A

B

C Fig. 31-4  In guinea pigs and chinchillas, orotracheal intubation is complicated by the fusion of the soft palate to the base of the tongue, creating the palatal ostium, which is highly vascular and easily traumatized (A). Intubation can be accomplished with the use of an otoscope or endoscope and placement of a stylet (B), then threading the endotracheal tube over the stylet (C).

433

434

  Table 31-1    Injectable Preanesthetic, Sedative, and Tranquilizer Drugs Used in Small Exotic Mammalsa

Glycopyrrolate Sedatives/ tranquilizers Acepromazine

Ferret

Guinea Pig

Chinchilla

Rat

0.2-1.0 SC, IM, IV 0.01-0.02 SC, IM, IV

0.05 SC, IM, IV 0.01-0.02 SC, IM, IV

0.05 SC, IM, 0.05 SC, IM, 0.05 SC, IM, IV IV IV, IP 0.01-0.02 0.01-0.02 SC, 0.02-0.5 SC, SC, IM, IV IM, IV IM, IV25

0.25-1.0 SC, IM

0.1-0.2 SC, IM25 ≥ 1 SC, IM, IV

0.5-1.0 SC, IM25 ≥ 1 SC, IM, IV

Hamster

Gerbil

Mouse

0.04 SC25

0.04 SC25

0.5 IM25

−

0.05 SC, Higher doses may be IM, IP necessary in rabbits 0.02-0.5 SC, − IM25

0.5-1.0 SC, IM ≥ 1 SC, IM, IV54

0.5-2.5 SC, IM, IP25 0.1-1.0 SC, IM, IV, IP25

−

3 IM25

−

−

Atipamezole

≥ 1 SC, IM, IV68,88

Dexmedetomidine

0.05-0.125 IM 0.05-1.0 μg/kg IM, IV

0.04-0.1 IM 0.05 SC 0.05-1.0 μg/kg IM, IV

0.05 SC

0.015-0.5 SC, IP

−

−

Diazepam

0.5-2.0 IV

0.5-2.0 IV

0.5-3.0 IV

0.5-3.0 IV

2.5-5.0 IV, IP

5 IM, IP25

5 IM, IP25

Flumazenil

0.05-0.1 SC, IM, IV 0.1-0.25 SC, IM 1-2 μg/kg IM, IV

0.05-0.1 SC, IM, IV 0.01-0.2 SC, IM25,70 1-2 μg/kg IM, IV

0.05-0.1 SC, 0.05-0.1 SC, IM, IV IM, IV54 0.1 SC 0.1 SC

−

−

−

0.03-0.1 SC, IP25 −

0.1-0.2 IP25

−

0.1-1.0 IM, IV, IP25 0.03-0.1 SC, IP25 −

−

0.5-2.0 SC, IM, IV 2-10 IM, IV

1-2 SC, IM, IV 1.0-2.5 SC, IM, IV; 5 IP25 2-10 IM, IV 1-5 IM, IV, IP

5 IM, IP25

1-5 IM, IP25

2 IM25

Yohimbine

0.2-0.4 IV

0.25-0.5 SC, IM, IV 0.1-0.5 SC, IM25 0.2-1.0 IM, IV

5 IP25

Xylazine

0.25-2.0 SC, IM, IV 2-5 IM, IV25

0.5-1.0 IM, IV

0.5-1.0 IM, IV 0.2 IV; 0.5 IM25

−

−

Medetomidine

Midazolam

Comments

2-5 SC, IM, IP25 ≥ 1 SC, IM, IV, IP

May induce seizures in gerbils 1:1 volume reversal of medetomidine or dexmedetomidine 0.015-0.5 Half the dose of SC, IP medetomidine; Microdoses combined with benzodiazepines and opioids for sedation, induction 3-5 PO, IP25 Significant irritation if given IM − Reversal of benzodiazipines 0.03-0.1 SC, Light sedation, rarely IP25 used alone; variable − effects in guinea pigs Microdoses combined with benzodiazepine and opioid for induction 1-5 SC, Lower doses for IM, IV25 premedication 5-10 IP25 Seldom used 0.5-1.0 IM, IV

Reversal for xylazine

aThese are suggested doses based on published information and the authors’ clinical experience. Species and individual variation in response to a given drug can be uncertain so adjust the dose depending on the clinical response of the animal. All doses are in milligrams per kilogram unless specified otherwise.

SECTION VI  General Topics

Preanesthetics Atropine

Rabbit

CHAPTER 31  Anesthesia, Analgesia, and Sedation of Small Mammals

INJECTABLE MEDICATIONS A considerable increase in the use of injectable anesthetic combinations in small exotic mammals has recently been documented.* Certain breeds or even strains of small mammals have differing responses to injectable anesthetics.5 There is little published work evaluating different injectable combinations for common surgical procedures in pet animals of differing age, sex, breed, and health status in the clinical setting,40,88 and so use caution when extrapolating injectable doses from laboratory animal studies. Using reversible injectables provides better control of anesthetic depth, less hypothermia, less cardiopulmonary depression, and shorter recovery times.54-56,98 Muscle necrosis and cutaneous reactions can occur in any patient administered IM injections, depending on the drug formulation or volume delivered. Drugs are often not uniformly absorbed when administered SC, resulting in erratic and unpredictable anesthesia. The intraperitoneal (IP) route of injection is commonly used in laboratory medicine, but errors during administration (intravisceral, SC, or intra-adipose tissue) can result in organ damage or delayed onset of action. Injections are usually performed into the lower left quadrant of the abdomen with the rodent restrained in dorsal recumbency and the cranial end of the rodent directed downward. Supplemental oxygen and assisted ventilation should always be available when injectable anesthetics are used, regardless of their route of administration.

SEDATIVES AND TRANQUILIZERS Acepromazine In ferrets, acepromazine provides adequate sedation for simple procedures such as ear cleaning,71 however, one author discourages its use in ferrets owing to its vasodilatory effects (see Table 31-1).64 Acepromazine has been used extensively in rabbits and rodents and reported doses vary widely.4,48,120,122 The peak effect in rabbits is often not seen for 30 to 40 minutes, even when given IV. It is used in laboratory rabbits specifically for blood collection because of the drug’s vasodilatory effects. Vasodilation occurs at very low doses of acepromazine, therefore low doses will not avoid this effect. Animals that are hypovolemic, anemic, or hypotensive before anesthesia should not receive acepromazine. Acepromazine has also been shown to decrease tear production in rabbits.35 Acepromazine is not recommended for use in gerbils because it may lower the seizure threshold.49

Benzodiazepines Diazepam can be administered PO or IV, but if used IV, the patient must be monitored for hypotension caused by the propylene glycol found in many diazepam formulations (see Table 31-1). IM and SC absorption is erratic because of the solution’s high osmolality, which is due to an increased pH with this carrier. Because parenteral formulations of diazepam are available, large volumes must often be given. Transient lameness after IM injection has been reported in ferrets.72,74 Diazepam can be used alone in rabbits to provide preoperative anxiolysis and sedation,15 but it is most commonly used in combination with other drugs to enhance their activity or decrease muscle rigidity. Midazolam is a short-acting benzodiazepine that is watersoluble, so it can be given IM (see Table 31-1). Midazolam has

*References 4, 40, 53-55, 72, 74, 88, 120, 122.

435

been used as a sole sedative for minor, nonpainful procedures in rabbits and rodents. Lower doses are most often used for preanesthetic sedation. Flumazenil will reverse midazolam, but because its half31-life is shorter than that of midazolam, resedation may occur. Titrate the flumazenil dose to avoid the reversal of the beneficial anxiolysis, sedation, and muscle relaxation. Benzodiazepines coupled with opioid medications provide sedation and preoperative analgesia.

Alpha-2 Agonists Xylazine, medetomidine, and now dexmedetomidine are the most commonly used alpha-2 adrenergic receptor agonists in small exotic mammals (see Table 31-1). Their major advantages are that they provide good muscle relaxation and can be reversed with yohimbine, atipamezole, or tolazoline (see Table 31-1). If all analgesic components are reversed, postoperative analgesia should be provided before reversal of anesthesia. These drugs can have significant cardiopulmonary effects, including respiratory depression, second-degree heart block, bradyarrhythmias, and increased sensitivity to catecholamine-induced cardiac arrhythmias (xylazine).53,54 Because of the many adverse effects reported for this class of drugs, their use is not recommended in small exotic mammal patients with evidence of cardiopulmonary compromise. Supplemental oxygen and assisted ventilation should always be available when these drugs are being used. Xylazine/ketamine given over multiple anesthetic episodes and detomidine alone or in combination with ketamine or diazepam have been associated with myocardial necrosis and fibrosis in rabbits.61,80 Significant breed and strain differences in response to medetomidine sedation doses have been reported for rabbits and rats.5,6,48 In rabbits, ketamine/medetomidine allowed for more rapid intubation, a greater isoflurane-sparing effect, and less esophageal temperature loss than with ketamine/ midazolam, but the rabbits thus treated were more prone to laryngospasm.40 Medetomidine/ketamine provided better quality and duration of surgical anesthesia (38.7 ± 30.0 minutes) in healthy rabbits than medetomidine/midazolam/fentanyl (MMF). Arterial pH and PaO2 were significantly decreased in both groups and apnea occurred postintubation with MMF.53 Atipamezole (10%-20% of calculated dose) may reduce these adverse effects without reversing their anesthetic and analgesic effects, but supplemental oxygen must be available. The effects of alpha-2 agonists with ketamine seem to be less uniform in guinea pigs, and they may not become sufficiently anesthetized.96 Chinchillas administered MMF IM had wellcontrolled anesthesia for 1.5 hours, but the heart rate (HR) and respiratory rate (RR) were decreased and recoveries were prolonged without reversal.54 Chinchillas receiving medetomidine/ ketamine had greater HR and RR depression, and recovery was longer than with MMF combinations (Table 31-2).54 Alpha-2 agonists are combined with ketamine for IP injection for surgical anesthesia with good muscle relaxation in rats and mice.4,96

INDUCTION AND MAINTENANCE OF ANESTHESIA Preoxygenation should be performed routinely, and always when there is potential for hypoxemia. Preoxygenation can produce an oxygen reservoir and the benefits are achieved in ≤1 minute of high inspired oxygen concentration in the healthy patient but may take ≥5 minutes in a patient with compromised respiratory function. Preoxygenation is accomplished with an

436

Rabbit

Ferret

Guinea Pig

Chinchilla

Rat

Hamster

Gerbil

Mouse

Comments

Injectable Anesthetics/Analgesics 1-2 IP20

Etomidate

1-2 IV

1-2 IV

Ketamine

5-50 SC, IM, IV25

5-50 SC, IM, IV70

5-40 SC, IM, IV

5-40 IM, IV

10-40 IM, IV; 50-100 IP25

50-100 IP25

100-200 IM, IV25

Ketamine (K)/ diazepam (D)

10-25 K/1-5 D IV25,48

10-25 K/1-3 D IV25,70

20-30 K/1-2 D IV

40-100 K/3-5 D IP33

70 K/2 D IP25

50 K/5 D25 100 K/5 D IP25,33

Ketamine (K)/ diazepam (D)/ butorphanol (B) Ketamine (K)/ medetomidine (M) Ketamine (K)/ medetomidine (M)/butorphanol (B) Ketamine (K)/ midazolam (Mi) Induction Anesthesia

−

15 K/3D/0.2 B IM, IV70

100 K/5 D IM25; 20-30 K/1-2 D IV −

−

−

−

−

−

4-8 K/0.085-35 K/0.25-0.5 M 0.1 M IM17,39,40,50,53,68,88 IM25,70 5 K/0.08 5-15 K/0.1-0.5 M/0.1-0.2 M/0.4-0.5 B SC, B IM70 IM50

5-40 K/0.050.5 M IM25

5 K/0.06 M IM54

100 K/0.25 M IP25

75 K/0.5 M IV25

−

−

45-75 K/0.30.5 M IM, IP42,51 −

−

−

50-75 K/0.11.0 M IP25,42 −

5 K/0.25-1.0 Mi IM, IV

5 K/0.1-0.3 Mi IM, IV

5 K/0.5 Mi IM, IV

5 K/0.5 Mi IM, IV

−

−

−

−

10-40 K/1-3 Mi IM, IV39,40,48

5-10 K/0.25- 20-50 K/0.50.5 Mi IM, 5.0 Mi IM, IV70 IV 25 K/1-2 X 20-40 K/1-2 IM25 X IM

20-50 K/0.55.0 Mi IM, IV 20-40 K/1-2 X IM

40-150 K/3.0- − 5.0 Mi IV, IP25 40-100 K/5-10 200 K/10 X X IM, IP25,33 IP25

−

−

0.05 M/0.02 − F/1 Mi IM54

40-150 K/3.0- Lighter anesthesia at 5.0 Mi IP25 lower dosages of ketamine 35-100 K/2.5- − 15.0 X IP25,42 − Must be prepared to intubate because apnea, respiratory depression common; complete reversal with atipamezole, flumazenil, naloxone

Ketamine (K)/ xylaxine (X)

10-50 K/3-5 X IM17,53

Medetomidine (M)/ 0.25 M/0.02 F/1 Mi fentanyl (F)/ IM53 midazolam (Mi)

−

−

50 K/2 X IP25 −

100-200 IM, IV25

Administer 15-20 minutes after benzodiazepine Lower doses for induction; can cause muscle necrosis in guinea pigs Poor analgesia Provided 20-25 minutes anesthesia in ferrets − −

−

SECTION VI  General Topics

  Table 31-2    Injectable Anesthetic Drugs and Drug Combinations Used in Small Exotic Mammalsa

3-10 IV10

−

−

3-25 IV25

3-10 IM, IV70 −

−

40-80 IV, IP25,96,104

50-80 IP25

−

40-80 IP25,96

Bupivacaine (local,