Manual of Exotic Pet Practice

11830 Westline Industrial Drive St. Louis, Missouri 63146

MANUAL OF EXOTIC PET PRACTICE Copyright © 2009 by Saunders, an imprint of Elsevier Inc.

ISBN: 978-1-4160-0119-5

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. Permissions may be sought directly from Elsevier’s Rights Department: phone: (+1) 215 239 3804 (US) or (+44) 1865 843830 (UK); fax: (+44) 1865 853333; e-mail: [email protected]. You may also complete your request on-line via the Elsevier website at http://www.elsevier.com/permissions.

Notice Knowledge and best practice in this field are constantly changing. As new research and experience broaden our knowledge, changes in practice, treatment and drug therapy may become necessary or appropriate. 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 the practitioner, relying on their own experience and knowledge of the patient, 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 assume any liability for any injury and/or damage to persons or property arising out of or related to any use of the material contained in this book. The Publisher Library of Congress Control Number: 2007925165

Vice President and Publisher: Linda Duncan Publisher: Penny Rudolph Developmental Editor: Shelly Stringer Publishing Services Manager: Pat Joiner-Myers Senior Project Manager: Gena Magouirk Design Direction: Julia Dummitt

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Contributors Maya Bewig, DVM Associate Grey Wolf Veterinary Hospital Sequim, Washington, USA Wildlife Deborah Carboni, DVM, MS Ontario Veterinary College Guelph, Ontario, Canada Marsupials M. Camille Harris, DVM, MS, BS PhD Student (Ecology of Infectious Diseases) Department of Biological Sciences Virginia Polytechnic Institute and State University Blacksburg, Virginia, USA Hamsters and Gerbils J. Jill Heatley, DVM, MS, Dip. ABVP (Avian) Clinical Associate Professor, Zoological Medicine Department of Clinical Sciences College of Veterinary Medicine and Biomedical Sciences Texas A&M University College Station, Texas, USA Hamsters and Gerbils Hedgehogs Megan Kirchgessner, DVM Clinical Veterinarian New England Wildlife Center South Weymouth, Massachusetts, USA Chelonians Stephen M. Miller, DVM General Curator North Carolina Zoo Asheboro, North Carolina, USA

Mark A. Mitchell, DVM, MS, PhD Associate Professor, Zoological Medicine Department of Veterinary Clinical Medicine College of Veterinary Medicine University of Illinois Urbana, Illinois, USA History of Exotic Pets Preparing Your Hospital for Exotic Pets Invertebrates Ornamental Fish Snakes Chelonians Rabbits Chinchillas Wildlife Natalie Mylniczenko, DVM, MS Associate Veterinarian Chicago Zoological Society Brookfield Zoo Brookfield, Illinois, USA Amphibians Javier Nevarez, DVM, PhD Assistant Professor, Zoological Medicine Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University Baton Rouge, Louisiana, USA Crocodilians Lizards Shannon M. Riggs, DVM Staff Veterinarian International Bird Rescue and Research Center Fairfield, California, USA Guinea Pigs Chinchillas

Ornamental Fish

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vi Thomas N. Tully, Jr., DVM, MS, Dip. ABVP (Avian), ECAMS Professor, Zoological Medicine Department of Veterinary Clinical Sciences School of Veterinary Medicine Louisiana State University Baton Rouge, Louisiana, USA Birds Marsupials Mice and Rats Kristine M. Vennen, DVM Clinical Veterinarian Loftin Veterinary Clinic Youngsville, Louisana, USA

MANUAL OF EXOTIC PET PRACTICE

Tiffany M. Wolf, DVM Clinical Veterinarian The Minnesota Zoo Apple Valley, Minnesota, USA Ferrets Trevor Zachariah, DVM, MS Veterinary Resident Chicago Zoological and Aquatic Animal Residency Program University of Illinois Urbana, Illinois, USA Invertebrates

Rabbits

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Preface Exotic companion animals are increasingly being recognized as a significant area of interest within veterinary medicine. For decades, veterinarians have been asked to consult on cases involving ornamental fish, rabbits, small rodents, birds, reptiles, or even an occasional spider or hermit crab. Many veterinarians have found it difficult to treat and consult on these cases because of a lack of published information; many are forced to make recommendations that are based on subjective or anecdotal information. Fortunately, the practice of exotic pet medicine has made great strides in the past two decades. Although there remains much work to be done, the current knowledge base regarding nontraditional pets has been improved through scientific discovery and communication among veterinarians. Historically, much of the exotic pet literature was based on an understanding of what was done with domestic species. Today, veterinarians have many more resources for information on the species mentioned in this book, including several peer-reviewed journals publishing exotic animal–specific articles that provide a scientific foundation to better treat and manage these species in captivity. It is

from an expanded information base that the editors and authors have compiled this text. This textbook is one of the first of its kind to include sections on all of the major exotic animal groups. As the editors of this text, we felt that there was a strong need for a general exotic pet textbook that could be used by veterinarians to manage any exotic animal that came their way. There is a large percentage of the veterinary profession practicing small or mixed animal veterinary medicine that has a small to moderate caseload of exotic pets. It is for this group of veterinarians that this text was written. There are certainly other excellent textbooks that go into extensive detail on the needs of each group of exotic pet animals and would certainly be beneficial to the veterinary practitioner. Unfortunately, to purchase each textbook on its own would require a significant monetary investment. Although initially intended as a general reference for those individuals looking for a general exotic pet text, this text will become an important reference for those individuals with more exotic pet experience.

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Acknowledgments We wish to thank each of the authors for their contributions to this text. Because of the spectrum of coverage this text provides, it would be difficult, if not impossible, to complete such a textbook without their contributions. We would also like to thank Ms. Verna Serra for her invaluable administrative assistance during the preparation of this text. We are also grateful

to our publisher, Elsevier, for providing us an opportunity to bring to fruition a textbook that will provide an invaluable resource to so many in our profession. Our primary contact at Elsevier, Ms. Shelly Stringer, was most helpful with keeping us on track and guiding this much-needed book to completion.

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Dedication

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n life, it is important to find that which inspires you. For me, my inspiration comes from my wife, Lorrie, and daughter, Mary Yansheng. Lorrie and Mary, it is your love that serves as my source of energy and ensures that I can succeed at whatever endeavor I pursue. I love you both. To my coeditor, Thomas N. Tully, thank you for your mentorship, friendship, and guidance during my tenure at LSU. You will always be my chief. (MAM)

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dedicate this book with love to my parents, Donna and Thomas N. Tully, Sr. Their guidance and support have been invaluable assets to my professional development. I wish to extend a heartfelt thank-you to my wife, Susie, and two daughters, Claudia and Fiona, for their day-to-day patience and inspiration. To my coeditor, Mark Mitchell, thank you for being the best colleague, friend, and office mate one could ask for over the past 10 years. (TNT)

Mark A. Mitchell

C H A P T E R

1

HISTORY OF EXOTIC PETS WHAT IS AN EXOTIC PET? The word exotic is used as an adjective to describe many different things in society. Generally, this term indicates something unique, dangerous, or exciting. In Merriam-Webster’s Collegiate Dictionary, 11th edition,1 the definition of exotic includes: “1. introduced from another country: not native to the place where found, 2. (Archaic) foreign, alien, and 3. strikingly, excitingly, or mysteriously different or unusual.” Any of these definitions are applicable to the species of interest in this book. However, there are other adjectives commonly used to describe exotic animals too, including nontraditional or nondomestic. Selecting the best adjective to describe the species addressed in this text can be difficult. For example, to call all of these pets exotic may not be correct, as certain species, such as box turtles (Terrapene carolina) or red-eared sliders (Trachemys scripta elegans), are kept as pets in the United States but are native to large regions of the country. The use of the term nondomestic or nontraditional would not necessarily be correct for certain species either, such as ferrets, as these animals have been associated with humans for over 2,300 years.2 For purposes of simplicity, the editors have selected the adjective exotic to describe the species described in the text because they are certainly considered by most to fit the third dictionary definition for the term exotic: “strikingly, excitingly, or mysteriously different or unusual.” There are those nonexotic (or traditional, domestic) species that the public acknowledges as common or usual, including the dog, cat, cow, horse, sheep, goat, pig, or chicken, and then there are those that are anything but common or usual, such as the goliath bird-eating spider (Theraphosa blondi), freshwater angelfish (Pterophyllum scalare), blue poison dart frog (Dendrobates azureus), blue-tongued

skink (Tiliqua scincoides), hawk-headed parrot (Deroptyus accipitrinus), or chinchilla (Chinchilla lanigera).

History of Exotic Pets Exotic pets have long held the interest of humans. During the early and mid-20th century, it was not uncommon for newly imported reptiles, birds, or mammals to stir the imagination of the public. The following is a brief historical review of how some exotic animals gained in popularity. Ornamental fish represent one of the oldest groups of exotic pets. Evidence suggests that the Sumerians were the first to keep fish in captivity (2500 bc), but it was for food.3 The Egyptians and Romans were likely the first groups to keep fish as something more than just a food source. However, it was the Chinese (Sung Dynasty: 960-1279) who were the first to actively keep and breed fish for their aesthetics. Goldfish were the first fish to be actively maintained for this reason. It was not until the 17th century that these ornamentals made their way to the Western world (Europe). A major problem encountered by those who did not have regular access to natural spring water was the inability to maintain the health of the fish. Losses were likely great in those days from elevated ammonia and nitrite levels in the water. Issues of water quality were first addressed by Robert Warrington in the 19th century. His theory for a successful aquarium was to use plants to produce oxygen for fish and snails to eat the detritus. It was not until the early and mid-20th century that the importance of aeration and filtration was acknowledged. Ornamental fish arrived in the United States in the late 19th century/early 20th century. The ornamental fish hobby grew with the advent of commercial travel. Fish could be moved globally by ship, railroad, or

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2 plane. The cost of the fish and equipment, although expensive, finally became affordable after the 1940s. Over the past 2 to 3 decades, significant advances have been made in filtration techniques, water quality standards, and fish nutrition. The ornamental fish industry remains an important contributor to the overall pet market; however, to date, veterinarians have not developed any significant inroads into this field. Reptiles have only recently become popular as pets and something more than a “dime store” fancy. From the 1940s to the 1970s, the primary reptiles being sold in the United States were native species, such as the red-eared slider turtle and green anole (Anolis carolinensis). In the earlier decades of the 20th century, hatchling turtles were collected from wild nests and offered for sale. Starting in the 1950s and 1960s, turtle farming became popular in the southern states (e.g., Louisiana). Green anoles and other native reptiles were also routinely captured from the wild for the pet retail trade. After the U.S. Food and Drug Administration instituted the 1975 regulation restricting the interstate and intrastate sales of chelonians under 10.2 cm (4″), turtle farmers began exporting the turtles. During the latter half of the 20th century, reptiles were imported from Australia, Africa, South and Central America, and Asia; however, it was not until the 1980s and 1990s that a real explosion in the pet reptile trade occurred. Much of the initial trade revolved around green iguanas, although boids (e.g., boa constrictor, Boa constrictor) and pythons (e.g., ball python, Python regius) were also being imported in large numbers. In 1997 alone, more than 566,000 green iguanas, 94,000 ball pythons, and 29,000 boa constrictors were imported.4 During the 20th and into the 21st century, there has been a move toward captive propagation of reptiles. Many of the “designer” reptiles that are currently available, including corn snakes (Elaphe guttata), leopard geckos (Eublepharis macularius), bearded dragons (Pogona vitticeps), and ball pythons, have been derived from the establishment of specialized genetic lines. With the current high dollar value of many of these animals (e.g., $5,000-$30,000 for ball pythons), individuals are often interested in obtaining veterinary care for their pets. The current status of the reptile trade in the United States is based on a combination of both wild-caught and captively propagated species. Wild-caught animals continue to be primarily distributed through retail pet stores, where the client’s knowledge regarding the care of these animals is limited, whereas captive animals, especially high dollar animals, are trading hands by more experienced herpetoculturists. Psittacines may represent the class of exotics that have been kept the longest in captivity. Records in Egypt suggest birds were kept in captivity for purposes in addition to food since 4000 bc.5 With the advent of open water sailing during the 15th to 18th centuries, the movement of exotic birds became more commonplace. Many of the birds considered common, such as canaries (Serinus canaria), parakeets (Melopsittacus undulatus), and zebra finches (Poephila guttata), have been bred in captivity since the 18th and 19th centuries. In the United States, aviculture became very popular with the turn of the century. With the advent of flight, it became even easier to transport birds across the country. Through most of the 20th

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MANUAL OF EXOTIC PET PRACTICE

century, a large proportion of birds being sold in the United States were wild-caught animals. These could be distinguished from captive-born birds by their open leg band. Fortunately, over the past 2 decades, the number of birds being imported has declined, and the majority of the psittacines offered for sale are captive-born (closed leg band). Ferrets are one of the exotic species of animals that have a long and documented history with human civilization. Originally brought into captivity around 350 bc, these animals have held many roles in captivity, including hunting partner, vermin control, and companion animal. Ferrets are thought to have been introduced into the United States during the 1700s. These animals would have been brought over during the great migration to the New World. Their value for hunting and mousing certainly would have gained them favor among their caretakers. Today, these animals have a lifestyle that is very different from their past. Ferrets, no longer considered “working animals,” spend the majority of their day slumbering and serve as companion animals. In certain parts of the world, such as the United Kingdom, ferrets remain active working animals, assisting hunters with the capture of rabbits. Domestic rabbits, like ferrets, have been associated with human civilization for over 1000 years. Oryctolagus cuniculus, the domestic rabbit, was originally found on the Iberian Peninsula at the end of the Pleistocene era.6 Historical records suggest that it was the Phoenicians who first exported rabbits to Spain in 1100 bc.7 However, it was a group of French monks that were credited with domesticating and selectively breeding the rabbits that are consistent with the animals known today. Rabbits were prized by the Romans and English. There is little documentation regarding the landing of domestic rabbits in the United States before the 20th century. It is likely that animals were imported from Europe before this time, but not in any great numbers. The Belgian hare was the first rabbit to catch the American public’s attention. This large breed was prized not only because of its aesthetics but also because it could provide meat and fur. The first organized rabbit association, the National Pet Stock Association, was formed in 1910. The organization, which has since changed its name to the American Rabbit Breeders Association, has over 37,000 members. Rabbits remain popular today in the United States for production of meat and fur, as research animals, and as companion pets. Chinchillas remain exotic animals of interest in the United States. These animals, like ferrets, have served many different roles since being acknowledged by humans, including providing fur and meat, acting as research models for auditory research, and serving as companion pets. Chinchillas originallly served as a prey species, providing meat and fur to indigenous peoples of the Andes mountains. An international trade for chinchilla fur was founded in the 1500s, but by the late 1800s native populations had been decimated.8 Chinchillas were originally introduced into the United States in 1923 by Mathias F. Chapman. This founder stock comprised 11 animals, 3 of which were females. Although chinchillas are still being raised for fur production, they are now very popular as pets.

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Chapter 1

WHY ARE EXOTIC PETS BECOMING MAINSTREAM? A Survey of Exotic Pet Ownership For veterinarians to be successful, they must have patients. When a veterinarian attempts to establish a site for a veterinary hospital, it should be based on the potential to have access to a sufficiently sized patient population. In some areas of the United States, animal populations can be estimated based on rabies vaccine registrations for dogs and cats or Coggin’s test results for horses. Unfortunately, this is more difficult to do for exotic species. With the exception of vaccinating ferrets for rabies, there is no community-based method for identifying the number of exotic pets in a geographical area. Because of this, veterinarians must rely on larger, national surveys to identify the size of their potential caseload. The American Veterinary Medical Association (AVMA) and the American Pet Product Manufacturer’s Association (APPMA) are two national organizations that routinely survey the pet population landscape in the United States. Their surveys provide an estimate of the number of households that maintain companion pets in the United States and also attempt to estimate the number of animals in a given household. This information is derived primarily by these two organizations to provide their respective industries with the information they need to develop future strategies regarding the sale and care of pets in the United States. The last published report by the AVMA was in 2002.9 The results of the survey suggested that exotic pets are not uncommon in U.S. households. Interestingly, 60% of pet households owned more than one type of pet. As many as 65.4% and 49.4% of bird owners were likely to have dogs and cats, respectively. This is vital information for veterinarians attempting to decide whether to incorporate additional species (e.g., birds) into their practices, as many pet owners would likely prefer the convenience of visiting a single veterinarian rather than two or more veterinarians. Of all the exotic species included in the survey, fish represented the largest group being kept as pets (49.2 million). This is interesting because veterinarians play a limited role in providing health care for ornamental fish. Birds represented the second largest group of exotic animals being kept as pets (approximately 10 million); however, according to the survey, the number of birds kept as pets had decreased by almost 20% compared to the number found 5 years earlier (1996). Rabbits represented the third largest group of exotic

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History of Exotic Pets

animals being kept as pets (4.8 million). Reptiles (lizards, snakes, chelonians, and others) accounted for approximtately 2.9 million animals in the survey. If all rodents were combined (e.g., hamsters, gerbils, guinea pigs, other rodents), then the total number of these animals being kept as pets was approximately 2.6 million. Ferrets represented the smallest number of animals being kept as pets (approximately 1 million); however, these animals are illegal to own in California, and the survey probably underestimated the true number of ferrets in the United States. When attempting to derive estimates of a pet animal population from published surveys, it is important to recognize that these types of surveys can underestimate the actual number of cases. For example, the 2001 AVMA survey suggested that approximately 2.9 million reptiles were being kept as pets in the United States, whereas the 2000 APPMA survey suggested the number might be as high as 9 million.10 This represents a 68% discrepancy in the number of proposed animals. With over 1 million animals being imported annually, it would suggest that more reptiles are being kept as pets than the AVMA survey suggests. Although it is arguable that many of the imported animals succumb during transport, these numbers do not account for the thousands of captive-born reptiles being sold in the U.S. market. Amphibians were also not included in the AVMA survey; however, they would not be expected to account for such a discrepancy between the studies.

REFERENCES 1. Merriam-Webster’s Collegiate Dictionary, ed 11, Springfield, Mass, 2003, Merriam-Webster. 2. Fox JG: Biology and Diseases of Ferrets, ed 2, Baltimore, 1998, Lippincott Williams & Wilkins. 3. Aquarium, from Wikipedia, http://en.wikipedia.org/wiki/Aquarium, May 20, 2006. 4. U.S. Fish and Wildlife Service, Law Enforcement Management Information System, from www.fws.gov, 1997. 5. Rutgers A, Norris KA: Encyclopedia of Aviculture, vol 3, Dorset, UK, 1977, Blandford. 6. Grzimek: Grzimek’s Encyclopedia of Mammals, vol 4, New York, 1990, McGraw-Hill. 7. Brown M: Exhibition and Pet Rabbits, Surrey, UK, 1982, Spur. 8. Chinchilla, from Wikipedia, http://en.wikipedia.org/wiki/Chinchilla, May 20, 2006. 9. Wise JK, Heathcott BL, Gonzalez ML: Results of the AVMA survey on companion animal ownership in US pet-owning households, J Am Vet Med Assoc 221(11):1572-1573, 2002. 10. American Pet Product Manufacturer’s Association: 2001-2002 APPMA National Pet Owners Survey, Greenwich, Conn, 2001, APPMA.

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Mark A. Mitchell

C H A P T E R

2

PREPARING YOUR HOSPITAL FOR EXOTIC PETS ADVANTAGES AND DISADVANTAGES OF INCOROPORATING EXOTIC PETS INTO A VETERINARY PRACTICE When considering whether to incorporate exotic species into a veterinary practice, veterinarians should first identify the potential advantages and disadvantages associated with such a change. Incorporating exotic pets into a veterinary practice can have multiple benefits (e.g., provide new challenges for veterinarians, provide clients a single hospital resource for all [most] of their pets, and increase hospital revenue). The potential disadvantages for adding these species to a practice include the additional time that will be required for acquiring continuing education, additional monies required to purchase equipment and literary resources for the veterinary library, and the time required to publicize the practice’s entrance into the field of exotic pet medicine. Veterinarians are an ambitious lot. The pursuit of a veterinary professional degree requires commitment, intelligence, and ambition. Because these are traits that are inherent to veterinarians, it is important that they continually challenge themselves to satisfy their intellectual and emotional needs. Upon graduation, veterinarians may find veterinary medicine challenging because of the “newness” of it all; however, after this “newness” wears off, many veterinarians find themselves not satisfying their desires for intellectual challenge. Working with exotic pets can provide veterinarians with this much needed challenge. Although mastering the majority of issues regarding dogs and cats would be feasible in a career, it is unlikely that anyone could become highly competent working with the thousands of exotic species that could be presented

to the veterinarian. Although many veterinarians call what they do “veterinary practice,” it could not be any more true than with exotic animals. Some might argue that attempting to work with so many species would be difficult and that veterinarians should focus more on the types of animals they already work with. It is important that veterinarians understand their limitations. For example, if a veterinarian has limited orthopedic surgical experience, he or she should not attempt to place a dynamic compression plate on a femoral fracture in a poodle. The same logic applies in the case of exotic species too. The 2001 American Veterinary Medical Association (AVMA) survey confirms that the majority of U.S. households with pets have multiple pets. This is an important consideration for veterinarians. Once a veterinary-client releationship has been established, it is easier for the client to maintain it for all of his or her pets. Using a single veterinary resource would be preferred by most clients, for both emotional reasons and convenience. Many clients would not hesitate to bring their dog or cat to the veterinarian for an annual examination but are unaware that their exotic pets need the same level of care. If the veterinarian is not making these types of recommendations, then the client’s exotic pets will not be provided the same level of health care. The inclusion of exotic pets into a veterinary practice can also be financially rewarding. Yes, there are those clients that will remind the veterinarian that their pet hamster can be replaced for $5, but there are also those who will pursue any level of care for their $5 hamster because of the human-animal bond they have developed with their pet. The same mentality is still held for “outdoor” cats and “pound” puppies. Until there is a real recognition of the responsibility associated with

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Chapter 2

5

Preparing Your Hospital for Exotic Pets

TABLE 2-1

Professional Exotic Species Associations

Organization

Contact Information

Journals

American Association of Zoo Veterinarians

http://www.aazv.org/

Association of Avian Veterinarians

http://www.aav.org/

Association of Exotic Mammal Veterinarians

http://www.aemv.org/

Association of Reptilian and Amphibian Veterinarians International Association for Aquatic Animal Medicine National Wildlife Rehabilitator’s Association Wildlife Disease Association

http://www.arav.org/ http://www.iaaam.org/

Journal of Zoo and Wildlife Medicine (Quarterly) Journal of Avian Medicine and Surgery (Quarterly) Journal of Exotic Pet Medicine (Quarterly) Journal of Herpetological Medicine and Surgery (Quarterly) IAAAM News (Quarterly)

http://www.nwrawildlife.org/ http://www.wildlifedisease.org/

Wildlife Rehabilitation Bulletin Journal of Wildlife Diseases (Quarterly)

owning a pet, there will always be clients (e.g., dog, cat, horse, and exotic) who believe veterinary care is expensive and unnecessary. I will not focus on them in this chapter. Instead, I will focus on the fact that being a thorough diagnostician can be financially rewarding. Exotic animals have evolved to mask their illness to avoid predation. Therefore, to fully ascertain the status of an exotic pet requires the pursuit of diagnostic tests. A physcial examination alone will not suffice. At minimum, a database for exotic pets should include a complete blood count, plasma chemistry, urinalysis (mammals), and fecal examination. In many cases, radiographs, ultrasound, or endoscopy are also required. Veterinarians that successfully treat exotic pets are able to explain to their clients the necessity of these diagnostics for evaluating their patient’s condition while, at the same time, generating income for their practice. Working with exotic pets requires more “client time” than does working with domestic pets. Often it is important to spend additional time with the client, covering husbandry and nutrition issues that would be unnecessary with a dog or cat client. Because of the additional time required to work with these animals, veterinarians should institute fees commensurate with the time allocated to the examination and consultation. I, along with the other editor of this text, believe that the potential disadvantages associated with incorporating exotic pets into a practice are truly negligible. One of the potential disadvantages mentioned previously was the need to acquire continuing education and literature to expand the veterinarian’s medical knowledge base. With the large number of available national and international continuing education opportunities (e.g., Central Veterinary Conference, Kansas City, MO; North American Veterinary Conference, Orlando, FL; Western Veterinary Conference, Las Vegas, NV) that incorporate both domestic and exotic species, the potential to obtain the necessary knowledge and hands-on laboratories to expand clinical skills in exotic pets while maintaining learning opportunities with domestic pets is great. Those interested in further expanding their opportunities to learn exotic pet medicine and remaining current on new trends and scientific

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Figure 2-1 An Ellman radiosurgery unit can be used to perform surgery on exotic and domestic species. I find these units invaluable for hemostasis in all patients, regardless of size.

discovery in the field of exotic pet medicine can become members of the various exotic species professional veterinarian organizations (Table 2-1). The argument that the inclusion of exotic pets into a practice will lead to significant expenses for equipment is not valid for high-quality small animal practices. The majority of the equipment that is used to treat and diagnose exotic pets is also invaluable for domestic cases. For example, a radiosurgery unit (Ellman International, Inc., Oceanside, NY) (Figure 2-1) is used extensively in my practice to manage hemorrhage and minimize tissue trauma for small patients. This same unit can, and in many cases should, be used by those working with domestic pets for the same inherent reasons. The VetScan (Abaxis Inc., Union City, CA) (Figure 2-2) chemistry analyzer provides the exotic animal veterinarian with the ability to analyze chemistries from 100 μL of whole blood or plasma. This is important when dealing with small patients, such as budgerigars or geckos. However, those of us who work with (or have worked with) domestic pets can certainly remember

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6

MANUAL OF EXOTIC PET PRACTICE

ing these individuals with information about exotic pet care will help ensure that the pet retail employees make the appropriate recommendations to potential pet owners. Historically, veterinarians do not visit with a pet owner until after they have purchased the animal and the equipment. In many cases, veterinarians may have to educate the client on the incorrect purchases they made, which for many, is upsetting. A proactive approach of training pet retailers can prevent unnecessary animosity between the veterinarian and the pet retailer. By establishing a solid relationship with the pet retailer, the veterinarian can expect to receive referrals, which will expand his or her caseload and be financially rewarding.

DEVELOPING A KNOWLEDGE BASE

Figure 2-2 The VetScan chemistry analyzer can be used to perform whole blood or plasma chemistries on a variety of species, including both domestic and exotic species. Because of the light weight of the unit (11.2 pounds; 5.1 kg), it can easily be taken into the field to use with exotic or domestic species.

those times when no more than 100 μL of blood could be collected from a puppy or fractious cat. Again, the “proposed” expense would not only be useful for incorporating exotic pets but would simplify diagnostic testing for domestic species as well. “If you build it, they will come” is the famous line from the 1989 movie Field of Dreams. Can the same be said for an exotic practice? By expanding a practice, will clients inherently come? If veterinarians promote the fact that their practices accept exotic pets to their current client base, and, their clients have both domestic and exotic pets,1 veterinarians may realize an immediate expansion of their practices. A detailed history for domestic pets should include a question asking clients if they have other pets. This information can then be used to promote the practice’s treatment of exotics. In addition to promoting exotic pets with established clients, veterinary practitioners can also pursue new clients by establishing relationships with local retailers and local specialty clubs. Pet retailers are generally the primary source of exotic animals in a community. It is worthwhile to schedule time with a local retailer to discuss the specific needs of the different exotic pets they have for sale. It also can be beneficial for the veterinarian to offer a training seminar for the pet retail employees. Provid-

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As was mentioned previously, there are numerous opportunities to obtain the knowledge required to work with exotic pets. From the standard continuing education meetings to the more specialized scientific meetings, every need can be met. Each of the professional organizations in Table 2-1 has an annual scientific meeting, which allows veterinarians to share information in both formal and informal venues. These professional organizations also publish quarterly, peer-reviewed journals that provide cutting edge information related to the specialty. There are also learning opportunities through the Internet. Because there is a large amount of non–peer-reviewed and unregulated material on the Internet, veterinarians should be cautious and discerning. The two most accessible and useful Internet sources of information on exotic pets are the Veterinary Information Network (http://www.vin.com/) and the AVMA Network of Animal Health (http://www.avma.org/). Both of these networks provide veterinarians the opportunity to communicate with specialists in exotic pet medicine. There is a wealth of other literature available to veterinarians too. To assume that this single text could provide all that is needed to practice exotic pet medicine would be shortsighted. This book is intended to provide its readers with a collective introduction to the numerous exotic species that may present to their practices. Veterinarians that develop a special interest should obtain additional texts that go into more depth regarding the husbandry, medicine, and surgery for a class of animals. Box 2-1 represents a list of individual texts that can provide more detail on exotic animals (by class). Those interested in the ultimate pursuit of specialization in exotic pet medicine should consider advanced specialty training. Currently, there are two opportunities: board certification with the American Board of Veterinary Practitioners (AVBP)(Avian) and the American College of Zoological Medicine. Future certification opportunities in exotic mammal and reptile/amphibian medicine are being evaluated. There are two paths that can be followed to become board eligible: (1) Perform a residency at a certified institution, or (2) have at least 6 years of clinical practice in the field. For those interested in obtaining additional information regarding these programs,

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Chapter 2

Preparing Your Hospital for Exotic Pets

BOX 2-1

Textbooks That Can Be Used to Further Pursue the Husbandry, Medical, or Surgical Needs of Exotic Pets by Class

Amphibians Wright KM, Whitaker B, editors: Amphibian Medicine and Captive Husbandry, Malabar, Fla, 2001, Krieger, 499 pp. Avian Harcourt-Brown N, Chitty J, editors: BSAVA Manual of Psittacine Birds, ed 2, Gloucester, UK, 2005, British Small Animals Veterinary Association, 323 pp. Tully TN, Lawton MPC, Dorrestein GM, editors: Avian Medicine, Boston, 2000, Butterworth-Heinemann, 411 pp. Mammals Harcourt-Brown F, Textbook of Rabbit Medicine, Boston, 2002, Butterworth-Heinemann, 436 pp. Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2004, WB Saunders, 461 pp. Ornamental Fish Noga EJ, Fish Disease: Diagnosis and Treatment, Ames, 2000, Iowa State Press, 367 pp. Stosskopf MK, editor: Fish Medicine, Philadelphia, 1993, WB Saunders, 882 pp. Reptiles Girling SJ, Raiti P, editors: BSAVA Manual of Reptiles, ed 2, Gloucester, UK, 2004, British Small Animal Veterinary Association, 383 pp. Mader DR, Reptile Medicine and Surgery, ed 2, St Louis, 2006, WB Saunders, 1242 pp.

7 veterinary hospital has an established relationship. As mentioned previously, this type of relationship can be mutually beneficial. Regular staff meetings offer an opportunity to expand the knowledge base of the reception staff. Information from these meetings can be compiled for the staff and strategically placed near the phone for ready access. Veterinarians are finally recognizing the true potential of employing a certified veterinary technician. A competent veterinary technician can do many of the tasks veterinarians have historically done (e.g., processing hematologic, parasitic, and radiographic diagnostic test materials), providing the veterinarian with the opportunity to address other concerns in the hospital. In many of the technician programs, exotic pet medicine is becoming available to those who are interested in the field. For many veterinary technician graduates, like veterinary graduates, experience with exotic pets can be an important consideration for prospective employers. The additional training provides the “something extra” that enables that applicant to stand out among other competent individuals. Hiring technicians with experience in exotic pet medicine can help expedite the incorporation of an exotic pet practice into a hospital. There are now numerous continuing educational opportunities open for support staff. Many of the large continuing education meetings (e.g., Central Veterinary Conference, North American Veterinary Conference, Western Veterinary Conference) have tailored specific all-day programs for veterinary technicians, and exotic animal medicine is routinely a “track” offering. The major scientific meetings have also recognized the importance of providing training for veterinary technicians, and the meetings often have lectures and laboratories designed to meet the specific educational needs of these individuals.

PREPARING THE WAITING ROOM visit the following websites: http://www.abvp.com/ and http://www.aazv.org/.

PREPARING THE STAFF The reception staff at a veterinary hospital is arguably the most important “face” of the hospital. Regardless of the veterinarian’s reputation, an unimpressive first phone call or answer to a question can be sufficient to disuade a client from scheduling an appointment. Therefore, it is important to provide the reception staff the training necessary to guarantee their success in managing clients. The reception staff should be provided resources (e.g., veterinarian, veterinary technician, pet retailer, continuing education, and “cheat sheets”) that they can use to promptly and accurately answer questions. The veterinarinan is likely the best resource but may not always be available. In those cases, an experienced veterinary technician could assist in answering any questions. When the questions are husbandry related, directing the client to a pet retailer would be appropriate. The pet retailer should be an individual with whom the

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Preparing a waiting room for the inclusion of exotic pets can take some forethought. First, it is important to realize that not everyone finds 6 foot (1.8 m) long boa constrictors something they want to examine up close. To ensure the comfort of all clients, it may be wise to establish two waiting rooms. If this is not possible, then scheduling appointments so that the exotic and domestic pet appointments are at different times may be useful. By its very definition, the waiting room is not a place most people want to spend much time. Because people live busy lives and waiting is generally not a favorite pastime, it might be a good idea to change the name. It could be called the educational room or the information room, for example. Regardless of what the room is called, it should be set up to maximize clients’ time before their pet is examined. The reception staff can initiate the educational experience by providing species-specific brochures (http://www.exoticdvm.com/index. cfm?fuseaction=books.showIcons&) that provide information regarding the client’s pet. The provision of preliminary information can save the veterinarian and veterinary technician time later that otherwise would be used for client education.

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8

MANUAL OF EXOTIC PET PRACTICE

The preexamination period is a good time to acquire initial historical information. Providing the client an easy-to-read and thorough history form to complete before the examination can again save time later for the veterinary staff and keep the client involved in thinking about his or her pet. The reception area should reaffirm the hospital’s interest in exotic pet medicine. Information charts and appropriate exotic pet artwork can be used to generate a feeling of warmth for the client and provide additional educational information. Some veterinary hospitals will maintain live exotic pets in their waiting rooms. This can be used to reinforce the appropriate care of exotic pets. Easyto-read signs that discuss husbandry and care of the animal can be placed around the animal’s habitat. The placement of display animals reinforces that the veterinary hospital has an interest in caring for exotics and provides the veterinarian and his or her staff real-world experience in exotic pet management and husbandry. If live animals are kept in the hospital for display, it is important that the habitat be maintained to high standards.

NECESSARY EQUIPMENT Most veterinary practices have the equipment (e.g., isoflurane anesthesia) needed to provide a high standard of veterinary care for exotic pets. The additional equipment that veterinarians may use will depend on the species they accept and the extent to which they plan to practice on exotic pets. If the clincian’s desire is to provide basic health care and refer more complicated cases, then his or her equipment requirements will be minimal. If the clinician’s desire is to offer a comprehensive exotic animal care service, then it may be necessary to acquire additional specialized equipment. The following is a general list of equipment that veterinarians practicing exotic pet medicine should consider obtaining. (For information on specific equipment recommendations for particular species, see other chapters in this book.)

Housing Housing exotic pets in a veterinary practice is an important consideration. Many veterinary hospitals use stainless steel cages to hospitalize their dogs and cats. These cages can be used for some exotic mammal and avian patients, but they are generally less desirable for reptiles. I have used these cages for large reptiles, but had to use external heating sources to maintain an appropriate environmental temperature. Temperaturecontrolled incubators are a must for exotic animal practices (Figure 2-3). These incubators allow the veterinarian to hospitalize a range of exotic species, while meeting the specific environmental needs of these different animals. Environmental humidity can also be controlled in some incubators.

Diagnostics A high-quality light microscope is an important tool to have in a veterinary hospital. Unfortunately, with the advent of national diagnostics laboratories, veterinarians have become less reliant on analyzing their own patients’ samples. However,

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Figure 2-3 Temperature-controlled incubators can be used to provide exotic pets a thermally controlled environment.

there are certainly times when samples cannot be processed in a timely manner with a laboratory, and the clinician needs to be able to evaluate the sample. Hematologic samples provide important information regarding the health status of a patient. Veterinarians who regularly work with lower vertebrates (e.g., fish, amphibians, reptiles, and birds) and invertebrates realize that these animals have nucleated red blood cells, and to perform a complete blood count requires one of the estimation techniques (e.g., eosinophil unopette). A quality microscope capable of providing 400 to 1000 × magnification is required. The microscope can also be invaluable for interpreting fine needle aspirates, cytologic scrapings (e.g., skin scrape or gill biopsy in fish), and fecal examinations. Biochemistry can also provide a significant amount of information regarding the health status of a patient. One of the major drawbacks for some hospitals is that their laboratory requires a sizeable volume (0.5-1.0 ml) of blood to perform a chemistry panel. The general recommendation for collecting blood from exotic species is that no more than 0.5-1.0 ml/100 grams (body weight) be collected. Based on these minimum requirements, it would be difficult to collect sufficiently sized blood samples on small psittacines (e.g., budgerigar, cockatiel), passerines (e.g., finch, canary), small reptiles (e.g., leopard gecko, Jackson chameleon), or fish to perform biochemistries. The VetScan (see Figure 2-2) is one tool that I have found useful for processing these small samples. In addition, the samples can be processed in less than 14 minutes, which is an important consideration when managing a critical case.

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Chapter 2

Radiographic imaging is a valuable diagnostic tool for the veterinarian. However, only high-quality radiographs are of any value. When considering radiographic equipment for a small and exotic animal practice, it is important to purchase equipment that can manage a range of patient sizes (e.g., canary to bullmastiff). I prefer a machine capable of a wideranging kVp (e.g., 40-100). Rare-earth cassettes and singleemulsion films should be used for taking radiographs of exotic animals. These films will enable the veterinarian to detect small changes in anatomy. Digital radiography is another consideration and would benefit the veterianrian when working on either exotic or domestic pets. Digital radiography enables the veterinarian to produce films quickly for interpretation, which is important when they have anesthetized patients. Ultrasound is a diagnostic tool that has variable usage among species. For example, ultrasound has limited value for evaluating the coelomic cavity of avian species, because it is encased by the keel and ribs, whereas the same diagnostic tool can be used to readily evaluate the viscera of a reptile patient. For ultrasound to be useful, the veterinarian should pursue continuing education to fully understand the operation of the device. Endoscopy has become an essential diagnostic tool for the exotic animal veterinarian. Both flexible and rigid endoscopes are routinely used. Rigid endoscopes (2.7 mm) are used to perform coelioscopies and cloacascopies in birds and reptiles. I have also found them to be invaluable for assisted tracheal intubation in rodents and lagomorphs. Flexible endoscopes are frequently used to assess the proximal and distal gastrointestinal tract and respiratory system. Veterinarians attempting to justify the purchase of an endoscope should also consider the many applications these tools can also provide for their domestic pet patients.

Pathology Because exotic pets can mask their illness, many cases are presented to the veterinarian as “dead, but breathing.” In those situations where the client has multiple animals, it is important to recommend having any dead animals necropsied or to sacrifice diseased animals to obtain a definitive diagnosis. There are a number of different pathology services available to the veterinarian, but not all of these services have pathologists who are competent with exotic species. There are two pathology services in the United States that have excellent track records working with exotic species: Northwest Zoopath (Monroe, WA; http://www.zoopath.com/) and Zoo/Exotic Pathology Service (West Sacramento, CA; http://www.zooexotic.com/ html/who.html).

SURGERY Loupes Because many of the exotic patients we work with are small, magnifying loupes can be very helpful in characterizing small changes during an examination of blood vessels and nerves during a surgery. Basic head loupes are inexpensive and can be

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9

Preparing Your Hospital for Exotic Pets

Figure 2-4 Magnification loupes can greatly enhance a veterinarian’s visual field during surgery.

purchased for $30 to $50. For higher magnification, clinical magnification systems (Figure 2-4) should be used. These systems can be used with or without additional lighting. A variety of systems are available (http://www.surgitel.com/), and these systems can cost between $1,000 and $2,000. I find magnifying loupes extremely useful during surgical procedures.

Radiosurgery When performing surgery on small patients, hemostasis is essential. What may appear as a small volume of blood in a larger animal can represent a significant blood loss in a smaller patient. For example, a 3-ml blood loss in a german shepherd would be considered unremarkable, while in a 100-g cockatiel it would represent 30% of the animal’s blood volume. Radiosurgery can be used to minimize blood loss. I would recommend a dual frequency unit (http://www.ellman.com/) that enables the veterinarian to incise, incise and coagulate, and fulgurate. Again, this type of unit could also be used for domestic pet procedures (e.g., incision, mass removal).

THERAPEUTICS Because of the small size of many of our exotic pet patients, it can be difficult to deliver therpeutics without significant dilution. For this reason, it is important to establish a relationship with a compounding pharmacy. A major benefit to using compounded drugs is that multiple concentrations can be created for the same drug. For example, I routinely have several concentrations of ivermectin (Merial Limited, Iselin, NJ) (stock 10 mg/ml; dilution 1 : 1 mg/ml, dilution 2 : 0.1 mg/ml) or butorphanol (Torbugesic, Fort Dodge Animal Health, Ft. Dodge, IA) (stock: 10 mg/ml, dilution 1 : 2 mg/ml, dilution 2 : 0.5 mg/ml) made to simplify dosing. The other benefit of using compounded drugs is that they can be flavored according to the animal’s taste. For example, my pharmacy routinely flavors ferret compounds with meat-based flavoring agents and

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10 rabbit compounds with fruit-based flavoring agents. One potential drawback associated with compounded drugs is that the drug may not be evenly distributed throughout the carrier. To evaluate the concentration of a compounded drug, I suggest sending the medication to a private analytical laboratory for evaluation.

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MANUAL OF EXOTIC PET PRACTICE

REFERENCE 1. Wise JK, Heathcott BL, Gonzalez ML: Results of the AVMA survey on companion animal ownership in US pet-owning households. J Am Vet Med Assoc 221(11):1572-1573, 2002.

12/4/2007 11:57:38 AM

Trevor Zachariah Mark A. Mitchell

C H A P T E R

3

INVERTEBRATES Invertebrates represent the largest group of animals on Earth, with approximately 1 million species characterized to date. Additional species are being discovered all of the time, and it has been estimated that as many as 30 million invertebrates may reside on planet Earth. The diversity among invertebrates is enormous, with over 30 different phyla and myriads of subsequent taxa. Differences between the groups are so great that the only common feature among them is that, as their name implies, they do not possess a true vertebral column. Though it is easy to overlook, invertebrates are vital not only to the natural world but also to the lives of humans. Many invertebrates play important roles in the fields of agriculture, ecology, biology, medicine, and commercial and industrial trades, to name just a few. Invertebrates have also become increasingly popular as pets and display animals in zoological collections. There are numerous reasons for this trend, including the “awe factor,” relative inexpense of the animals, and their low-maintenance husbandry requirements. Because of their importance and increasing popularity, it is expected that invertebrate species will be presented to veterinarians with increased frequency. Unfortunately, like many other exotic pet species, little is known about the medical needs of these animals. To date, only a limited amount of research in this field has been published, and there is a strong need to expand the current knowledge base of invertebrate medicine. This chapter serves as an introduction to the biology and medicine of captive invertebrates.

COMMON SPECIES KEPT IN CAPTIVITY The only unifying feature common to all invertebrates is an absence of a vertebral column. Both because of this fact, and the diversity of species, the taxonomy of invertebrates is

dynamic. Because of the large numbers of invertebrate taxa, it is not possible to review all groups in this chapter. Instead, this chapter reviews those groups important to the captive pet trade or aquaculture.

Cnidarians There are approximately 10,000 different species of cnidarians, and the majority of these animals are found in the marine environment. Cnidarians are classified either as polyps or medusae. The polyps include the corals (Figure 3-1), hydrae, and anemones (Figure 3-2). The medusae include the true jellyfish and the box jellyfish. Cnidarians can be found as individuals or colonies, depending on the group. Most are carnivorous, subduing and killing their prey with specialized cells called cnidocytes (see Anatomy and Physiology). Jellyfish are one species with cnidocytes, which also are venomous to humans.

Gastropods There are approximately 60,000 different species of gastropods, and these animals are primarily aquatic, although terrestrial forms exist. Among the aquatic gastropods, the majority of animals are found in benthic habitats. Gastropods are the most numerous and diverse class within the phylum Mollusca and are the only group to have members that have evolved a terrestrial lifestyle (the Pulmonata). The gastropods are primarily represented by the snails (Figure 3-3) and slugs.

Arachnids There are approximately 70,000 different species of arachnids, and more than 80% of the animals in this group are spiders

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12

Figure 3-1 Maze brain coral (Platygyra sp.). These stony corals are popular in the aquarium hobby. (Photo by Trevor Zachariah.)

Figure 3-2 Bubble tip anemone (Entacmaea quadricolor). As indicated by this photo, these anemones are popular with clown fish. (Photo by Trevor Zachariah.)

MANUAL OF EXOTIC PET PRACTICE

Figure 3-3 Turbo snail (Turbo sp.). These marine gastropods are commonly recommended for marine aquaria. In this image, note the dextral twist of the shell, the extended antennae, the oral opening between them, and the large foot. (Photo by Trevor Zachariah.)

Figure 3-4 Chilean rose spider (Grammostola rosea). Because of its hardy and docile nature, this species of giant spider is commonly found in captivity. (Photo by Trevor Zachariah.)

and mites. The majority of arachnids (Figure 3-4) are terrestrial and carnivorous. The arachnids, like the cnidarians, may possess specialized tools (e.g., venom) to capture and kill prey. In some species (e.g., spiders and scorpions), the venom can also be harmful to humans. Certain species of arachnids, such as the mites and ticks, have evolved to live as parasites on vertebrate hosts.

all of these animals are nocturnal detritivores (Figure 3-5). Diplopods possess more legs than any other animal. Unlike the chilopods, diplopods are secretive and prefer to hide from predators rather than challenge them. Diplopods do not pose any real danger to humans.

Myriapods

Insects

There are approximately 13,000 different species of myriapods, all of which are terrestrial. Centipedes (≈3000 species) are from the order Chilopoda, and most of these animals are nocturnal predators. These invertebrates possess fangs, which they use to envenomate their prey or potential predators. Millipedes (≈10,000 species) are from the order Diplopoda, and

The insects represent the largest group of invertebrates, with over 900,000 described species. It is estimated that more than 75% of the animal species on Earth are insects. The greatest diversity among any group of living animals is seen with the insects. Most are terrestrial, but some have also developed the ability to fly. Some species of insects have developed parasitic

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Chapter 3

13

Invertebrates

Figure 3-5 A fire millipede (Aphistogoniulus sp.). Millipedes of a variety of colors and sizes are available to hobbyists. (Photo by Trevor Zachariah.)

Figure 3-6 A common sea star found in the marine trade. These echinoderms remain quite popular with hobbyists. (Photo by Trevor Zachariah.)

life cycles, many of which include human involvement. Some species of insects can pose a danger to animals and humans through their defensive mechanisms (e.g., bees). Numerous species are of economic importance to humans as pests.

Crustaceans There are approximately 42,000 different species of crustaceans, most of which are aquatic. The taxon is a diverse group, with many of the inconspicuous species playing a central role in ecologic webs. Some of the larger species are important to aquaculture.

Echinoderms There are approximately 6,000 different species of echinoderms. Many of these animals are common to the commercial aquarium trade, including the sea stars, brittle stars, sea urchins, sand dollars, sea cucumbers, and sea lilies (Figure 3-6). All of these species are marine and benthic. The echinoderms share a five-part radial body plan, also known as pentamerous symmetry, and have the ability to voluntarily move connective tissue, known as catch connective tissue (see Anatomy and Physiology).

ANATOMY AND PHYSIOLOGY Veterinarians working with any species must develop a basic understanding of the anatomy and physiology of that animal. This is no different with invertebrates. However, due to the great diversity of form and function among this large group, only a basic introduction to the anatomy and physiology of the invertebrates is presented here. For additional information, see the Suggested Readings at the end of the chapter.

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Figure 3-7 Sebae anemone (Heteractis sp.). In the center of the image is the slit mouth of the anemone, surrounded by a number of tentacles. (Photo by Trevor Zachariah.)

Cnidarians Whether in the form of a polyp or medusa, all cnidarians have a basic body plan that is radially symmetric. There are two tissue layers: the epidermis, which lines the outside of the animal, and the gastrodermis, which lines the inside of the animal. These layers are separated by a nonliving layer of elastic, gelatinous material known as the mesoglea, which provides structure and buoyancy without metabolic cost. Tentacles ring the mouth, the single opening to the digestive system (Figure 3-7). All surface tissues perform direct gas exchange. In the tentacles, and sometimes in the living tissue layers of the body, cnidocytes can be found. Cnidocytes contain a specialized, secreted organelle called a cnida. When stimulated by chemical or tactile cues, the cnidae use osmotic pressure to

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14 propel a hollow tube with great force out of the cell toward prey or predators. The tube can be used to physically subdue or inject venom into prey. The venom used in cnidae can be quite potent and may also be used for defense. Each cnida can be used only once, after which the cnidocyte must secrete a new one. The nervous system is comprised of a web of neurons, and attached to it are sensory systems that belie the simplicity of this web. Statocysts assist with balance, ocelli sense light, and sensory lappets register tactile stimuli. Box jellies take this one step further and have complex eyes that may be capable of forming images. The muscles of cnidarians are comprised of epitheliomuscular cells. Cnidarian muscles serve two purposes: (1) contractile functions to assist with locomotion, and (2) intracellular food digestion. The muscles are arranged in longitudinal and circular arrangements, allowing for a variety of types of movement. The most common locomotive movement for jellyfish is jet propulsion, and that is accomplished by contracting and releasing the body against the mesoglea, which acts as a spring. The mouth opens into the coelenteron, a cavity that is lined with the gastrodermis. The coelenteron may be partitioned into pouches or canals to increase the surface area of the gastrodermis and distribute nutrients throughout the animal. Many species also harbor mutualistic algae, known as zooxanthellae, which provide another source of nutrition. Cnidarians have an amazing ability to heal and regenerate. This is also reflected in the fact that asexual reproduction by budding or fission is common to cnidarians. Sexual reproduction also occurs, and most cnidarians are gonochoric. Fertilization is external and is followed by the development of a ciliated, planktonic, nonfeeding larva, known as the planula.

Gastropods Many gastropod species are characterized by the presence of a shell. The shell of most species is coiled in a dextral manner (e.g., right-handed), whereas a few species possess sinistral shells (e.g., left-handed). The central axis of the shell is the columella, and the opening at the base of the shell is the aperture. The first revolution of the shell is known as the body whorl, and this is where most of the visceral mass of the animal resides. The remaining whorls are collectively known as the spire, which culminates at the peak, or apex, of the shell. There are also many species with reduced or nonexistent shells. Gastropods are connected to their shells by a columellar muscle, which extends into the foot and to the operculum, if present. The foot is a mass of muscles and connective tissues, and it functions as the locomotory organ of the animal. Some species possess an operculum on the foot, which is a rigid disc that acts as a door to the aperture when the animal is retracted into its shell. The internal anatomy of gastropods is determined by their development, which includes a 180-degree torsion of the body. This leaves the majority of the mass of the body (and the shell, in those species which possess one) atop the head and foot.

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MANUAL OF EXOTIC PET PRACTICE

The process of torsion also causes a reduction or absence of some of the organs—ctenidia, osphradium, kidney, heart auricle—on the side corresponding to the direction of the torsion (e.g., left side for sinistral and right side for dextral). The space within the body whorl created by the torsion process is known as the mantle cavity. As the name implies, the mantle cavity is lined with the tissue known as the mantle, which secretes the shell. In the terrestrial species, the mantle cavity is modified into a sac-like structure with increased vasculature and an opening called the pneumostome. Thus equipped, the mantle cavity functions as a primitive lung. In the aquatic species, the mantle cavity contains the ctenidia, or gills, adjacent to which is the osphradium, an organ specialized for chemoreception. The heart lies within the pericardial sac and is located within the mantle cavity. The heart is a single ventricle and supplies hemolymph to an open circulatory system. Some species possess an elongated siphon for intake of water into the mantle cavity. Gastropods have a tubular digestive system that begins with a mouth and ends with an anus. The basic organs of the digestive tract include a buccal cavity, esophagus, stomach, intestine, and rectum. There are many variations on this theme, including a crop as part of the esophagus, a crystalline style, a gizzard, and a cecum. The crystalline style is found in the stomach of herbivorous species and provides mechanical and enzymatic assistance with digestion. The nudibranchs, or sea slugs, have a highly branched digestive tract. They prey upon cnidarians, and the digestive tract delivers undigested cnidocytes to the many tentacles that sprout from their bodies. Some gastropod species possess a proboscis within which the buccal cavity resides. Within the buccal cavity is a specialized feeding organ known as the radula. The radula is a layer of chitinous teeth that lie over a mass of cartilage and muscle known as the odontophore. The odontophore supports and moves the radula, which has evolved into many different forms. In some species, the radula serves as a harpoon, whereas in others, it serves as a rasp. The function of the radula is based on the feeding mode of the gastropod. The teeth of the radula vary in number and are continually worn and replaced. Gastropods sense the world through a variety of organs. Most gastropods have two eyes, which can be found on eyestalks or at the base of the cephalic tentacles. In a few species, the eyes may be capable of forming images; however, the majority of gastropods are able only to sense light through their eyes. Tentacles primarily serve chemoreceptive and mechanoreceptive purposes. The osphradium serves a chemoreceptive purpose. A pair of statocysts can be found in the foot and provide a sense of orientation. Also, a few species of gastropod appear to have magnetoreceptors. Almost all gastropods reproduce sexually. Primitive gastropods are gonochoric and perform external fertilization. More evolved species, including those that are terrestrial, are hermaphroditic and rely on internal fertilization. The planktonic larval stage of gastropods is called a veliger, and it is characterized by a large, ciliated organ known as the velum. The velum is used for locomotion, food collection, and gas exchange.

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Chapter 3

Invertebrates

Figure 3-8 Mexican redknee spider (Brachypelma smithi). Line A delineates the region of the prosoma, and line B delineates the region of the opisthosoma. The pedicel is the narrowing that connects the two regions. (Photo by Trevor Zachariah.)

15

Figure 3-9 Mexican redknee spider (Brachypelma smithi). Note the large chelicerae and the attached fangs. The opening of the mouth is found between the bases of the chelicerae. (Photo by Trevor Zachariah.)

Arachnids The body plan of the arachnids has two major components: the prosoma and the opisthosoma (Figure 3-8). The prosoma is comprised of a fused head and thorax that is covered dorsally by a carapace and ventrally by a sternal plate. The pleurae join the two and are flexible, which allows them to move relative to each other. The opisthosoma, or abdomen, contains the majority of the internal organs. In spiders, the prosoma and opisthosoma are joined by a narrow bridge called the pedicel. In scorpions, the two body segments are fused, and the opisthosoma is segmented and divided into two parts: the anterior mesosoma and the posterior metasoma, or tail. The metasoma is comprised of five to seven segments and, at its terminus, has a telson with a stinger. Most of the appendages originate on the prosoma. The most cranial pair of appendages are called the chelicerae. The chelicerae help to grasp and tear prey and bear the fangs in those species that have them (Figure 3-9). After the chelicerae, the next pair of appendages are called the pedipalps. The pedipalps come in a variety of forms and can serve different functions. In spiders, the pedipalps are similar to the legs but lack the metatarsal segment, whereas in scorpions, they terminate with pincers. In both groups of arachnids, the pedipalps are used to grasp prey and assist with copulation. The remaining prosomal appendages are the walking legs, of which there are four pairs. Each leg has seven segments, including (proximal to distal) the coxa, trochanter, femur, patella, tibia, metatarsus, and tarsus. The distal end of each leg bears a tarsal claw and, in many species, scopulae, dense tufts of hair that allow the arachnid to climb. The last set of spider appendages originate on the posterior end of the opisthosoma and include the three

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pairs of spinnerets. The spinnerets are variously modified to meet the needs of each species. Any of the appendages of arachnids may be autotomized, or voluntarily detached, and regenerated after several molts. The external openings to the arachnid respiratory organs are found on the abdomen. In spiders, the small openings, or spiracles, allow air to enter the multilayered, internal book lung, so called because hemolymph and air spaces interdigitate and look like the pages of a book in cross-section. Scorpions have four pairs of book lungs, one for each anterior opisthosomal segment. Primitive spiders, like the giant spiders, or tarantulas, have two pairs of book lungs. More evolved spiders have branching, tubular tracheae, which perform gas exchange directly with the tissues. The book lungs are the site of gas exchange. Hemolymph is circulated throughout the body of arachnids by an open circulatory system. The heart is located on the dorsal midline of the opisthosoma, surrounded by a pericardium, and connected to an open-ended arterial system. The movement of hemolymph throughout the body is achieved by a combination of pressure and suction from the heart and its sac. Hemolymph is light blue in color due to the oxygencarrying molecule hemocyanin, which contains two copper atoms rather than iron as in the hemoglobin of mammals. The opisthosoma also contains the majority of the digestive system. This includes the extensive diverticula of the midgut where most of digestion takes place. Also in the opisthosoma are the Malpighian tubules and stercoral pocket, which are involved in excretory processes. The gonads and silk glands are also present in the abdomen.

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16 The prosoma contains the anterior portion of the digestive tract, including the diverticula of the midgut, which extend through the pedicel; the sucking stomach; and the mouth, which is just posterior and ventral to the chelicerae. A pair of venom glands are also found in the prosoma of spiders. These glands are under voluntary control and are connected directly to the fangs. The prosoma is highly muscular and contains pseudoskeletal, cartilage-like structures called endosternites, which serve to anchor the muscles. The prosomal muscles maintain hemolymph pressure by contracting and relaxing the carapace and sternal plate, which in turn allows for extension of the appendages. Sensory perception in arachnids is achieved through a number of specialized organs. Spiders are covered in different types of hairs that allow for sensing of tactile, seismic, and chemical stimuli. Scorpions possess paired pectines, which are paired comb-like organs used to detect chemical and seismic stimuli. The pectines are located caudal to the last pair of legs.1 Both spiders and scorpions possess eyes, although the number (up to 12) varies. The visual acuity of arachnids can also vary among species. It is generally believed that giant spiders have poor vision, whereas jumping spiders (family Salticidae) can make well-developed images. In many of the New World giant spiders, hairs are used as a defense mechanism. Urticating hairs, located on the opisthosoma, are small, barbed structures that can be discharged to ward off a threat. Giant spiders use their caudal pair of legs to rapidly kick these hairs into the air. When the urticating hairs settle on the body surfaces of a potential predator, they cause severe irritation. Individual spiders may have more than 1 million urticating hairs on their abdomen, at a density of approximately 10,000 hairs per square millimeter.2 The urticating hairs are replaced after each successive molt. Arachnids reproduce by internal fertilization; however, because of their predatory nature, copulation can be somewhat dangerous. Female spiders produce egg sacs, whereas female scorpions gestate their eggs internally and are ovoviviparous. Maternal care is rare among spiders, whereas scorpions invest significant energy into caring for their young. Female scorpions carry their newly delivered young on their dorsum to reduce predation. The life span of some scorpion species may reach 25 years.3 The most common species kept in captivity, the African emperor scorpion (Pandinus imperator), has a life span of 3 to 8 years.1 Male giant spiders live only a short time after reaching sexual maturity and have a life span of 6 to 18 months, on average.4 Female giant spiders, however, live significantly longer, with anecdotal reports of individuals surpassing 30 years of age. With proper captive care, it would not be uncommon for female giant spiders to live in excess of 20 years.

Myriapods The myriapod body plan is elongated and composed of numerous segments (Figure 3-10). Each segment, except for the head and anal segments, bears either one (centipedes) or two (millipedes) pairs of legs, although the first few segments in milli-

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Figure 3-10 Giant African black millipede (Archispirostreptus gigas). Note the large size and long, tubular body form. (Photo by Trevor Zachariah.)

pedes bear only one pair of legs. Despite their names, centipedes and millipedes do not have 100 and 1000 legs, respectively. In actuality, these animals generally have 40-60 and 150-200 legs, respectively. Most of the body mass of the myriapod consists of the trunk. Millipedes are cylindrical in shape, with a hard, calcified exoskeleton. Centipedes are dorsoventrally flattened, with no waxy outer cuticle layer. Centipedes are built for speed when catching prey, whereas millipedes are slow and built for powerful digging. Besides the legs, myriapods have other appendages that are important for their survival. A pair of antennae are located on the head and the jaws, providing sensory input. Centipedes also possess a pair of forcipules on the first body segment, which are essentially venomous fangs used for acquiring prey. On the anal segment of centipedes is a pair of anal legs. These structures can have various functions, including tactile, defense, and aggression, depending on the species.3 The digestive system in all myriapods is long and tubular. Centipedes have a pharynx and esophagus that represent the majority of the gut length, whereas the millipede gut consists primarily of midgut. Millipedes have salivary glands associated with the oral cavity, whereas centipedes have a variety of glands associated with the pharynx and esophagus.3 The paired Malpighian tubules serve as the primary excretory organs. Millipedes have a layer of tissue that surrounds the midgut that has both energy storage (e.g., glycogen) and detoxification properties.3 The heart is a tubular organ that lies dorsally along the length of the trunk. Ostia act to move blood in and out of the heart. The dorsal pericardial sinus is formed by a horizontal membrane, as is the ventral perineural sinus. The perivisceral sinus is located in the middle of these other sinuses.3 Myriapods have an open circulatory system. In myriapods, gas exchange occurs via tubular trachea that delivers oxygen directly to the tissues. The trachae open to the environment through spiracles in the exoskeleton. In most

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species, the spiracles cannot be closed, which greatly hampers water conservation and results in the necessity of maintaining a humid, moist environment. The spiracles are located ventrally in millipedes and laterally in centipedes. Sensory structures in myriapods consist mainly of the eyespots and antennae. The eyespots consist of a varying number of ommatidia (individual sensory units), depending on the species. In most myriapod taxa, the ommatidia are not clustered in densities high enough to form a true compound eye, such as is found in insects.3 Myriapods are not believed to be capable of forming images. Instead, it has been suggested that these animals are limited to sensing light and movement.3 The antennae are able to sense both tactile and chemical stimuli. Myriapods are gonochoric and practice internal fertilization. In general, female centipedes are protective of their egg masses until the young hatch and disperse.3 Centipedes generally have a life span of 4 to 6 years; millipedes live for 1 to 10 years.3

Insects The diversity of insect morphological forms is astonishing. However, all the forms are derived from modifications of a basic plan. For purposes of brevity, those important to captive insects will be presented. The insect body is divided into three body sections: the head, thorax, and abdomen. The head bears a single pair of dorsal antennae, a variable number of ocelli, a single pair of compound eyes, and the ventral mouthparts. The mouthparts are modified to reflect the feeding strategy of the taxa. For example, sucking mouthparts are found on moths and butterflies (order Lepidoptera); piercing and sucking mouthparts on aphids, cicadas, and assassin bugs (order Hemiptera); cutting and sponging mouthparts on flies (order Diptera); and chewing and sucking mouthparts on bees and wasps (order Hymenoptera).3 The insect thorax bears three pairs of legs and one or two pairs of wings. The legs each have six segments, including the coxae, trochanter, femur, tibia, tarsus, and pretarsus. Each leg ends in a pair of tarsal claws. The wings of an insect can be modified, even into different structures. Two extreme examples of this include the beetles (order Coleoptera), in which the cranial pair are hardened elytra (e.g., wing covers), and the flies (order Diptera), in which the caudal pair are reduced to gyroscopic halteres to aid in flight. The abdomen of insects is segmented and relatively devoid of appendages. A terminal pair of cerci are usually present as are, in some species, external genitalia.3 The abdomen is often the largest of the three basic body segments and houses the majority of the viscera. The digestive system of insects is divided into three regions: the foregut, midgut, and hindgut. The foregut consists of the mouth, pharynx, esophagus, crop, and proventriculus. The midgut is the primary site for food digestion. Attached to the anterior end of the midgut are two to six ceca.3 The hindgut is comprised of the intestine, rectum, and anus. A variable

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number of Malpighian tubules (e.g., 2 to 250, depending on the taxon) are attached to the anterior end of the hindgut.3 The hindgut serves as the major excretory center in the insect body and resorbs most of the water from the digestive system. The circulatory system of insects is open, and the tubular, ostiate heart is found in the dorsal abdomen. The aorta extends cranially from the heart and is the only hemolymph vessel. An accessory heart is present at the base of most appendages, and facilitates the delivery of hemolymph to the tissues. Oxygen is delivered directly to the tissues by tubular trachae, in a fashion that is similar to that of myriapods and some arachnids. Spiracles are found on the thorax and abdomen, but not on the head.3 Unlike the spiracles of myriapods, those of most insects can be closed to prevent loss of moisture or water. In insects, the perception of multiple types of sensory stimuli is performed by sensilla, or hair-like receptors. These structures are found all over the body, although the majority are found on the appendages.3 Chemical, tactile, temperature, and humidity receptors are found on the antennae and tarsi. The abdominal cerci contain tactile and seismic receptors. The ocelli are used to detect changes in light intensity and aid in orientation.3 Visual stimuli are detected by the compound eye, and each of the ommatidia has its own lens. Thus, contrary to popular misconception, the insect brain, similar to the brain of vertebrates, integrates the information from each ommatidium to form a mosaic image.3 Many insects also have tympanic organs for the detection of sound. Insects are gonochoric and practice internal fertilization. There are three types of development among the insects. Hemimetabolous development involves juveniles called nymphs, which are dissimilar from adults, are aquatic, and grow and molt until reaching a final molt into the adult form. Paurometabolous development involves juveniles that are also called nymphs; however, these are similar to adults and grow and molt into the adult form. Holometabolous development (e.g., metamorphosis) involves juveniles called larvae, which grow and molt until forming a pupa, from which the adult form emerges. Holometabolous development is a successful life strategy, as approximately 80% of insects (e.g., approximately 740,000 species) utilize it.3

Crustaceans The crustaceans, like the insects, represent a diverse group of animals, with variable body forms. One of the most commonly recognized forms, the order Decapoda (e.g., crabs, lobsters, crayfish, and shrimps), will be described here (Figure 3-11). The crustacean body is comprised of two sections: the head and trunk. The head is small relative to the trunk and, along with the anterior of the trunk, is covered by a carapace. Five pairs of appendages adorn the head: two pairs of antennae, a pair of mandibles, and two pairs of maxillae. The maxillae assist with feeding. The head also bears one pair of compound eyes that are located on movable stalks.

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Figure 3-11 Louisiana red swamp crawfish (Procambarus clarkii ). This species is a typical decapod crustacean. P. clarkii are farmed extensively for human consumption. (Photo by Trevor Zachariah.)

The anterior section of the trunk bears eight pairs of appendages. The anterior three appendages, or maxillipeds, assist with feeding, while the posterior five, pereopods, are used for walking. There are seven segments of the pereopods, including the coxae, basis, ischium, merus, carpus, propodus, and dactyl. In many species (e.g., crabs, lobsters, and crayfish), the first pereopod is modified into an enlarged cheliped or pincer that is used for defense, food acquisition, and courtship. Limb autotomy is a common occurrence in decapods and most often occurs as a result of combat with conspecifics or in defense against predators. The limbs regenerate with later molts. The posterior of the trunk is comprised of a variable number of segments, depending on the taxon. The terminal end of the trunk bears a telson and paired uropods, and together, these form a tail fan. The tail fan helps to create the backward thrust of shrimp, crayfish, and lobsters. The five pairs of appendages arising from the posterior trunk are the pleopods. The pleopods are biramous and may be modified to serve a variety of functions, including swimming, burrowing, creating ventilating or feeding currents, brooding eggs, gas exchange, and copulating.3 In crabs, the posterior abdomen and pleopods are reduced and found ventral to the carapace. The digestive system begins with the cranial mouth and leads into a two-chambered stomach. In the stomach, both mechanical and chemical digestion of food occurs. Absorption of food occurs in the ceca, which are connected near the junction of the stomach and intestine. The intestine follows the length of the abdomen and terminates at an anus in the telson. Decapods have a compact, ostiate heart. This organ is located dorsally under the carapace. The heart is connected to a system of arteries (seven main arteries leave the heart), capillaries, and venous sinuses.3 Hemolymph is transferred through the circulatory system to the gills for oxygenation. The gills, of which they may have up to 24 pairs, are also responsible for

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Figure 3-12 Bubble tip brittle star (Ophiarachna sp.). This is a view of the oral side of the central disc of a brittle star. In the center of the image is the opening of the mouth. (Photo by Trevor Zachariah.)

excreting nitrogenous wastes.3 Ion balance in crustaceans is maintained primarily by the antennal glands (e.g., green glands), which are located in the cranial aspect of the head. Urine is created and stored in a bladder that opens near the base of the ventral pair of antennae. Sensory perception in crustaceans is accomplished by the eyes, antennae, and appendages. The stalked eyes are somewhat mobile, and some crustaceans may be able to detect color.3 Setae (hair-like receptors) are capable of detecting chemical stimuli and are primarily found on the antennae and appendages. Aesthetascs, or collections of setae, are located on the dorsal pair of antennae. Statocysts are found at the base of the dorsal antennae and assist with orientation. In some crustaceans, statocysts may also be found on other appendages.3 Crustaceans are primarily gonochoric, with a few (hermaphroditic) exceptions. Internal or external fertilization is possible, depending on the taxon. In most decapod species, females carry the eggs until they hatch. The egg masses are held on the ventrum by the pleopods. Decapods go through an indirect development, and there can be a number of larval stages. Other crustacean taxa may have direct or indirect development.

Echinoderms The body plan of echinoderms follows pentamerous symmetry. Though a type of radial symmetry, the echinoderms are not closely related to the cnidarians. Pentamerous symmetry is based on a central axis around which five body regions aggregate. The body surfaces are described as oral and aboral (Figure 3-12). Sea cucumbers (class Holothuroidea) maintain pentamerous symmetry in an elongated body form, with the ends of the animals being described as oral and aboral.

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Besides their recognizable body forms, another unique feature of echinoderms is the water-vascular system (WVS). The basic anatomy of this system starts with a madreporite, an eccentric, porous opening that is found on the aboral surface of sea stars and urchins and on the oral surface of brittle stars. The madreporite opens into a stone canal, which then leads to a circumoral ring canal. Leading perpendicularly from the ring canal are the radial canals and blind sacs known as polian vesicles. The radial canals reach into the arms of sea and brittle stars and along the inside to the aboral surface in sea urchins and cucumbers.5 Multiple lateral canals direct water from the radial canals to ampullae, each of which is attached to a tube foot. The ampullae are used in a manner that is similar to the bulb of a turkey baster, increasing pressure to extend the tube feet and decreasing pressure to contract them. The function of the polian vesicles has not been fully determined, but it may serve in aiding maintenance of fluid pressures within the WVS. The madreporite and stone canal function to maintain fluid pressure within the WVS. The stone canal is supported by calcareous ossicles and is lined with cilia, which beat to create water flow. The fluid within the WVS is essentially seawater, with increased cellular, protein, and potassium concentrations.3,5 The body wall of echinoderms is comprised of regions called ambulacral areas and interambulacral areas. Ambulacral areas represent those regions that bear the tube feet. These two types of areas generally alternate around the oral-aboral axis of sea urchins and cucumbers, whereas only ambulacral areas are found on the oral side of the arms of sea and brittle stars. The exoskeleton of echinoderms contains numerous ossicles, comprised of calcite microcrystals, embedded in the dermis. Ossicle structure can vary at the species level and is often a trait used to identify echinoderms. Some ossicles are modified for specific purposes, such as the paxillae of burrowing sea stars. In these animals, the ossicles form a protective shield for the aboral surface. Pedicellariae are specialized ossicles that occur on many different echinoderms. Pedicellariae are stalked or sessile ossicles that have a set of jaws and are used for defensive purposes. Sometimes these pedicellariae are equipped with poison glands (see Human Health Hazards). Another unique feature of echinoderm anatomy is catch connective tissue. This tissue is mutable, which allows these animals to vary the rigidity of their bodies at will. The stiffness or softness of the dermis is due to the extracellular matrix, in which nerves have been found to terminate.3 Research has found that calcium ion concentrations vary proportionally to the rigidity of the tissues.3

HUSBANDRY Because of the enormity of invertebrate species, it is not possible to cover all of the husbandry needs of these animals in a single chapter. Instead, we will give a short review of the husbandry needs of the most common species; for a more detailed review, see Lewbart.6 In considering the captive

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Figure 3-13 Cnidarians are dependent on constant circulating water flow. If these animals are placed into an inappropriate system they can be damaged. (Photo by Mark A. Mitchell)

care of an invertebrate, it is best to have some background knowledge of the particular species’ natural history. Even though there is relatively little known about the needs of the myriad species of invertebrates, it is best to try to mimic the natural environment and diet as much as possible. Many times, it is best to maintain a simplistic approach, as more advanced attempts at maintaining these animals may have negative results.

Cnidarians As with most aquatic species, water quality is the most important factor associated with the successful management of cnidarians in captivity. Stoskopf 7 recommends the following guidelines: ammonia levels less than 0.1 ppm, nitrite levels less than 1 ppm, nitrate levels less than 10 ppm, dissolved organic matter levels between 0.5 and 3.0 ppm, undetectable phosphate levels, calcium levels between 400 and 450 ppm for corals, pH between 8.2 and 8.4, alkalinity between 3.2 and 4.5 mEq/L, and a specific gravity (as a surrogate measure for salinity) around 1.025 to 1.027. Trace elements are another important consideration, and these animals depend on the water to provide these essential nutrients. Unfortunately, little is known regarding the specific needs of these animals, so attempts should be made to mimic natural levels of trace elements based on the natural body of water from which these animals are derived. Water motion is particularly important for cnidarian species (Figure 3-13). Cnidarians are generally classified as being either mobile or sessile. Water motion is essential for both groups because it facilitates nutrient gathering and oxygenation. For mobile species, water motion is important also for transporting the organisms.7 Because sessile species cannot move away from their wastes or accumulated organics, water motion serves to

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20 disperse potential toxicants. Cnidarians exposed to excessive water motion can be injured. The force of the water movement can push cnidarians into the walls of the aquarium or other fixed objects within the aquarium.7 Even sessile cnidarians can be injured by excessive water movement, as the fixed organisms are battered by the substrate. Water motion can be provided by way of a power head, airstone, or wave maker. Water motion is generally measured in gallons of water moved per hour. Determining the most appropriate flow rate depends on aquarium size and volume. The authors generally look at the movement of the cnidarians in the aquarium to determine what is best. If the cnidarians appear to be moving too quickly or are being battered against the aquarium or substrate, then the flow rate should be reduced. Due to their diversity, the cnidarians as a group are well adapted to a variety of temperature ranges; however, individual species may have a relatively narrow tolerance for temperature changes.7 As a general guideline, tropical anemones and corals should be kept between 20° and 31° C (68°-87.8° F), with an optimal temperature being around 24° C (75.2° F).7 For temperate species, a temperature range of 20° to 24° C (68°-75.2° F) is considered more appropriate.7 Thermostatically controlled aquarium heaters are the best method to provide an appropriate temperature range within an enclosure. In larger aquaria, multiple heaters may be required to establish an appropriate temperature range. Thermometers should be placed in different areas of the aquarium to monitor temperature. Many cnidarians derive a significant amount of their nutrients, sometimes up to 90%, from symbiotic algae (e.g., zoochlorellae or zooxanthellae) that are embedded in their tissues.3 The loss of these organisms, due to improper lighting or water quality conditions, can be devastating for a cnidarian. Full spectral lighting that mimics the sun is considered ideal. The light should provide ultraviolet and visible light. Stoskopf 7 recommends photosynthetically active radiation (e.g., 400700 nm) with more flux density between 400 and 550 nm than between 650 and 700 nm. The lighting should be maintained close to the water surface (30%).37

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Multiple types of parasites affect echinoderms, including apicomplexans, amoebans, ciliates, algals, mesozoans, helminths, gastropods, and annelids.37 Lesions resembling neoplastic growths have been reported in one species of brittle star (Ophiocomina nigra) and one species of sea cucumber (Holothuria leucospilata).37 See Jangoux39 for a complete review of echinoderm infectious diseases. Evisceration is a defense mechanism practiced by some species and is a natural response to changing environmental conditions or a sequela to a pathological condition. Sea cucumbers are capable of forcefully discharging adhesive, sometimes toxic, Cuvierian tubules from their anus to entangle predators or intruders.3 Many sea cucumber species are also able to discharge parts of their digestive tract in response to stress or as a natural seasonal process.3 The Cuvierian tubules and digestive tract can be regenerated. Evisceration also occurs in severe cases of ulceration in sun stars (Solaster sp.). Sun star ulcers can also be much milder and are an idiopathic condition of captive individuals.5 General trauma occurs in echinoderms and will often lead to secondary infections. Regeneration is possible, even from severe injuries, although permanent defects may be present after healing.

THERAPEUTICS Cnidarians Treating cnidarians with chemical compounds can be a risky venture, as the accumulated knowledge and scientific research in this area are lacking. Whenever a treatment is attempted, it should be conducted, whenever possible, in a separate tank from the animal’s main environment and the animal monitored carefully. Methods of drug removal or dilution (e.g., activated carbon filtration, conditioned replacement water) should be on hand in case of problems and the animal must be rinsed off before being returned to the main enclosure.7 Antibacterial treatments appear to be well tolerated by many cnidarian species.7 In many species 5% Lugol’s iodine solution (5-10 drops/L, 10-20 minute bath per day) can be used to provide broad-spectrum antisepsis and cauterize damaged tissues, but this should be evaluated on a species by species level, as some coral species do not tolerate Lugol’s iodine well.7 Tetracycline baths (10 mg/L) are safe for many corals; however, this antibiotic can be chelated by calcium in the water, lowering the effective dose.7 In cases where the hardness of the water is high, the veterinarian may consider a higher dose of tetracycline. Chloramphenicol baths (10-50 mg/L) have been used with success.7 The dosing frequency reported can be highly variable (24-72 hours). Chloramphenicol baths may be used in conjunction with a pretreatment or posttreatment Lugol’s iodine bath.7 Stoskopf 7 recommends inactivating chloramphenicol by adding 60 ml of chlorine bleach to every 20 L of treatment water several hours before discarding it. Cnidarians can also be medically managed with topical therapeutics. Lugol’s iodine (5%) can be applied topically with

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swabs to cauterize and disinfect wounds. Topical application should not be for longer than 20 to 30 seconds.7 Pastes created from antibiotics, such as neomycin and kanamycin, or waterresistant compounding bases used by dentists have been used.7 As with any topical therapy, only a small quantity of the therapeutic should be applied initially to determine if a reaction may occur. Antiparasitic treatments against flatworms, protozoa, and metazoa generally follow similar plans for fish. Placing saltwater species in freshwater (1-3 minute bath) can be done with relative ease. Monitor the invertebrates closely for any obvious changes (e.g., color change), and remove immediately if they appear to be reacting negatively to the freshwater. Levamisole, an antiparasitic used regularly in vertebrates, can also be used as a bath treatment for cnidarians. Stoskopf 7 recommends an extended (8 mg/L, 24-hour) bath for cnidarians. Different species may react differently to these treatments, and the animals should be monitored closely during any treatment procedure. When considering treatment plans for aquatic animals, it is important to think outside the box. Medicating a feed that is being offered to a patient is a common practice, although these foods are often commercially prepared. Instilling a medication into a live prey item, such as brine shrimp, may be more difficult. One of the most common methods for medicating difficult-to-treat live prey is to use osmosis. Placing brine shrimp, a salt-loving species, into freshwater will lead to the movement of water into these organisms. If a medication is placed in the freshwater, it will move into the brine shrimp, too. One potential drawback of this technique is that it is difficult to ensure the quantity of medication in these prey species. More research is needed to elucidate this information.

Gastropods Delivery of antibiotics to gastropods can be achieved through several methods. Due to the porosity of the epithelium and gills of aquatic species, bath treatments are possible. The techniques described earlier will also work for these animals. Intramuscular injections via the foot, injection via the hemocoel in some species (e.g., opisthobranchs), and adding medication to the feed are other alternatives that can be used.30

Arachnids Systematic analytical studies on the use of antibiotics in arachnids have not been attempted to the authors’ knowledge. There are anecdotal reports, but most have varying results and are based on cases without a definitive diagnosis. Cooper 21 suggests the use of tetracycline solutions (e.g., 50 mg tetracycline in 25 ml fluid) for irrigating external lesions. Other preparations, primarily those used in the aquarium trade, have been used orally.4 Enrofloxacin has been used in giant spiders at doses of 10 to 20 mg/kg PO q24h without apparent toxicity to the animal.4 Trimethoprim-sulfonamides may be used topically or systemically,4 but little is known about their effect or correct dosages. Both fenbendazole (10-200 mg/kg PO) and

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33 oxfendazole (10-200 mg/kg PO q24h to biweekly) have been used safely in giant spiders, but they did not eliminate oral nematode infections.4 When medicating arachnids, the drug may be delivered orally with a syringe or injected into a prey item before it is offered to the spider. Again, injecting the compound into the prey species may result in less than optimal dosing. Mites are a common finding in imported arachnids. Most of the miticides used in veterinary medicine are toxic to all invertebrates, including arachnids. Although tedious, the method of removing mites with mineral oil on a cotton-tipped applicator is safe. Treatment recommendations for fungal infections in arachnids are limited.4,13 Pizzi has recommended the topical application of povidone-iodine (0.75% water-based solution) for giant spiders. Topical chlorhexidine can also be used to manage fungi on arachnids. The chlorhexidine or iodine can be placed on a cotton-tipped applicator and applied directly on the affected area of the exoskeleton. Other topical and parenteral medications may also prove effective, but further research needs to be performed to determine the efficacy of these treatments. Invertebrates that are dehydrated or suffering from hypovolemia (e.g., trauma with loss of hemolymph) should be given fluids to replace any losses. Many arachnids will consume water offered directly from a syringe.35 Animals that are active can be placed into a dish of shallow water and allowed to imbibe on their own will. It is important that the water not be too deep, as arachnids can drown if the water enters their respiratory openings.10,13,35 Giant spiders can be given parenteral fluids via the distal leg joint,13 heart and pericardial sac,22 and the opisthosoma.10 When considering rehydrating a spider, veterinarians should base the selection of a fluid on the type of dehydration. The majority of cases presented to veterinarians will be associated with a hypertonic dehydration. Only one study has attempted to determine the osmolarity and electrolyte balance of giant spider hemolymph.40 Based on that study, a “tarantula Ringer’s solution” was recommended. Interestingly, the osmolarity of the solution was rather hypotonic (201 mOsm/L). Physiologic (0.9%) saline (273 mOsm/L) has been recommended as an alternative fluid for rehydrating giant spiders,13 and we have been successful using this method. Pizzi4 also reports the successful use of lactated Ringer’s and Hartmann’s solutions for fluid therapy. An alternative to replenishing hypovolemic spiders with commercial fluids is to use a transfusion of hemolymph from a donor spider using a small-gauge catheter, as described by Visigalli16; however, the effects this procedure would have on the transfused spider are not known. The amount of injectable fluid that can be safely administered to a giant spider has not been clearly defined, but Stewart and Martin24 were able to safely administer 0.5 ml (4% body weight) of fluid into a 12-g spider. Johnson-Delaney22 has suggested that the fluid rate used for reptiles would be appropriate also for giant spiders. In a recent study by the authors, Grammostola rosea were given a single injection of physiologic saline into the opisthosoma representing up to 6% of their

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body weights, without any observed negative side effect. Additional research is required to determine the maximum volume and rate that fluids can be replenished in giant spiders and other arachnid species.

Myriapods The information available for therapeutic use in myriapods is severely lacking. There is anecdotal evidence that fluid therapy and oral antimicrobials have been attempted. For millipedes, fluids may be replenished by placing the animal in a shallow dish of lukewarm water. This will only work for active, alert animals. For centipedes, placing them in an enclosure with a high humidity has been recommended.14 This can be achieved by using dampened paper towels or moistened bedding. Injecting oral antibiotics into prey items is an appropriate method for dosing both millipedes and centipedes.14 Parenteral administration may also be attempted. Injection of watersoluble drugs into the hemolymph through the arthrodial membranes can be done, but dosing rates are empirical. The effect of parenteral antibiotics on the microbial flora of millipedes is unknown, but should be considered prior to administration.14

Insects As with the myriapods, there is little information available regarding the use of therapeutics in insects. The most studied species is the honeybee (Apis mellifera), and Cooper36 presents a limited formulary for insects in general (Table 3-1).

Crustaceans Because most crustacean species are aquatic and most captive species are cultured for human consumption, the compounds used for these species in the United States are regulated by the Food and Drug Administration.37 Animals that are not des-

TABLE 3-1

Formulary for Insects

Type

Name

Route

Fluids

Water Hypotonic saline* (0.2%-0.5%) Chlortetracycline Oxytetracycline Sulfadiazine Mineral oil Povidone-iodine Ketoconazole or nystatin Fumagillin Benomyl Povidone-iodine

PO, nebulization PO, nebulization, ICe PO, topical PO, topical PO PO Topical Topical

Antibiotics

Laxatives Antimycotics

Antiprotozoals Antiseptics

PO PO Topical

Adapted from Cooper JE: Insects. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell, p 218. *Do not use in stick and leaf insects. ICe, intracoelomic; PO, per os.

Ch003-X0119.indd 34

tined for human consumption, such as pets or institutional specimens, may be treated with medications in an extra-label manner.37 Table 3-2 provides a list of therapeutic agents and dosages used to manage penaeid shrimp. This formulary may be extrapolated for use in other crustaceans as well.

Echinoderms Treatment of echinoderms, like other invertebrates, is in its infancy. Harms5 has suggested that these animals can be treated with antibiotics via a bath or intracoelomic injection. He also suggested that chitin synthesis inhibiting parasiticides (e.g., lufenuron) might be applicable to these animals, although the dosing protocols, safety, and effectiveness of these compounds have not been described for these species.

SURGERY Cnidarians Surgical procedures of cnidarians are generally limited to debridement of lesions and fragmentation for propagation.7 When treating cnidarians, veterinarians should use sharp dissection to remove damaged tissue(s). Debridement should take place ideally in an enclosure separate from the animal’s primary enclosure to avoid contaminating the water system with debrided tissue, as this tissue can be infectious.7 There is a great deal of debate as to how to manage a defect that is created in a coral after debridement. Some veterinarians close the defect, whereas others manage the defect by second intention. Anecdotally, various substances have been used to fill defects with some success, including plasticine clay (with or without antibiotic impregnation), plaster of Paris, hydraulic cement, and specialized plastering compounds.7 In addition to these products, Stoskopf 7 has also recommended several other substances that deserve investigation, such as dental compounding bases, methacrylates, cyanoacrylate gels, and underwater epoxies. For an overview of coral fragmentation, see Stoskopf.7

Gastropods Gastropod shell injuries occur occasionally in captive collections. The gastropod shell can be repaired using the same basic concepts used for repairing chelonian shells. First, the shell should be disinfected. Dilute povidone-iodine or chlorhexidine and saline should be used for this purpose. It is important not to introduce these chemicals into the coelomic cavity, as they can be caustic to unprotected tissues. Dental acrylic, surgical epoxy/glue, and tape can be used to stabilize fractures. In severe, displaced fractures, cerclage wire can be used to reduce the fractures.

Arachnids To the authors’ knowledge, only one known elective surgical procedure has been described for arachnids. Passive integrated transponders were implanted into the opisthosomas of 12 giant

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TABLE 3-2

Formulary for Penaeid Shrimps

Type

Name

Dosage

Antimicrobials Antibiotics

Benzalkonium chloride Nitrofurazone Nifurpirinol Furazolidone Oxytetracycline Ormetoprim-sulfamethoxazole Malachite green Methylene blue Trifluralin Copper sulfate Zeolite Formalin Potassium permanganate

0.6-1.0 ppm, 24 h 1-2 ppm 1-2 ppm 10-20 ppm, 24 h 40-60 ppm, 24 h 50-100 mg/kg PO, 14 days 0.5-0.8 ppm, 24 h 8-10 ppm, 24 h 0.01-0.1 ppm 0.5-1.0 ppm, 10-12 h 100-120 kg/m2, applied to environment 0.5-1.0 ppm, 24 h 25-30 ppm, 30-60 min

Antimycotics

Antiprotozoals Antiparasitics

Modified from Noga EJ, Hancock AL, Bullis RA: Crustaceans. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell, p 192. PO, per os.

spiders, without complication, by Reichling and Tabaka.41 Though this procedure was developed for research purposes, it may prove to have clinical application in the future. Most surgical procedures performed on arachnids arise as a result of traumatic injury. Arachnids can naturally lose a limb through autotomy. When an arachnid suffers an injury to an appendage as a result of trauma or dysecdysis, the limb can be removed by the veterinarian with little damage, as the process is a natural response for the host.13,35 It is important to remove injured limbs, as they can lead to excessive loss of hemolymph. To remove an injured leg or pedipalp, the appendage should be grasped with hemostats at the level of the femur and separated with a sharp upward tug on the leg. Several authors suggest that because autotomy is a voluntary act, arachnids should not be anesthetized during the procedure.13,20 However, the process does result in tissue damage, which initially should mimic a noxious stimulus or aversion behavior. Some veterinarians take a similar approach with vertebrates (e.g., iguana tail autotomy, salamander limb autotomy). Trauma to the exoskeleton of an arachnid requires immediate medical attention, as a significant loss of hemolymph is likely to be fatal. If not extensive, traumatic or iatrogenic (e.g., hemolymph collection, surgery, induced autotomy) exoskeleton defects can be repaired. There are various methods for doing this, including the application of cyanomethacrylate (e.g., surgical tissue adhesive), cyanoacrylate (e.g., super glue), or nail hardener.4,13,20,35 Multiple applications may be needed depending on the extent of the lesion. Larger exoskeleton defects may be closed with 5-0 or 6-0 synthetic suture,22 although Pizzi4 discourages this practice as ineffective.

Myriapods The only described surgical procedures for myriapods are wound repair and appendage removal. For both millipedes and centipedes, the application of adhesives may be done in a

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manner similar to that described for arachnids. Large defects in centipedes may also benefit from closure with 5-0 to 8-0 synthetic suture.14 Damaged appendages may be removed by sharp dissection and may be sealed with adhesive.

Insects Although there are a number of surgical procedures described for insects, most of these are experimental in nature and not conducive to clinical medicine.36 However, Cooper36 does suggest that attempts at limb amputation, lesion removal and debridement, and wound repair (including suturing) be attempted in cases when deemed appropriate by the veterinarian.

Crustaceans As with many of the taxa discussed in this chapter, little information exists regarding surgical procedures in crustaceans. However, this should not preclude attempts at minor surgical procedures such as amputation and wound repair. Presumably, these would be performed in a manner similar to what has been described for other arthropod species, with concern taken for the unique aspects of crustacean anatomy and physiology. For example, the autotomy reflex of decapod crustaceans, with a fracture plane between the basis and ischium segments, may serve as a method of amputating limbs.

Echinoderms No surgical procedures have been described for echinoderms. Ophiuroids have an amazing ability to autotomize appendages, and this, along with their impressive regenerative capabilities, could be considered in cases where the animal suffers an injury.5

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MANUAL OF EXOTIC PET PRACTICE

HUMAN HEALTH HAZARDS Cnidarians The cnidarians include some of the most venomous species of animals in the world. Certain species of box jellies, in particular, maintain highly potent nematocysts that are capable of inflicting extreme pain and even death in humans. The most notorious cnidarians are Chironex fleckeri (e.g., Australia’s “sea wasp”), which is responsible for an average of two deaths each year, and Chiropsalmus quadrumanus (southeastern United States), which is responsible for at least one human fatality annually.3 Other species of cnidarians are also capable of inflicting intense pain on humans; however, most are harmless. Although not intended to be injurious, the sharp edges of some corals are known to cause lacerations in humans.

Gastropods Most gastropods are harmless. The family Conidae (e.g., cone snails), a group of carnivorous, marine snails, however, is an exception. Cone snails have a radula that is modified into a hollow harpoon that they use to envenomate prey. This weapon can also be used against humans. The venom of some species is toxic to humans and has been responsible for at least a few deaths.3 Fortunately, these snails are not common in the commercial trade.

Arachnids Arachnids are feared the world over, and this is unfortunate considering these animals prey on many of the insects considered to be pests or vectors for disease. One of the reasons that humans fear these animals is because of the perceived concern associated with their venom. With very rare exceptions, spiders possess venom. However, most are either unable to bite or their venom is not dangerous to humans. The World Health Organization lists four genera as exceptions: Atrax, Latrodectus, Loxosceles, and Phoneutria. The most common, from the standpoint of their relative abundance and popularity in the United States, are widow spiders (Latrodectus spp.) and recluse spiders (Loxosceles spp.). Widow spiders produce a neurotoxic venom that can cause abdominal and leg pain, high cerebrospinal fluid pressure, nausea, muscle spasms, and respiratory paralysis.3 Death is infrequent, and signs usually resolve within 3 to 7 days after symptomatic treatment with or without antivenin therapy.42 Recluse spiders produce cytotoxic venom that induces a necrotic ulcer at the site of the bite. It is rare that these envenomations become systemic.42 Symptomatic treatment is usually sufficient, and reports of mortalities are uncommon.42 Species from both of these genera are relatively common in captivity, although this practice is not recommended. Less commonly encountered, though particularly more potent in venom and aggressiveness, are the funnel web spiders (Atrax spp.) and the South American “banana spiders” (Phoneutria spp.). Giant spider bites are not considered serious. In general, Old World species are believed to have more potent venom

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than New World species.43,44 Most individuals envenomated by giant spiders develop mild to severe local pain, itching and tenderness, edema, erythema, joint stiffness and swollen limbs, burning feelings, muscle cramps, and temporary paralysis.43 Schultz and Schultz10 reported on the symptoms associated with the bites of various species of giant spider and found that all of them were mild and short-lived. Rather than toxicity, the real significance of giant spider bites may be the potential for mechanical injury. Fangs can be large, with those of adult Theraphosa blondi reaching up to 3 cm in length.31 As with any insult that compromises the skin, injuries from arachnid bites or stings have the potential to lead to secondary infections. Opportunistic pathogens may originate from the human or from the invertebrate. The microflora of the mouthparts of various captive giant spider species have been described, and a range of Gram-positive and Gram-negative bacteria found: Staphylococcus aureus, Staphylococcus spp., Bacillus megaterium, B. subtilis, B. cereus, Pseudomonas diminuta, P. aeruginosa, P. fluorescens, P. cepacia, Pseudomonas spp., Aeromonas hydrophilia, Acinetobacter calcoaceticus, Micrococcus varians, M. mucilaginosus, Coryneforms, and other Enterobacteriaceae.29,45 At least three of these bacteria (S. aureus, P. aeruginosa, A. hydrophilia) are known human pathogens.45 Infections with oral nematodes from the family Panagrolaimidae have been reported in giant spiders, and some species of this family are known to cause zoonotic disease.31 Giant spiders carry another potential risk to humans, their urticating hairs. As mentioned previously, New World species have urticating hairs on their opisthosomas. When alarmed, the spiders can brush off the hairs with their hind legs, which allows them to become airborne. Exposure to these hairs can occur from direct contact with the spider or indirectly from the animal’s environment, especially when cleaning and replacing substrate. Each urticating hair is covered by hundreds of small hooks and can cause severe itching when it makes contact with the skin.33 Cooke et al.2 have shown that these hairs can embed themselves to a depth of 2 mm in human skin. A severe skin reaction has been observed by the authors in a colleague working with the Theraphosa blondi colony at the Louisiana State University School of Veterinary Medicine. This individual was changing the substrate in the cages and had indirect contact with the spiders. A severe, intense, pruritic cycle ensued that affected the hands and forearms (Figure 3-24). The reaction lasted for 14 days. A single case of papular dermatitis with edema resulting from urticating hair exposure has also been described.46 Although dermatologic exposure to urticating hairs is a self-limiting condition, urticating hairs can be much more harmful to the human eye. The hairs can penetrate the cornea and cause a multitude of symptoms, including panuveitis, iritis, anterior synechiae, vitritis, chorioretinitis, and keratitis. A review of the literature revealed more than a dozen case reports of ophthalmia nodosa (a granulomatous disease) caused by contact with a giant spider. Ocular disease caused by urticating hairs can be serious and chronic in nature. It is recommended that latex or nitrile exam gloves be worn whenever handling or cleaning the enclosure of a giant spider. Eye pro-

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Invertebrates

deliver. Various insects (e.g., bees, wasps, ants) are able to affect humans through chemical means (e.g., venom, acidic secretions) as well. However, barring allergic reactions or mass attacks, the effects of insects on humans are rarely more serious than the discomfort that they cause. Wild-caught insects can serve as vectors for infectious diseases. Testing subsets of captive populations for those diseases of concern can be done to limit the likelihood for zoonotic disease transmission.

Crustaceans

Figure 3-24 Human dermatitis resulting from giant spider urticating hairs. This colleague came into contact with airborne urticating hairs from a goliath birdeater spider (Theraphosa blondi ). (Photo by Clare Guichard.)

The crustaceans in general do not pose much of a threat to humans. The most common injuries involve the pincing ability of the orders Decapoda and Stomatopoda (e.g., “mantis shrimps”). Such injuries may be severe when dealing with crustaceans of large size (e.g., lobsters). Grasping these animals at the cephalothorax can generally reduce the likelihood of being pinced.

Echinoderms tection may also be indicated. During the activity, it is best to avoid touching one’s face or any area of skin. Washing one’s hands afterward is also strongly recommended. Most scorpions produce a venom that is painful but not dangerous to humans. The exceptions are found in the family Buthidae, which includes Androctonus and Centruroides.3 Death from the neurotoxic venom is generally preceded by convulsions, paralysis of the respiratory muscles, and cardiac failure. Fortunately, some effective antivenins are available.3 Veterinarians working with these species should be preemptive and have antivenins on hand. Clients should be made aware of the potential hazards to owning these animals.

Myriapods Millipedes are generally slow, passive animals. However, many species are able to exude or squirt noxious substances from the glands found on the body segments.14 These substances can affect the human skin by staining, irritating, or blistering and can cause potentially serious injuries if the human eye comes into contact with them.14 As with arachnid species, latex exam gloves and/or eye protection is recommended when handling these animals. Large centipedes have fangs of commensurate size, and their venom can be quite painful. Deaths resulting from the bite of a centipede are difficult to authenticate, although the potential for this may exist for Scolopendra.3 Most species have the ability to move rapidly, so handling should be avoided if at all possible.

Insects Insects pose a variety of health hazards to the humans that interact with them. The most obvious is through the physical injury (e.g., bite, sting, or pinch) that many species are able to

Ch003-X0119.indd 37

Sea urchins can inflict severe injuries to humans. The spines of these animals can cause both direct cell injury and, in some species, a toxin insult.3 Delayed hazards include secondary opportunistic bacterial infections and chronic granuloma formation.5 Some species also possess venomous pedicellariae, which are meant to protect the animal from predators and can produce a painful reaction in humans.3

SUGGESTED READINGS Breene RG: The ATS Arthropod Medical Manual: Diagnosis and Treatment, The American Tarantula Society. Breene RG: Concise Care Guide for the 80 Plus Most Common Tarantulas, The American Tarantula Society. Brownell P, Polis GA: Scorpion Biology and Research, Oxford University Press. Daly HV, Doyen JT, Purcell AH: Introduction to Insect Biology and Diversity, Oxford University Press. Foelix RF: Biology of Spiders (ed 2), Oxford University Press. Frye FL: Captive Invertebrates: A Guide to Their Biology and Husbandry, Krieger. Hopkin SB, Read HJ: The Biology of Millipedes, Oxford University Press. Keegan HL: Scorpions of Medical Importance, University Press of Mississippi. Lewbart GA: Invertebrate Medicine, Blackwell. Polis GA: The Biology of Scorpions, Stanford University Press. Ruppert EE, Fox RS, Barnes RD: Invertebrate Zoology: A Functional Evolutionary Approach (ed 7), Brooks/Cole. Schultz SA, Schultz MJ: The Tarantula Keeper’s Guide, Barron’s.

REFERENCES 1. Frye FL: Scorpions. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 2. Cooke JAL, Miller FH, Grover RW, Duffy JL: Urticaria caused by tarantula hairs, Am J Trop Med Hyg 22(1):130-133, 1973. 3. Ruppert EE, Fox RS, Barnes RD: Invertebrate Zoology: A Functional Evolutionary Approach, ed 7, Belmont, Calif, 2004, Brooks/Cole-Thomson Learning. 4. Pizzi R: Spiders. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell.

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38 5. Harms CA: Echinoderms. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 6. Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 7. Stoskopf MK: Coelenterates. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2007, Blackwell. 8. Frye FL: Captive Invertebrates: A Guide to Their Biology and Husbandry, Malabar, Fla, 1992, Krieger. 9. Marshall SD: Tarantulas and Other Arachnids, Hauppauge, NY, 2001, Barron’s Educational Series. 10. Schultz SA, Schultz MJ: The Tarantula Keeper’s Guide, Hauppauge, NY, 1998, Barron’s Educational Series. 11. Breene RG: Concise Care Guide for the 80 Plus Most Common Tarantulas, Carlsbad, N Mex, 1998, American Tarantula Society. 12. Cappelletti A, Visigalli G: What every veterinarian needs to know about giant spiders, Exot DVM 5(pt 6):36-42, 2004. 13. Pizzi R, Cooper JE, George S: Spider health, husbandry, and welfare in zoological collections. In Proc Brit Vet Zoo Soc, pp 54-59, 2002. 14. Chitty JR: Myriapods (centipedes and millipedes). In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 15. Williams DL: Invertebrates. In Meredith A, Redrobe S, editors: BSAVA Manual of Exotic Pets, ed 4, Gloucester, UK, 2002, British Small Animal Veterinary Association. 16. Visigalli G: Guide to hemolymph transfusion in giant spiders, Exot DVM 5(pt 6):42-43, 2004. 17. Cooper JE: Invertebrate anesthesia, Vet Clin North Am Exot Anim Prac 4(1):57-67, 2001. 18. Applebee KA, Cooper JE: An anaesthetic or euthanasia chamber for small animals, Anim Technol 40(1):39-43, 1989. 19. Berzins IK, Smolowitz R: Diagnostic techniques and sample handling. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 20. Breene RG: The ATS Arthropod Medical Manual: Diagnosis and Treatment, Carlsbad, N Mex, 2001, American Tarantula Society. 21. Cooper JE: A veterinary approach to spiders, J Small Anim Pract 28(3):229-239, 1987. 22. Johnson-Delaney C: Exotic Companion Medicine Handbook, Lake Worth, Fla, 2000, Zoological Education Network. 23. Davies RR, Chitty JR, Saunders RA: Cardiovascular monitoring of an Achatina snail with a doppler ultrasound probe, Proc Brit Vet Zool Soc, Autumn Meeting:101, 2001. 24. Stewart DM, Martin AW: Blood pressure in the tarantula, Dugesiella hentzi, J Comp Physiol 88(2):141-172, 1974. 25. Williams DL: Sample taking in invertebrate veterinary medicine, Vet Clin North Am Exot Anim Prac 2(3):777-801, 1999. 26. Williams D: Studies in arachnid disease. In Cooper JE, Pearce-Kelly P, Williams DL, editors: Arachnida: Proceedings of a Symposium on Spiders and Their Allies, pp 116-125, 1992.

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27. Murray MJ: Euthanasia. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 28. Fiorito G: Is there pain in invertebrates? Behav Processes 12(4):383-388, 1986. 29. Stefano GB, Salzet B, Fricchione GL: Enkelytin and opioid peptide association in invertebrates and vertebrates: immune activation and pain, Immunol Today 19(6):265-268, 1998. 30. Smolowitz R: Gastropods. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 31. Pizzi R, Carta RL, George S: Oral nematode infection of tarantulas, Vet Record 152(22):695, 2003. 32. Baker AS: Acari (mites and ticks) associated with other arachnids. In Cooper JE, Pearce-Kelly P, Williams DL, editors: Arachnida: Proceedings of a Symposium on Spiders and their Allies, pp 126-131, 1992. 33. Foelix RF: Biology of Spiders, ed 2, New York, 1996, Oxford University Press. 34. Williams D: Integumental disease in invertebrates, Vet Clin North Am Exot Anim Prac 4(2):309-320, 2001. 35. Cooper JE: Emergency care of invertebrates, Vet Clin North Am Exot Anim Prac 1(1):251-264, 1998. 36. Cooper JE: Insects. In Lewbart GA, editor, Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 37. Noga EJ, Hancock AL, Bullis RA: Crustaceans. In Lewbart GA, editor: Invertebrate Medicine, Ames, Iowa, 2006, Blackwell. 38. Edgerton BF, Evans LH, Stephens FJ et al: Synopsis of freshwater crayfish diseases and commensal organisms, Aquaculture (1-2):57-135, 2002. 39. Jangoux M: Diseases of echinodermata. In Kinne O, editor: Diseases of Marine Animals, vol 3, Hamburg, Germany, 1990, Biologische Anstalt Helgoland. 40. Schartau W, Leidescher T: Composition of the hemolymph of the tarantula Eurypelma californicum, J Comp Physiol 152(1):73-77, 1983. 41. Reichling SB, Tabaka C: A technique for individually identifying tarantulas using passive integrated transponders, J Arachnol 29(1):117-118, 2001. 42. Diaz JH: The global epidemiology, syndromic classification, management, and prevention of spider bites, Am J Trop Med Hyg 71(2):239-250, 2004. 43. Escoubas P, Rash L: Tarantulas: eight-legged pharmacists and combinatorial chemists, Toxicon 43(5):555-574, 2004. 44. de Haro L, Jouglard J: The dangers of pet tarantulas: experience of the Marseilles Poison Centre, J Toxicol-Clin Toxicol 36(1-2):51-53, 1998. 45. McCoy RH, Clapper DR: The oral flora of the South Texas tarantula, Dugesiella anax (Araneae: Theraphosidae), J Med Entomol 16(5):450-451, 1979. 46. Ratcliffe BC: A case of tarantula-induced papular dermatitis, J Med Entomol 13(6):745-747, 1977.

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Stephen M. Miller Mark A. Mitchell

C H A P T E R

4

ORNAMENTAL FISH

Ornamental fish present an unusual paradox in that they are both well known and unknown to veterinarians. These animals are well known because they can be seen every day in the home aquarium, ornamental pond, pet store, and public aquarium,1 while at the same time they are unknown because knowledge regarding their health care is limited (e.g., in the areas of antibiotic residuals, antibiotic resistance, emerging diseases, antiquated or undocumented diagnostic and surgical techniques, and pain management issues). The purpose of this chapter is to address some of the current and former issues related to the health and well-being of ornamental fish.

FRESHWATER It would be impossible and impractical to categorize every species of freshwater fish in a single book chapter, so we will present information about the two major groups of fish that are commonly kept in the captivity (freshwater temperate and tropical species) and refer to them as a general classification of bony fishes known as Actinopterygii. In addition, we will also provide more detailed information regarding three of the important groups of captive freshwater fishes: catfish, cichlids, and cyprinids.

Freshwater Temperate Fish

COMMON SPECIES KEPT IN CAPTIVITY Fish represent the largest class of vertebrates, with more than 20,000 different species. This group also represents the largest number of species kept in captivity. While there may be tens or even hundreds of different species from another class of vertebrates kept in captivity, there are likely more than 1000 different species of fish that have been maintained in captivity. A visit to a local pet store will often reveal 50 to 100 different species of fish available for sale at any given time. There are three major groups of fish kept in captivity: freshwater, brackish water, and saltwater. The fundamental difference among the three groups is the relative density of the water in which they live. Some fish can move between freshwater to brackish water or saltwater to brackish water, but relatively few fish can live in the two extremes (freshwater and saltwater). The various physiologic and anatomic characteristics among freshwater and saltwater fish will be addressed in the following sections.

Most of the major taxonomic groups are represented by the freshwater temperate species.2 For many, this group represents the species commonly considered as sport fish in the United States. The most common genera of freshwater temperate fishes maintained in captivity include the sturgeon (Acipenser spp.) (Figure 4-1), paddlefish (Polyodon spathula), eels (Anguilla spp.), pike (Esox spp.), bass (Micropterus spp.), sunfish (Lepomis spp.), walleye (Sander vitreus), mullet (Mugil spp.), spotted sea trout (Cynoscion spp.), salmonids, and cyprinids. Although many of these fish are raised by hobbyists, the majority of these species require much larger systems as might be represented in a public aquarium display. Consult an introductory ichthyology text for a more detailed description of the major taxonomic groups of freshwater temperate fish.

Freshwater Tropical Fish Freshwater tropical fish represent the largest numbers of animals typically found in home aquaria. Generally, this group of fishes is readily available for a small to moderate investment

39

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MANUAL OF EXOTIC PET PRACTICE

and can be easily maintained by the novice aquarist or hobbyist. Families of freshwater tropical fish include characins (e.g., tetras: Gymnocorymbus spp., Hemigrammus spp., Hyphessobrycon spp., Paracheirodon spp., Moenkhausia sp.), cyprinids (barbs: Barbus spp.), catfish (e.g., Corydoras spp., Pimelodus spp.), killifish (e.g., Aphyosemion spp., Fundulopanchax spp.), rainbowfishes (e.g., Iriatherina spp., Melanotaenia spp., Glossolepis sp.), gouramies (e.g., Colisa spp., Osphronemus spp.,

Figure 4-1 The sturgeon is a primitive freshwater temperate species. This genus has a wide range of species, representing relatively small sized animals to one of the largest freshwater species of fish. Although they are best known for their eggs (e.g., caviar), at least one species is offered for sale in the pet trade, and several others are routinely maintained in public aquaria.

Trichogaster spp.), livebearers (e.g., Alfaro spp., Poecilia spp., Xiphophorus sp.), and cichlids (e.g., Pseudotropheus spp., Labidochromis spp., Iodotropheus sp., Dimidichromis sp., Copadichromis sp., Neolamprologus spp., Apistogramma spp., Microgeophagus sp., Aequidens spp., Cichlasoma spp.).2,3

Catfish Catfish (e.g., Ictalurus spp., Lacantunia spp., Corydoras spp., Ancistrus spp., Pimelodus spp., Arius spp., Kryptopterus spp., Phractocephalus spp.) represent one of the largest groups of freshwater fishes, with more than 2000 species. Catfish have a cosmopolitan distribution. Catfish are an important group because they serve many different roles, including as ornamentals (Figure 4-2, A), as food fish in aquaculture (Figure 4-2, B), as research animals, and for sport fishing. Most catfish are found in freshwater, although there are two families that contain saltwater species.2,3 Although catfish have a cosmopolitan distribution, more than 50% of all catfish species are native to South America. There is a high degree of variability in the size and weight of these fish, with animals ranging from 10 cm to over 2 m in length and 10 g to over 300 kg in weight. Most species of catfish are nocturnal. Catfish are primarily benthic or bottom-dwellers. Because of their benthic lifestyle, catfish have sensory structures, barbels that assist them with characterizing food and nonfood items and substrate types in a low-light setting.

A

B

Figure 4-2 A, Corydoras sp. B, Ictalurus sp. Catfish represent a diverse group of fishes, with a significant amount of variability in morphology and physiology among genera. Although both are benthic, the catfish in A and B are shaped very differently.

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Figure 4-3 Cichlids represent one of the most popular groups of ornamental freshwater fishes kept in captivity.

Cichlids The cichlids represent one of the most diverse groups of fish, with representation in North America, Central America, South America, and Africa. These fish are prized for their diversity in size, shape, and color. African lake cichlids are one of the most evolutionarily diverse groups in the world, as many different species have evolved within limited ecologic niches (Figure 4-3). With this rapid evolution have come highly variable morphology, feeding strategies, and dietary needs. Although cichlids can look highly variable from genus to genus, they all have a common “break” in their lateral line system. The lateral line is a mechanosensory structure that assists fish with interpreting events in their environment (e.g., shifts in water pressure suggesting a predator is approaching).

Cyprinids Goldfish (Carassius auratus), koi (Cyprinus carpio), and carp (Cyprinus spp.) are members of the largest family (>2200 species) of freshwater fish, Cyprinidae. Cyprinids have the widest area of distribution of any of the freshwater fish. Southeast Asia is the center of origin for this family of fish; however, many species have been introduced around the world and have readily adapted to these new environments.2 The carp are the largest species from this family and may hybridize with goldfish. Koi (Figure 4-4) are colorful domestic mutations of carp that are highly valued by hobbyists and the commercial pet trade. The koi can be divided into two major classifications: the nonmetallic koi and the metallic koi.2 This division is based on typical color patterns, scale types, shape, and size. Goldfish are not koi but rather descendants of the Crucian carp (Carassius carassius).2 Goldfish remain one of the most popular ornamental species and are often found in garden ponds and home aquaria. These fish generally do best if kept

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Figure 4-4 Koi (Cyprinus carpio) represent one of the most coveted groups of fish. These animals are prized for their color, size, and longevity. Their desirability is reflected in their value, with prized individuals selling for tens to hundreds of thousands of dollars.

in cool waters (55°-68° F, 12°-20° C) with abundant plant life and adequate aeration. There are more than 30 varieties of ornamental goldfish available in the pet trade today.

SALTWATER Many marine fish species exist, and, again, it would be impossible to cover the diversity of these fishes in a single chapter. Therefore, this chapter will focus on the species most commonly seen in captivity, including marine tropical fish, marine coldwater fish, and the elasmobranchs (e.g., sharks, skates, and rays).2-4

Marine Tropical Fish There are approximately 23 categories of marine tropical fish. These 23 groups can be arbitrarily divided into four major groups by their feeding attributes and compatibility with other species (Figure 4-5). Most of these groups are well represented in the aquarium trade and in public aquaria. The first category of marine tropical fish is the “rapid eater” and includes the angelfish (e.g., Pomacanthus spp.), damselfish (e.g., Amblyglyphidodon spp.), squirrelfish (e.g., Holocentrus spp.), triggerfish (e.g., Balistapus spp., Balistoides spp.), and groupers (e.g., Cephalopholis spp., Variola spp.).2 These fish do well if kept at low densities in the aquarium or if they are provided a large area where overcrowding is not an issue (e.g., large reef tanks in public displays). Hostile interaction is a major concern for these animals, as they can be very food aggressive.2-4 These animals may be kept together quite readily if provided an abundance of food and a diverse diet. Often they are kept in live coral reef tanks with anemones and other species of animals from the same group.2,3 The slow eaters represent the largest subgroup of marine tropical fish. This group includes, but is not limited to, the

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The last group of animals to be categorized as marine tropicals includes the snappers (e.g., Lutjanus spp., Pristipomoides spp., Nemadactylus spp., Etelis spp.) and grunts (e.g., Haemulon spp.). These are reasonably large, territorial schooling fish that are, for the most part, kept at public facilities. They can best be described as “gluttons,” not only for the manner in which they feed but also for the almost insatiable hunger they exhibit. They are virtually fearless in their feeding habits and will attempt to remove food from the jaws of even large predators.2

Marine Coldwater Fishes

Figure 4-5 Marine tropicals should be housed based on their feeding strategy. This figure represents the typical setting for “rapid eaters.”

anemone clown fish (e.g., Amphiprion ocellaris), parrotfish (e.g., Sparisoma spp.), puffers (e.g., Carinotetraodon spp., Tetraodon spp., Colomesus spp.), surgeonfish (e.g., Acanthurus sohal), wrasses (e.g., Cheilinus spp., Halichoeres spp., Hemigymnus spp., Thalassoma spp.), and trunkfish (e.g., Aulostomus spp., Strophiurichthys spp., Ostracion spp.). Compatibility is an issue with this group of animals, and they do best if maintained in large displays with a large amount of hiding area. Although these fish are grouped based on their feeding strategy, there remains a great deal of variability in the diets of these animals. Another group extreme in feeding habits includes those animals that have difficulty competing for food. This group of animals includes some of the more unusual species, such as the seahorses (Hippocampus spp.), jawfishes (Opistognathus spp., Stalix spp., Lonchopisthus spp.), pipefish (Stigmatopora spp., Lissocampus spp., Corythoichthys spp.), and batfish (Dibranchus spp., Halieutea spp., Halieutichthys spp., Malthopsis spp., Ogcocephalus spp., Zalieutes spp.). Many of these fish are slow swimming and are easily outcompeted by faster swimming fish. Members of this group should be housed together with similar species and monitored closely to ensure that they obtain sufficient calories.

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Although some of the smaller marine coldwater species are used for public display, many are too large to be accommodated in anything other than large aquaria. The majority of the species in this general category are used as commercial food fish. The first group, which represents approximately 70 different species, includes the gadids or cod (e.g., Gadus spp.), haddock (e.g., Melanogrammus spp.), hake (e.g., Urophycis spp.), and pollock (e.g., Theragra spp.). These species are valued as food and for their medicinal value. There are very few public aquaria that exhibit these large species, because they require systems in excess of 1 million gallons of seawater. Tuna (e.g., Thunnus spp., Euthynnus spp., Katsuwonus spp.) have become popular in large open ocean exhibits at public facilities, but only through improved and advanced life support technology and husbandry techniques has it been possible to exhibit this spectacular group of animals. By far the most common category of fish in this group is the flatfish. This group includes, but is not limited to, the flounder (e.g., Platichthys spp., Scophthalmus spp., Limanda spp., Pleuronectes spp., Atheresthes spp.), halibut (e.g., Hippoglossus spp.), and rock sole (Lepidopsetta bilineata). The sablefish (Anaplopoma fimbria) and lumpfish (Paraliparis fimbriatus) are also in this group of fish and are only occasionally represented in public displays.

Elasmobranchs There are more than 350 species of sharks in the world, and only a small percentage are represented as display animals or kept by private aquarists and hobbyists. The species that are found occasionally in home aquaria include the carpet sharks (e.g., Parascyllium spp.) (Figure 4-6), catsharks (e.g., Galeus spp., Scyliorhinus spp.), and horned sharks (e.g., Heterodontus spp.). Most of the other species of sharks are much too large to be kept in anything less than a public aquarium or large commercial facility. Many public facilities display saw sharks (e.g., Pristiophorus spp.) and multiple species of ornamental sharks, including angel sharks (e.g., Squatina australis), dogfish (e.g., Squalus spp., Scymnodon spp., Deania spp., Centroscymnus spp.), and frilled sharks (e.g., Chlamydoselachus spp.). Skates and rays are closely related to sharks; however, they are markedly different in appearance (Figure 4-7). There are both freshwater and saltwater varieties of stingrays. The skates and rays are dorsoventrally flattened animals and usually have at least one venomous spine on the dorsal caudal fin (tail).

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Figure 4-6 The carpet shark (Parascyllium collare) is one of the few species of elasmobranchs that can be kept in captivity by hobbyists.

Figure 4-7 Sharks and stingrays have distinct morphologic features. Note the dorsoventrally flattened shape of the benthic (bottom-dwelling) ray compared to the streamlined pelagic (open water–dwelling) shark.

There are multiple freshwater species (Figure 4-8) that are small enough to be kept by the private aquarist; however, the majority of these animals are also found in public facilities. There are a number of ornamental species of saltwater rays, such as the guitarfish (e.g., Rhinobatos spp.), butterfly rays (e.g., Gymnura spp.), and cow-nose rays (e.g., Rhinoptera spp.), that have been maintained and successfully reproduced in public facilities. Many of the larger, saltwater rays, such as the southern stingray (Dasyatis americana), Atlantic stingray (Dasyatis sabina), eagle ray (Myliobatis aquila), and giant manta ray (Manta birostris), are also now being kept in public aquaria.

UNIQUE ANATOMY AND PHYSIOLOGY When veterinarians begin to work with a new species of animal, it is imperative that they develop a basic understanding of the animal’s anatomy and physiology. A background knowledge of anatomy and physiology will prove beneficial when collecting diagnostic samples or administering therapeutics. The following is a review of unique anatomic and physiologic features

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Figure 4-8 Freshwater stingrays are routinely found for sale in the pet trade. These animals originate from South America.

of fish. (See other resources for additional information regarding this subject.2) Fish are covered with a mucous coat that is produced by cells in their integument. This mucous coat is an important component of the innate immune system and serves as the first line of defense against pathogenic organisms (e.g., bacteria, fungi, and viruses). The mucous barrier contains various sized proteins (e.g., immunoglobulins) that bind pathogens and prevent invasion. If this protective barrier is penetrated, fish have minimal protection against pathogens. To ensure that this barrier remains intact and undamaged, fish should be handled only when necessary. The scales of a fish are located in the dermis and provide protection over the musculature. There are four types of scales found on fish: placoid, ganoid, cycloid, and ctenoid. Placoid scales are found on elasmobranchs, and ganoid, cycloid, and ctenoid scales are found on teleosts (e.g., bony fishes). The ganoid and cycloid scales are common on the more primitive species of teleosts, whereas the ctenoid scales are found on the more evolutionarily advanced fish. The scales serve as a protective armor, and damage or loss of the scales may result in the introduction of opportunistic infections. Handling should be minimized to avoid traumatizing the scales. Teleosts typically have two sets of paired fins (e.g., pectoral and pelvic) and three unpaired fins (e.g., dorsal, anal, and caudal) (Figure 4-9). Fins are used for steering, balancing, and braking. Certain species have modified fins to adapt to certain niches. For example, the anal fin of the knifefish is a large, single fin located on the ventrum of the animal. This fins serves as the animal’s primary source of locomotion. Spines may be associated with some fins and serve as a defense mechanism. The lionfish (Pterois volitans) produces venom that can be injected into a potential predator, causing significant pain and discomfort. Knowledge of the species that produce venom is essential to prevent injury to the handler. Fish may damage their spines when captured in a net. To prevent this, fish may be scooped into a plastic cup or bucket to facilitate removal from the aquarium. The respiratory system of fish is vastly different from the respiratory systems of higher vertebrates (e.g., reptiles, birds, and mammals). Gills are the primary respiratory organs of most fish, although certain species use accessory organs to aid in the absorption of oxygen. Gills serve to absorb oxygen, excrete waste products (e.g., ammonia and carbon dioxide),

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and regulate ion and water balance. Teleosts have four pairs of gills; elasmobranchs can have five to seven pairs. The gills are attached to a bony gill arch, and each gill is comprised of primary and secondary lamellae. The secondary lamellae are the site of gas exchange (Figure 4-10). Exposure to parasites and toxic compounds, such as ammonia, results in excessive production of mucus, which can impede gas exchange. The presenting symptoms of affected animals include rapid opercular movements and gulping for air at the water’s surface; among these animals sudden death may occur. Fish have a simple, inline, two-chambered heart comprised of a single atrium and ventricle. The heart is located ventral to the pharynx and cranial to the liver. Unoxygenated blood is pumped from the heart to the gills, where it is oxygenated and distributed to the rest of the body. Fish possess two portal

Figure 4-9 Fish generally have two sets of paired fins, the pectoral and pelvic, and three unpaired fins, the dorsal, anal, and caudal.

systems: a renal portal system, which drains blood from the caudal musculature, and a hepatic portal system, which drains venous blood from the digestive tract. The lateral line is an important mechanosensory structure used by fish to detect changes in sound waves and water pressure. The lateral line originates on the head, around the eyes and nares, and extends along the lateral body wall. When maintained in captivity, certain groups of marine fish, including tangs and angelfish, may develop head and lateral line erosions. The specific causes of this syndrome have not been elucidated, but dietary deficiency, water quality, and infectious disease are all suspected. Fish can be classified into three different feeding strategies: herbivore, omnivore, or carnivore. The length of the digestive tract can vary depending on feeding strategy. For example, the length of an herbivore’s digestive tract is generally much longer than that of an omnivore or carnivore. The stomach is absent in some species, such as goldfish and carp. Pyloric cecae are found in some species of fish. These structures secrete digestive enzymes and increase the absorptive surface area of the digestive tract. Pyloric cecae are used as a taxonomic indicator in some species. The fish liver is a large structure and is located in the cranial coelomic cavity. The normal color of the liver should be red-brown; however, yellow, fatty livers are a common finding at necropsy. This finding is often the result of diets rich in fats and protein. The fish kidney is a single structure that is comprised of two segments: anterior and posterior. The kidney is located dorsal to the swim bladder along the body wall. The fish kidney serves as both an osmoregulatory and a hematopoietic organ. The anterior kidney and the interstitium of the posterior segment serve as the primary sites for blood cell and immunoglobulin production in fish, as these animals do not have bone marrow. The posterior kidney primarily regulates electrolyte and urine output. Fish that are found in saltwater (hypertonic) environments tend to lose water and absorb salts.

B

A

Figure 4-10 A, Healthy gill. B, Abnormal secondary lamellae. The secondary lamellae are the site of gas exchange and ammonia excretion in fish. Damage to these gills can result in reduced gas exchange, ammonia excretion, and death. (Courtesy Dr. Wes Baumgartner.)

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To prevent dehydration, these fish must drink water and excrete excess electrolytes, such as sodium and chloride, through the kidney and gills. Fish that live in freshwater (hypotonic) environments constantly absorb water by osmosis. To prevent overhydration, freshwater fish excrete large volumes of dilute urine.

HUSBANDRY Environmental Considerations It is easy to recognize aquatic facilities (institutional or pet retail) that have the most disease problems on the basis of their appearance. There is usually an accumulation of trash, dirty exhibits, dead animals, and a general “unhealthy” appearance to the collection. This does not mean that a “clean” facility is free of disease but that they traditionally have fewer and less severe outbreaks of disease.5 The general husbandry and maintenance of the aquatic facility and ecosystems have a direct relationship to the overall health of the animals. Maintaining clean areas behind the exhibits and nonpublic areas is essential to good health practices.3,5 The same can be said for retail aquarium facilities. Accumulation of feces, excess food, and detritus are predisposing factors to poor water quality and can serve as substrates for facultative pathogens. Cleaning and disinfecting seines, nets, buckets, and tanks are imperative to maintain a high-quality health program at a facility. A dynamic team effort is required to maintain a clean facility and healthy animals, but a clean facility will pay dividends by providing a safer workplace, reduced disease incidence, increased production, and generally healthier fish.

Given ideal environmental conditions, ecosystems require a certain amount of space to fully develop, though quantitatively how much space is a debatable matter. Generally, to place an ecosystem in a very large aquarium or other holding space is not a major ecologic problem, although it might be considered an engineering endeavor. The difficulty arises when veterinarians attempt to scale down and include many components in a much smaller space than which would normally occur in the wild. To miniaturize an ecosystem and place it into a small space for observation, education, or research, veterinarians are immediately faced with a major dilemma, which is to scale the miniature so that it can still function as a reasonable facsimile of the wild ecosystem. This question is intimately related to the entire problem of how veterinarians affect wild environments and how they restore them, as well as how they construct their aquaria.5,6

Creating an Aquarium There is little biologic reason for the traditional “box type” aquarium shape; however, the common reason for its existence is associated with availability and mechanical, aesthetic, and economic convenience (Figure 4-11). For many scientists and aquarists, the ease of purchase and setup of a ready-made tank outweighs all other factors when a water-based system is desired. The presence of tank walls that can support benthic communities or allow excessive light is undesirable. A weakly translucent cylindrical tank that minimizes attachment surface for a given volume and has a rotating, cleaning mechanism to keep

A

B

Figure 4-11 Historically, all fish tanks were rectangular (A). More recently, there has been a movement by commercial tank manufacturers to create new tank shapes (e.g., bow front tank) (B). These oddly shaped tanks do little for the aquatic ecosystem, and are primarily for aesthetics.

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the surface free of sediment is highly desirable but very expensive. For the hobbyist, aquarist, or scientist who wishes to construct an efficient model ecosystem, the materials (e.g., glass, plexiglass) are readily available to fabricate any shape or form of enclosure. Only after determining the ideal shape of the desired system, should one be concerned with the aesthetics, viewing, and construction of the aquarium.

TANK SIZE Choosing the correct size of tank is an easy task. The main principle to remember is that bigger really is better. The smallest tank size we recommend is the commercially available 20gallon tank (24″ × 12″ × 20″; 61 cm × 30.5 cm × 50.8 cm); however, the larger tanks provide an even more stable environment for fish. A larger tank also will offer the benefit of a much more liberal air-to-water surface area for the fish. However, with the advancements in life support technology today, it is possible to provide more than adequate life support to maintain large densities of fish in smaller aquaria while also maintaining a greater diversity of animals.4,7,8

TEMPERATURE The term tropical places too much emphasis on the idea of “high temperature” for all exotic fishes. A number of these ornamental fish are not from the tropics, and quite a few from the tropics do not come from particularly warm water. It is important to recognize that a large number of exotic ornamental fish cannot thrive in cool, chilly water (86° F or 30° C), because they need more oxygen than the water has the ability to carry. There is no exact degree of heat that is best suited to each species. Most fish tolerate a 10° F fluctuation over time and can stand a 5° F change in a short period of time (e.g., minutes) without consequence.4,5 It is almost impossible to find a place in nature where the temperature falls into the controlled ranges that veterinarians try to achieve in the captive environment, and it would be reasonable to believe that temperature changes can be beneficial and stimulating to the animals. In practice, the aquarium environment should be geared toward the temperature needs of the species of fish that will inhabit the aquarium. It is best to create fish communities from similar ecosystems and attempt to maintain the temperature range as close as possible to the native waters of those particular species. Temperature changes will occur, but if they occur within the basic guidelines mentioned previously, these changes will be beneficial to the well-being of the animals.

FILTRATION In nature, the waste products produced by fish are diluted into the vastness of the body of water and carried away by flowing water, reducing the potential dangers to fish. In closed systems (e.g., home or public aquarium), wastes and toxins can accumulate to levels that are harmful to fish. To avoid this problem, closed systems must be managed using some form of filtration. There are three primary types of filtration: mechanical, bio-

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Figure 4-12 Mechanical filtration is used to remove detritus from an aquatic system. This type of filtration can be combined with the other methods to improve the overall water quality in a system. For example, this power filter cartridge is comprised of floss (mechanical) and carbon (chemical). Once in the system, it will also be colonized with nitrifying bacteria (biologic).

logic, and chemical. These different filtration mechanisms work independently of one another and can be used in combination.

Mechanical Filtration Mechanical filtration represents one of the original forms of filtration used in the home aquarium. This type of filtration removes organic debris from the water by passing it through a filter material (e.g., floss, fiber, or paper cartridge) (Figure 4-12). The amount of filtration that can be accomplished using this type of filter depends on the type and size of the filter material and rate that the water is recirculated through the filter media. A densely packed fiber or small pore size will restrict the size of waste that can pass the filter media, resulting in less waste in the aquarium. Maintenance of these filters requires cleaning or replacement of the floss or cartridge. Mechanical filtration remains an important method of filtration in home aquaria, outdoor ponds, and public aquaria. This type of filtration is best used with other types of filtration (biologic or chemical) to improve the overall quality of water in the system. When a series of filters is used inline, it is best to place the mechanical filter first. This ensures that the heavy organic material will be removed before contaminating or clogging the other filters (e.g., chemical or biologic-sand filter). Mechanical filtration does have limitations; for example, it is not effective in trapping finite particles or chemicals.

Biologic Filtration Ammonia is the primary end product of protein catabolism in fish. In a closed system, this waste product can be fatal to fish. It was the advent of biologic filtration that led to our ability to maintain large densities of fish in small volumes of water.

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and the organic load on the system. New systems should be started slowly, adding only a small number of fish at a time. Closely monitoring the ammonia, nitrite, and nitrate levels on a daily basis is strongly recommended for new systems. Commercial microbial products (Fritzyme; Fritz Industries, Dallas, TX) are available that can expedite the establishment of the filter by seeding the system with bacteria. Water samples, filter pads, or aquarium substrates from established systems have also been used to seed a tank. However, the addition of these products to a new system may lead to the introduction of potential fish pathogens.

Chemical Filtration

Figure 4-13 Biologic filtration is essential to maintaining fish in closed aquatic systems. There are many different ways to increase the overall surface area in a system to colonize nitrifying bacteria. This filtration method uses plastic balls as a surface for the bacteria.

Biologic filtration is the most common type of filtration used in the home aquarium and outdoor pond, and it comes in many different forms (e.g., under-gravel filters, bio-wheels, sand or bead filters, and wet-dry filters) (Figure 4-13). A biologic filter should be selected based upon the expected load on the system. If a large density of fish is going to be maintained in a system or the fish are going to be fed large quantities of food to ensure growth (e.g., aquaculture), then a large surface area for bacteria is needed. The biologic filter is comprised of nitrifying bacteria. Although there are numerous types, the two most common genera discussed are Nitrosomonas spp. and Nitrobacter spp. Nitrosomonas spp. are important in denaturing ammonia, the primary waste product produced by fish, into nitrite. Both ammonia and nitrite are toxic to fish. Nitrobacter spp. are responsible for further reducing nitrite to nitrate. Once an aquarium and biologic filter are colonized, the system becomes self-sufficient. However, there are several factors that can affect the function of a biologic filter, including temperature, oxygen content, and drugs/therapeutics. Nitrobacter spp. are not cold tolerant. Therefore, in outdoor ponds when the water temperature drops below 65° F (18.5° C), nitrite will not be converted into nitrate as rapidly. During those times when the water temperature may drop below 65° F, fish should be fed less food to reduce the load on the system. Nitrosomonas spp. and Nitrobacter spp. are aerobic bacteria. In the well-aerated home aquarium, oxygen levels are often adequate for the bacteria; however, in outdoor ponds, oxygen levels can become depleted on warm summer nights. To reduce the likelihood of biologic filter failure, it is recommended that outdoor ponds be aerated during warm, summer days and nights. A biologic filter requires time to become established. The amount of time depends upon the temperature of the water

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There are numerous types of chemical filters in the pet trade. Chemical filtration refers to those filters that remove toxic compounds by binding them or converting them into nontoxic substances. The original form of chemical filtration was activated carbon. Carbon serves as a nonspecific binding agent for a number of different substances. When the binding sites on the carbon are full, they no longer act as filters and need to be replaced or cleaned. Other forms of chemical filtration are more specific, such as the resins that only bind ammonia. There are other forms of chemical filtration, such as ultraviolet (UV) sterilizers and protein skimmers, that alter or trap compounds. UV sterilizers expose compounds to short wavelength light, altering their form and rendering them harmless. UV sterilizers can be used to control certain pathogens and algae. A UV sterilizer has a UV bulb encased in a waterproof sheath within a cylinder. As water passes through the cylinder, the water is exposed to UV light, which can alter the DNA or RNA of the organism. The amount of time that it takes for the water to pass the bulb and the bulb wattage determine the effectiveness of the UV sterilizer. A low-wattage bulb in a short cylinder will have little effect on pathogens. These systems have also been used with great success at controlling algae and pathogens in outdoor ponds. Protein skimmers trap proteins in bubbles so that they can be separated from the water and removed. Chemical filtration, in combination with mechanical or biologic filtration, can improve the water quality dramatically, creating a “healthy” environment for fish.

LIGHTING Correct lighting for the aquarium system depends on several factors, including the quality of the light emitted from the selected light source. Full-spectrum lighting is preferred. This type of lighting provides the three primary light spectrums: ultraviolet, visible, and infrared. The various spectrums can be affected by water depth, with certain spectrums (e.g., ultraviolet) being removed in the epilimnion (upper water level). Another important factor is the function of the system. If the tank is going to house deep-water African cichlids, lighting becomes less important; however, if the lighting system is needed for a reef tank that is going to be stocked with corals, a significant quantity of high-quality full-spectrum light will be required. The primary disadvantages associated with excess lighting are algae overgrowth and overheating the water in

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smaller systems; otherwise, it is virtually impossible to have too much light in an aquarium system.4 A 12 hour photoperiod is considered appropriate for most fish. For those individuals interested in breeding fish, the photoperiod should be lengthened to mimic a spring/summer season.

scaling. For the aquarist, fine sediment bottoms should not be ignored. Given their full reign, with the proper environment and biota available, they can be important buffering systems and provide necessary stability for a healthy environment.

SUBSTRATE

There are a myriad of commercial accessories available for aquaria today. These accessories range from simple items, such as heaters and aeration devices, to some of the most advanced biotechnology available in the field, such as chillers and wave machines. Most commercial accessories available on the market are well made, and the deficiencies have been minimized over the years. When choosing a particular item, selection should be based on function not brand. For example, with a heater, it is important to match the wattage of the instrument to the volume of water to be heated, taking into account the general temperature required for the environment in which the aquarium is to be located.4 There are several types of heaters available, including basic submersible heating elements, solar units, flow through units, or a combination of similar systems. Water filtration units should be selected based on desired type of filtration, the volume requirements of a particular system, and its intended use. For example, a sand filtration unit should be selected on the basis of the volume of water to be used, the turnover rate expected for that volume of water, and the frequency with which the sand filter must be backwashed. Sometimes it is preferable, if not necessary, to combine several methods of filtration (e.g., under-gravel filtration in combination with sand filtration and power filtration). This enables the water to be “cleaned” using multiple methods. The use of multiple filters inline is particularly important in a public facility where a large water volume and bioload are used. Most saltwater aquaria utilize some form of ozonator in combination with a protein skimmer. Protein skimmers utilize the age-old technology that has been used by sewage treatment plants for years: Minute air bubbles are passed through water with a high organic waste content, and the protein is captured and trapped as foam at the surface of the water. The greater the degree of organic pollution there is, the more stable the foam will be. The skimmer collects the foam, and the foam is then collapsed to a liquid and removed from the aquarium without tainting the water.

Historically, aquarists have tended to ignore the substrate of an aquatic system, reducing it to a noninteracting element. In the aquaria of past decades, “clean,” relatively inert gravel and under-gravel filters were used to provide environments for all but the most specialized natural situations. In nature, with the exception of gravel bottoms in relatively unproductive hard rock mountain streams or sandy beaches with sandstone composition, rarely is the substrate material neutral. In most aquatic and marine environments, soft substrates are rich in organic reservoirs and harbor a myriad of important invertebrates and microbes that support rich plant growth. Limestone substrates control water chemistry, and reef corals and rocks determine the very character of the organisms growing on the surface of the reef. The interest in so-called live rock in coral reef tanks in recent years is beginning a tendency to replace sterile environments with live ecosystems. The addition of “trickle trays” with calcium carbonate pebbles also shows a developing interest in further elucidating the carbonate cycle and control over the pH. Conversely, acid, black water streams are most likely to occur in granite or sandstone areas where the natural acidity of the rain and tannic acid from the forest litter cannot be neutralized. To recreate this environment, the aquarist is advised to use hard rock and silica sand. Coarse sand or gravel is perhaps the most difficult of benthic environments for organisms to adapt to, and within sand and gravel habitats there are relatively few common species. In a stream or small lake that is not large enough for significant wave activity, or in a bay or coastal lagoon along a sandy coast, the sediment becomes progressively finer from gravel, to sand and silt, to a soupy, silty-clay mud substrate. To remain sand, the bottom must stay in constant motion; therefore, special adaptations are required by any organism to adapt to it. Even bacterial numbers tend to be limited in sand and gravel because their organic substrates are often washed out.9,10 It can be difficult to maintain a sandy, shore style substrate in a closed, captive system, as it can have a rather long profile in the energy regime required to maintain the sand. It would be impossible to recreate a wave-break, sandy scenario within a miniature system unless the benthic community is the only one desired. Traditionally, the general approach to filtration followed by most aquarists was to avoid the natural detrital processes in an aquatic system and to keep bacteria in filters that mimicked the benthic community. However, in a filter, the variety, density, and capability of the bacteria are limited. Thus, aquarium procedures of the past have tended to short-circuit the natural cycling processes, which resulted in the loss of valuable energy to the many members of the community. Aquaria that do not have a fine sediment component should have a separate sediment trap that can be partially drained of sediment, especially if it is intended to drive a system faster than normal for

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ACCESSORIES FOR AQUARIA

WATER QUALITY ANALYSIS The aquatic environment is a very complex system that is subject to constant physical and chemical changes. A waterassessment program should be devised for monitoring an aquatic ecosystem. Many factors need to be taken into consideration when designing a water quality monitoring program, including the volume of water in the aquarium/aquaria, the number and type of animals that will be present, the type of life support system, and the period of time the aquarium has been set up. Ideally, a water-assessment program should be established before designing and setting up a system, keeping in mind that most of the water quality analysis is done to keep its inhabitants healthy.8

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Chapter 4

Ornamental Fish

Water testing should be done daily for new aquaria until the system has cycled. When collecting a water sample, it is important to collect a mid-water sample, as samples closer to the surface or substrate will reflect more extreme values.2,3 Portable water analysis kits use premeasured reagents and simple analytic methods that may compromise the degree of accuracy. However, they are suitable for most private aquarists and small backyard ponds. Most large, public aquaria use more advanced analytic methods, such as mass spectrometry, saturometers, and sophisticated chemical analysis, for evaluating water quality. The acquisition and utilization of many of these processes and techniques are beyond the scope of what most private aquarists can afford. Commercial water quality laboratories can perform a complete water analysis for a modest price, and it may be a good idea to submit a sample to a laboratory for a baseline measurement. This method also offers the added benefit of comparing standardized values with the parameters derived from veterinarians’ own testing methods. Water quality is very important to the health of a fish, and poor water quality can prove fatal. There are two types of systems that can be used: open and closed. In open water systems, the water in the aquarium is continually replenished using a fresh water source. An individual who lives near the ocean may collect seawater for a home aquarium, although that is not recommended because of potential contaminants in the water. Open water systems are rarely used because they are labor intensive and require regular exchange of the entire system. The majority of home aquaria utilize closed recirculating systems, which recirculate the same water over and over again using a filter. In the closed system, fresh water is added only after evaporation or at the time of a water change.

Ammonia, Nitrite, and Nitrate Ammonia is produced in fish as an end product of protein catabolism. This waste product is primarily excreted via the gills, although some excretion in the feces also occurs. Ammonia is also generated in the aquatic system from the breakdown of uneaten food and detritus. Ammonia nitrogen can occur in two forms: ammonium (NH4+) and ammonia (NH3). Ammonia is the more toxic form for fish.11 The relative concentration of each form varies with pH and water temperature.12 Ammonia is soluble in water, and minimal amounts are lost through evaporation. In a closed system, such as an aquarium or backyard pond, ammonia levels can build up to toxic quantities (>1 ppm). Even low levels of ammonia can be toxic to the gills and skin, resulting in increased susceptibility to infections. Fish suffering from ammonia toxicity appear irritated, gasp at the water’s surface, and rub against rocks in the aquarium as a result of the irritation caused by the toxin. Ammonia levels should be monitored closely in closed systems, especially those with high fish densities. Testing for ammonia should be done weekly using a standardized commercial test kit, which is available at local pet retailers. In an established system, ammonia levels should be zero. If the ammonia levels begin to rise, then the system should be reevaluated. Overfeeding and overstocking an aquarium can overburden the biologic filter. Severe temperature fluctuations and insufficient oxygen levels may

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49 also result in a significant loss of the biologic filter. This is especially common in ponds that have significant summer algal blooms. New systems require time to become established, and new fish should be added gradually to prevent an overload of the biologic filter. Because ammonia is a common waste product in the aquarium, most problems can be prevented by limiting the numbers of fish in the aquarium and regulating the amount of feed being offered. A general rule of thumb for feeding fish is to only offer what they can eat in a 2- to 5-minute period. In cases where ammonia levels are creating problems, the first recommendation is to remove 25% to 50% of the water from the system and replace it with fresh, dechlorinated water. There are commercial products available that can chelate the ammonia source, but they are only a temporary solution. The primary cause of the elevated ammonia levels must be diagnosed and corrected. Ammonia is a colorless, odorless substance that can cause significant mortalities in a home aquarium. Most inexperienced aquarists tend to single out infectious diseases when they experience fish losses; however, poor water quality (e.g., excessive ammonia or nitrite) is often a primary/secondary cause of mortalities in these animals and should be tested on a regular basis. The nitrogen cycle eliminates ammonia by converting it to less toxic compounds. The first step in the cycle is to convert ammonia to nitrite. Nitrosomonas spp. are the primary bacteria associated with this process. Unfortunately, nitrite is also toxic (>0.1 ppm) to fish and can be rapidly absorbed across the gills. Affected animals may develop a methemoglobinemia and have a characteristic “brown blood.” This blood dyscrasia leads to reduced erythrocyte oxygenation and respiratory compromise. Fish with nitrite toxicity may behave similarly to fish with ammonia toxicity, and be found gasping for air at the water surface and die suddenly. When fish show clinical signs associated with nitrite toxicity, they should be removed from the toxic water and placed into a fresh, dechlorinated, welloxygenated system. A significant water change (25%-50%) should be made in the original aquarium or pond and the biologic filter reestablished. Salt can also be used to diminish the toxicity associated with salt. The second step of the nitrogen cycle occurs when Nitrobacter spp. oxidizes nitrite to nitrate. Nitrate levels less than 0.5 ppm are generally regarded as safe; levels less than 5.0 ppm are associated with stress and may predispose fish to opportunistic infections; levels greater than 10 ppm are considered toxic for some species. Reports of nitrate toxicity are rare in freshwater and saltwater fish, but at elevated levels they may be stressful and predispose the animals to opportunistic pathogens. Nitrate is utilized by plants and algae as a food source. Nitrate can be removed from an aquatic system by performing regular water changes. Ammonia and nitrite levels in an aquatic system may rise soon after treatment of the water with antibacterial compounds or a reduction in water temperature. Antibiotics added to the water are nonselective and may kill both pathogenic and commensal organisms. If these compounds kill enough of the

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50 bacteria associated with the biologic filter, then oxidation of ammonia and nitrite may stop. Nitrosomonas spp. are more temperature tolerant and will recolonize before Nitrobacter spp., so ammonia levels should be expected to decrease before nitrites. Therefore, elevated nitrite levels are often detected soon after a reduction in water temperature.

“New Tank” Syndrome “New tank” syndrome is a common occurrence with beginner aquarists and primarily occurs when fish are overstocked in a new aquarium. If large numbers of fish are added to an aquarium, and the biologic filter is not established, the system will be unable to eliminate the ammonia produced by the fish. In most cases, the owners report an acute mortality event with clinical signs consistent with ammonia and nitrite toxicity. These problems can be prevented if the new owner is patient and realizes the importance of providing a break-in for the filter (4-6 weeks). Fish should be stocked gradually, usually one to two fish per week. A standard rule of thumb for a freshwater stocking density is 1 to 1.5 inches of fish per gallon of water; in saltwater systems, the stocking density should be 2 to 2.5 inches of fish per gallon of water. With the advent of new filtration systems, stocking densities will continue to increase; however, if the filter becomes compromised or fails, the results would be disastrous.

Oxygen In the aquatic system, oxygen diffuses into water at the surface when the surface tension of the water is broken. For home aquaria, this occurs regularly when external filters are used that “drop” the water back into the tank like a waterfall or when airstones are used. The amount of available or dissolved oxygen (DO) within the system can be measured using special equipment. In most cases, a DO greater than 5 ppm is sufficient to maintain fish. For the home hobbyist, oxygen depletion is generally only a major problem with outdoor ponds during the summer months. During the day, plants produce their own food (photosynthesis) by removing carbon dioxide from the water and using energy produced by the sun. As plants make their food, they release oxygen into the water. During the night when plants or algae cannot undergo photosynthesis, they actually consume oxygen. In ponds with a large number of plants or algae, the oxygen levels in the water can fall to dangerously low levels (0.1 mg/L Species dependent 5.5-6.5; 8.6-9.5 >78 mmHg Unknown Unknown >5 ppm

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Chapter 5

79

Amphibians

SEMITERRESTRIAL (STREAMSIDE ANIMALS) Semiaquatic animals need standing water as well as areas to haul out. By providing this, it is possible to create humidity gradients that enable the animal to regulate its moisture. Aquaria can be tilted or accessories placed such that the bottoms have standing water and top layers are drier and well drained.

FOSSORIAL AND TERRESTRIAL Fossorial and terrestrial vivaria are similar in every regard, except that there should be deeper soil for fossorial animals. Standing water receptacles are reasonable in these settings, but the water level should not be deeper than the animal is tall. Moisture gradients in the soil are very important for fossorial animals, and this is easily accomplished by tilting the aquarium to retain different levels of moisture. Leaf litter placed on the surface will allow the animal to forage while remaining covered. Live plants are not recommended for fossorial species, as the plants may be uprooted when the amphibian forages.

ARBOREAL The enclosure should be tall with accompanying tall branches and plants. It is important to provide an adequate number of plants to provide ample hiding places.

Creating a Vivarium Glass tanks are commonly used to house amphibians. They are well accepted because they allow excellent visualization of the amphibians, are relatively inexpensive, and hold moisture. The primary disadvantage associated with them is that they are not well ventilated. One way to improve ventilation is to have a secure, well-ventilated lid. The lid also helps prevent escape. Plastic tanks are also appropriate for making vivaria and may be more flexible for modification, though scratches can occur that obscure direct visualization of the animal. Lids that are hinged allow servicing to a portion of the enclosure while maintaining cover for the rest of the area. Solid plastic or glass lids should be considered when maintaining a high humidity is important. When these types of lids are used, it is important to still place some ventilation holes in them to prevent stagnant airflow. Volatile organic compounds, such as those found in glues, need to be cured appropriately (under strict manufacturer conditions) before being used around amphibians; otherwise the fumes can leach into the environment and be irritating to the skin and respiratory epithelium. Deionized water should not be used in an amphibian’s vivarium, as it can disrupt osmolarity. Chlorinated tap water also is not recommended. Distilled water or reverse osmosis water is ideal.

BOX 5-1

Environmental Temperature Ranges for Amphibians Based on Natural Habitat

Tropical lowland Tropical montane (moist, cool, coniferous strata) Subtropical Temperate, summer Hibernation Aquatic, tropical lowland Aquatic, tropical montane Aquatic, subtropical Aquatic, temperate stream (summer) Aquatic, temperate pond (summer)

24°-30° C 18°-24° C 21°-27° C 18°-24° C 10°-16° C 24°-30° C 18°-24° C 21°-27° C 16°-21° C 18°-24° C

should be provided vertical enclosures. Aquatic and fossorial species should be provided an enclosure with a large surface area (length × width).

TEMPERATURE The environmental temperature range provided an amphibian should be based on its natural habitat (Box 5-1).6,7 There should always be a gradient with a range of 5° to 8° C. When changing water, make sure the temperature matches the enclosure. Basking spots are necessary, and ceramic heaters (which produce no light) or spotlights can usually provide the necessary temperatures. Care and diligence are required for appropriate wattage and distance. Spot heaters are best placed outside of the enclosure to avoid trauma via thermal burns. Do not use heat rocks for amphibians. Aquarium heaters may cause thermal burns for caecilians if they wrap around these devices. To prevent this, place mesh or PVC piping around the heater to prevent contact. It is ideal to use a thermometer that dually measures the warmest and coolest spot; the more advanced thermography units allow detection of temperature gradients as well.9 Hibernation is important in some species for stimulating breeding, but it is unknown whether it is necessary for long-term health and success in captivity.6 Many salamander species prefer cooler temperatures,10 and some vivaria might require chillers.

HUMIDITY A fine mist can be sprayed into the vivarium several times daily (manually or via misting systems). When doing so, distilled or aged water should be used. An air stone in a bowl of water or live plants can also be used to help maintain the environmental humidity. Relative humidity above 70% will suit most amphibians. Tree frogs have a natural behavior to adduct their heads and press against surfaces to retain water—if they do this exclusively, the environmental humidity is not adequate. Conversely, waxy tree frogs may develop dermatitis with excessive environmental humidity.6

ENCLOSURE SIZE

LIGHTING

Enclosure sizes are highly variable and largely dictated by the environmental re-creation that is attempted. Arboreal species

Full-spectrum lights are generally recommended for amphibians, but most do not provide radiation in the wavelengths

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80 consistent with vitamin D3 synthesis. Ultraviolet B (UVB) (290-320 nm) radiation may be important for vitamin D3 metabolism in amphibians, but the role of UVB radiation has not been fully examined. I prefer to be conservative and recommend the use of full-spectrum lights. Replacement of the bulbs is recommended every 9 months to ensure that the appropriate wavelength is emitted. UVB does not transmit through glass or plastic, and the depth at which it transmits is typically no greater than 9 to 18 inches away from the source. Care must be taken that the lighting does not adversely affect the animals. Low-level nocturnal lighting (moonlight simulation) may be useful for nocturnal species to ensure that the animals are not startled when the lights are turned on or off. Dimming lights gradually is ideal. Amphibians should be provided with a natural photoperiod of 12 hours of light and 12 hours of darkness. If animals are going to be bred, the light cycle may need to be altered to mimic the normal reproductive season of the animal.

SUBSTRATE Substrate selection is an important consideration for a vivarium. When considering different types of substrates, it is important to try to mimic an amphibian’s natural habitat. Smooth or small pebble gravel may be used as a substrate, but it can become a gastrointestinal foreign body if it is ingested. This can be prevented by using large pieces of gravel (e.g., bigger than can fit in the oral cavity) or feeding animals away from such substrates. Soil substrates should be organically rich and pH balanced (neutral). For burrowing caecilians, soil depth should be 3 to 10 cm.5 For fossorial species, there should be a moisture gradient and the soil should be loose enough to allow for tunnel formation. Soil should be replaced every 2 to 3 months. Substrates that need to be sterilized should be baked. Soil and leaf litter substrates should be sterilized to prevent arthropod and helminth infestation. To sterilize soil, bake at 200° F (95° C) in a thin layer (1 m).

counts can be performed on samples stored in either anticoagulant, although in some species the choice of anticoagulant may have an affect on the quality of the blood smear. I usually perform complete blood counts from EDTA samples and plasma biochemistry testing from lithium heparin samples. Blood smears should be made from fresh (no anticoagulant) blood samples. To reduce the likelihood of white blood cell damage during the smearing process, I premix the blood sample with 22% albumen (Gamma Biologicals, Inc., Houston, Texas). One drop of albumen to 5 drops of blood can significantly reduce the number of destroyed white blood cells in birds and reptiles (Mitchell, unpublished data). Historically, white blood cell estimates were done using the eosinophil unopette technique (Becton Dickinson Co., Franklin Lakes, New Jersey). Unfortunately, this commerical product is no longer available; however, the stain, phloxine B, can still be acquired and used to perform a white blood cell estimate.

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Chapter 7

Snakes

Figure 7-17 The heterophil of the snake has fusiform, eosinophilic stained granules. These cells are associated with the acute inflammatory response of snakes.

Plasma biochemistries can be performed using a variety of different methods. I prefer the Abaxis VetScan (Abaxis, Inc., Union City, California), because the analyzer only requires 0.1 ml of whole blood to perform a complete biochemistry analysis. Another benefit of the system is that it does not require any time to calibrate the machine.

HEMATOLOGY Snake erythrocytes are nucleated. Although snakes generally have fewer erythrocytes than higher vertebrates, they are longer lived (600 days).3 Snake hematocrits tend to fall between 20% and 35%, although clinically normal snakes may have a hematocrit less than 20 or more than 35. Each individual should be evaluated on a case-by-case manner. Erythrocyte numbers can be affected by environmental conditions, reproductive status, season, and nutrition. Heterophils are acute inflammatory cells that are frequently identified in snakes with inflammatory leukograms. Heterophils have oval to lobed nuclei and are filled with eosinophilic fusiform granules (Figure 7-17). Toxic heterophils are a common finding in chronic inflammatory responses. It is important to determine if a species depends on its lymphocytes or heterophils as the predominant cells. In species where the heterophil is the predominant cell type, the heterophil-tolymphocyte ratio should be between 1 : 1 and 3 : 1. Eosinophils are granulocytic cells that are often confused with heterophils. Eosinophils are filled with circular eosinophilic granules. Snakes have the largest eosinophils of all the reptiles. Although eosinophils are usually associated with allergic reactions and parasites, they likely serve other functions in an inflammatory response. Eosinophil counts may vary with stress and seasonal effects. Eosinophils are generally less than 0.5 × 103 cell/μl. Basophils are a generally the smallest granulocytes in snakes. The nucleus of the cell is often obscured by the basophilic

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147

Figure 7-18 The snake lymphocyte is similar in appearance to the mammalian and avian lymphocytes. The classic lymphocyte has an acentric nucleus and high nuclear-to-cytoplasmic ratio.

metachromatic granules that fill the cell. The exact function of basophils in snakes in not known; however, the cells are considered to play a role in histamine release. Basophils are generally less than 0.5 × 103 cell/μl. Snake lymphocytes look very similar to avian and mammalian lymphocytes, with an acentric nucleus and a basophilic cytoplasm (Figure 7-18). Reptile lymphocytes function in a similar manner to lymphocytes from higher vertebrates, including moderating the immune system and producing globulins. Lymphocytes may be the predominant white blood cell in some species of snakes. Snake lymphocyte counts less than 1.5 × 103 cell/μl should be considered lymphopenic. Lymphocyte counts may vary with stress and seasonal effects. Monocytes are the largest white blood cell encountered in snakes. The snake monocyte is similar in appearance to the monocytes described in mammals, a large nonsegmented nucleus and a blue-gray cytoplasm (Figure 7-19). Azurophils are monocyte-like but have an azure staining cytoplasm. Monocytes are primarily associated with chronic inflammatory responses and antigen processing. Absolute monocyte counts greater than 1.5 × 103 cell/μl are generally associated with a chronic inflammatory response. Reference material for hematology in snakes is sparse. Because snake hematology can be affected by species, environmental, seasonal, and hormonal influences, interpretation of reference ranges should be made taking this information, relative to the sample population, into account. An inflammatory response in a snake may be associated with infectious agents, foreign bodies, neoplasia, traumatic injuries, and toxic exposures. In general, snakes can have an inflammatory response without an overt leukocytosis (>15 × 103 cells/μl). In many of these cases, the leukocyte count may be within established reference ranges (5-15 × 103 cells/μl), but the animal will have a severe monocytosis or azurophilia (>1.5 × 103 cell/μl).

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Figure 7-19 Snake monocytes are associated with the chronic inflammatory response. In some cases it can be difficult to discern a large lymphocyte from a monocyte. However, the monocyte cytoplasmic-to-nuclear ratio is higher than that of the lymphocyte.

PLASMA BIOCHEMISTRY ANALYSIS Because ophidians have evolved to mask their illness, characterizing a disease process in a snake generally requires more than a physical examination. Plasma biochemistry analysis is an important diagnostic tool and can be used to derive important physiologic information regarding a patient. Reference material for plasma biochemistries for snakes is also sparse and variable.4 In snakes, plasma biochemistries can be affected by environmental conditions, reproductive status, season, and nutrition. Blood glucose is highly variable in snakes. Some species will be clinically normal with blood glucose values of 40 to 50 mg/ dl. In general, blood glucose levels in snakes range from 50 to 150 mg/dl. Calcium and phosphorus values may vary with diet and reproductive status of a female snake. In general, the calciumto-phosphorus ratio in snakes is approximately 1.5-2 : 1. Female snakes mobilize calcium and phosphorus during the reproductive cycle and can develop significant hypercalcemias (>13 mg/dl) and hyperphosphatemias (>6 mg/dl). Snakes with renal impairment frequently have an inverse calcium-tophosphorus ratio. Hyperphosphatemia may occur with refeeding syndrome. Reference ranges for the electrolytes are extremely variable. Potassium levels may vary from 2 to 6 mEq/L. Hypokalemia may occur with dietary restriction, diarrhea, and renal disease. Hyperkalemia may occur with re-feeding syndrome. Sodium levels are generally 135 to 170 mEq/L. Hypernatremia can occur with dehydration or increased salt intake. Hyponatremia is a common finding in snakes with chronic diarrhea. Chloride levels are generally between 90 and 130 mEq/L. Chloride levels, like sodium levels, increase with dehydration and dietary intake and decrease with diarrhea.

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Aspartate aminotransferase (AST) is an enzyme presumed to be in all body tissues in snakes. However, an elevated AST level (>250 units/L) is generally associated with muscle or hepatic disease. Creatinine kinase (CK) is generally derived from skeletal, smooth, and cardiac muscle. An elevated CK (>300 U/L) is a strong indicator of muscle necrosis. Alanine aminotransferase (ALT) is presumed to be derived from many tissues in the body; however, unlike with AST and CK, there are no specific tissues that appear to release this enzyme in greater concentrations. Total protein is comprised of pre-albumin, albumin, and the globulins. Protein concentrations generally increase with dehydration and inflammatory responses. Serum electrophoresis may be used to further differentiate the globulins into specific fractions. Hypoproteinemia is a frequent finding in snakes with chronic anorexia and renal disease. Hyperproteinemia is generally associated with dehydration and inflammatory disease. Uric acid is the end product of protein catabolism in most snakes. In general, uric acid levels are 15-17 mm) and assess follicle appearance to predict ovulation (Figure 7-22). Repeated measurements should confirm that the follicle is growing in size.

Microbiology Bacterial and fungal diseases remain significant pathogens for captive snakes. Many of the infectious agents encountered in captive snakes are directly related to inappropriate husbandry. Snakes that are not provided an appropriate thermal gradient, or are housed on a soiled substrate, may become exposed to opportunistic bacterial and fungal pathogens. Many of the bacterial and fungal pathogens diagnosed in snakes are ubiquitous. Therefore, the techniques used to isolate these pathogens in other species (e.g., mammals) also can be used for snakes. When attempting to culture bacteria or fungi from reptiles, some advocate that the cultures should be maintained at the same environmental gradient under which the snake is housed. However, I have found that the pathogens are more likely (and more quickly) to be isolated using standard techniques (37º C). It is important to remember that many of these organisms are capable of infecting a range of hosts, including both ectotherms and endotherms. Veterinarians interested in isolating a specific pathogen can request this of their diagnostic laboratory. Certain species, such as Salmonella spp., have specific requirements for growth. If these considerations are not taken into account before the sample is submitted to the laboratory, the organism may be outcompeted by more common organisms.

Parasitology

Figure 7-20 Dorsoventral radiographs in snakes may have limited value because of the prominence of the spine.

Endoparasites are a common finding in captive snakes. Although the majority of the parasites are associated with the gastrointestinal tract, there are others that can be found in the respiratory system (e.g., acanthocephalans) and integument (e.g., aberrant cestodes). When deciding which diagnostic tests should be used to confirm a diagnosis, it is best to consider the origin or location of the parasite. The presence or absence

Figure 7-21 The lateral survey radiograph provides the most information regarding the viscera. In this image the heart (thin arrow), lung (thick arrow), and liver (dashed arrow) can be seen.

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MANUAL OF EXOTIC PET PRACTICE

Figure 7-22 Ultrasound can be used to assess the reproductive status of a snake. Follicle size can be measured and used as a method for determining when to breed snakes.

Figure 7-23 When passing a tube through the oral cavity and into the esophagus, it is important to avoid the glottis (arrow). The glottis is located on the floor of the oral cavity. Snakes have conscious control over the glottis.

of endoparasites was once considered useful in characterizing whether a snake was captive born or wild caught; however, this approach is often of limited value as captive-bred and imported snakes are routinely mixed in pet stores, reptile shows, and private collections. The diagnostic tests used to confirm the presence of gastrointestinal parasites in other vertebrates are appropriate for snakes. Fecal samples should be evaluated using both a direct saline smear and a fecal flotation. The direct saline smear can be used to confirm and define bacterial and protozoal densities, whereas the fecal flotation is used to confirm the presence of trematodes, cestodes, and nematodes. Regurgitated food also can be evaluated using these techniques. If Cryptosporidium serpentis is suspected, then an acid-fast stain and/or Merifluor assay (Meridian Diagnostics Inc., Cincinnati, Ohio) should be considered. Any snake found to have a negative sample should be reevaluated, as all of these parasites can be transiently shed. In cases where there is no fecal output or regurgitant, a gastric lavage and/or enema can be done. A gastric lavage can be done using a red rubber feeding tube or stainless steel gavage tube and saline. This technique requires a minimum of two individuals: one to restrain the snake and another to collect the sample. Additional assistance may be required for larger specimens. The tube should be premeasured to ensure placement into the stomach. The stomach is approximately one half the distance from the snout. The tube can be inserted into the mouth without a speculum. As the tube is being passed through the oral cavity, it is important to avoid passage into the glottis (Figure 7-23). A small quantity of lubricant can be placed on the tube to facilitate passage down the upper gastrointestinal tract. Be sure not to place excessive quantities of lubricant on the tube, as it may be aspirated into the trachea. Once the tube has been passed to the desired level, the saline solution should be instilled into the stomach. I recommend infusing no more than 5 to 10 ml/kg of saline into the stomach for this procedure. After 10 to 15 seconds, the

fluid should be aspirated back into the syringe. Once the sample has been collected, the solution should be centrifuged at 1500 rpm for 5 minutes to concentrate any pathogenic organisms. Samples should be submitted for cytologic and microbiologic testing based on the clinician’s differential list. An enema can be used to collect samples for direct visualization or as a stimulus for an animal to defecate. A red rubber feeding tube can be used for this procedure. Passing a tube into the colon of a snake is easier than it is in lizards, as snakes have no urinary bladder. The tube should be premeasured to the distance it will be passed. It is often possible to palpate feces in a colon of a snake. The tube should be lubricated to facilitate passage. I usually instill warm (snake body temperature) soapy water or 0.9% saline at a rate of 5 to 10 ml/kg. The samples should be reviewed using a direct smear and fecal flotation. Parasites associated with the respiratory tract can generally be diagnosed via oral examination, fecal examination, or endoscopy. The eggs of lung parasites are generally passed through the trachea and into the oral cavity, where they are swallowed and passed with the feces. Sputum samples can be reviewed under light microscopy for eggs, or a direct saline smear/fecal flotation can be done to evaluate the sample for parasite eggs. A flexible endoscope can be inserted into the trachea of a snake to directly examine the lung(s) for the presence of parasites, or a rigid endoscope can be used to examine the lungs via a coelioscopy. The length of the endoscope can be a limiting factor in these cases. Ectoparasites, including mites and ticks, are a common finding in wild-caught snakes. Adult ticks are generally easy to identify, whereas nymphs and larvae may be more difficult to visualize. Ticks are generally found around the eyes, oral cavity, and vent, although they can occur anywhere on the body. Mites are smaller in stature than ticks and thus can be more difficult to identify. Snake mites are generally found in the same areas where ticks are found, although I routinely find

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Snakes

then its immune function may become suppressed. Treatment should be based on antimicrobial sensitivity testing.

FUNGAL

Figure 7-24 The gular fold should be inspected for ectoparasites.

them in the gular fold as well (Figure 7-24). Mites or ticks collected from a snake should be examined under a light microscope to be keyed to species.

COMMON DISEASE PRESENTATIONS Infectious Diseases BACTERIAL Bacterial infections remain a common problem in captive snakes. The majority of the problems encountered with these animals are related to opportunistic Gram-negative bacteria, including Escherichia coli, Pseudomonas spp., Aeromonas spp., Citrobacter freundii, C. braaki, Proteus spp., Serratia spp., Klebsiella spp., Enterobacter spp., and Salmonella spp. Although less common, Gram-positive bacteria also have been associated with clinical disease in these animals and should be considered in a differential list. Bacterial infections can affect any system. However, most infections tend to be associated with the respiratory, gastrointestinal, and integumentary systems. Most bacterial infections are diagnosed using standard microbiologic techniques. Because certain organisms can become injured during sample collection and transport, or are in limited numbers, an additional enrichment period may be needed. However, some organisms, such as Chlamydophila spp. and Mycobacterium spp., are difficult to isolate using standard techniques. With the advent of new molecular techniques, such as the polymerase chain reaction technique, these fastidious organisms can generally be characterized based on their 16sRNA. Most bacterial infections in snakes are preventable. Snakes that are maintained at inappropriate environmental temperature ranges may be predisposed to opportunistic infections. Reptiles are ectotherms and rely on their environmental temperature to regulate their core body temperature. If a snake is unable to achieve an appropriate core body temperature,

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Fungal dermatitis is a common finding in snakes maintained in moist to wet environments with poor ventilation. These conditions are frequently observed after hibernation; thus it is important to closely monitor snakes during this period. It is not uncommon to isolate multiple species of fungi from a lesion. Reptiles may harbor a large number (0-15) of different saprophytic and pathogenic fungi on their skin at any given time.5 Cytologic examination of a skin lesion or biopsy is essential to directing the clinician’s diagnostic plan. A confirmed diagnosis of a fungal disease can only be made when culture and histopathology confirm the specific pathology associated with the organism. Treatment should follow antifungal sensitivity testing. Misuse of antimicrobials may predispose snakes to fungal infections. Mycotic gastroenteritis has been reported in snakes maintained at suboptimal environmental conditions and administered inappropriate antibiotics.6

VIRAL Isolating viruses from snakes is a relatively recent event. Historically, most of the infectious diseases characterized in snakes were reported to have a bacterial origin. It is possible that some of these findings were the result of limited diagnostic capabilities. With the advent of new diagnostic methods, such as enzyme-linked immunosorbent assays and the polymerase chain reaction technique, more viral diseases can be expected to be identified in the future. Herpes virus or herpes-like viruses have been isolated from Indian cobras (Naja naja), Siamese cobras (N. naja kaouthia), and banded kraits (Bungarus fasciatus).7-9 The isolates from the Indian cobra and krait were incidental findings from the snake’s venom.7,8 The herpes-like virus isolated from the Siamese cobra was associated with pathologic changes in the venom gland. A herpes-like virus also has been isolated from juvenile boa constrictors (Boa constrictor constrictor).10 Intranuclear inclusions were found in the liver, pancreas, kidney, and adrenal cortex in two of the snakes. The limited number of herpes virus infections reported in snakes might suggest that the virus is rare in ophidians; however, a lack of confirmed diagnoses may also be associated with limited diagnostic test availability or a failure of veterinarians to pursue these cases. To date, there has been no specific treatment recommendation for ophidian herpes virus infections, although acyclovir may be used to suppress infections. Adenovirus has been isolated from a single adult boa constrictor, two rosy boas (Lichanura trivirgata), and a royal python (P. regius).11-13 The histopathologic findings in the snakes were similar to those lesions described for higher vertebrates, including hepatocellular necrosis and basophilic intranuclear inclusions. The route of transmission for this virus has not been described, although minimizing contact with any secretion from an infected snake is strongly recommended.

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152 Paramyxovirus (PMV) is one of the best-documented viral infections in snakes. The first report of this virus was from a collection of Fer-de-lance (Bothrops atrax) from Switzerland.14 Since that time, PMV has been isolated from both viperid and nonviperids. All snake species should be considered susceptible to PMV. The route of transmission for this virus is likely through contact with contaminated respiratory secretions. Affected snakes may have nasal discharge and purulent hemorrhagic discharge from the glottis. Neurologic disease has also been reported to occur with PMV.15 Diagnosis can be made using a hemagglutination inhibition assay or by viral isolation. Because the hemagglutination inhibition assay is a serologic assay, serial samples are required to confirm an active infection. In the USA, there are at least 3 laboratories performing this assay. Unfortunately, there is limited agreement between the results of these laboratories, so clinicians should use caution when interpreting the test results (M. Allender and M. Mitchell, unpublished data). Pulmonary hemorrhage and caseous debris are common postmortem findings.There is no current therapy for this virus. Trials evaluating a PMV vaccine have proven ineffective.16 Quarantine of new acquisitions is strongly recommended. The quarantine period should be at least 3 months. A thorough examination and a minimum of two negative assays should be performed before releasing the animal from quarantine. Inclusion body disease (IBD) can be a devastating disease in snake collections. The suspected cause of IBD is a retrovirus; however, this has yet to be confirmed. Although this disease is generally associated with boas and pythons, IBD has been reported in other species of snakes.17 In general, boa constrictors present with a history of regurgitation and mild central nervous system dysfunction, such as a postural change (stargazing), slowed righting reflex, and impaired locomotion (Figure 7-25). Pythons generally present for more severe disturbances of the central nervous system, including a loss of the righting

Figure 7-25 Boa constrictors with inclusion body disease often develop severe neurologic disease. This particular animal had no righting reflex.

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response, opisthotonos, and paralysis. As the disease progresses, snakes generally experience weight loss, dysecdysis, and secondary opportunistic infections, such as stomatitis, dermatitis, and pneumonia. The severe neurologic disease characterized in pythons generally limits their ability to acquire food; therefore regurgitation is rarely noted in pythons. The route of transmission for IBD has not been determined. However, clinical disease has been identified in snakes that have direct contact with infected individuals and with snake mites (Ophionyssus natricis). Diagnosis can be attempted antemortem from biopsies of the kidney, pancreas, esophagus, and liver. Eosinophilic intranuclear inclusions may be observed within the tissues when using a hematoxylin and eosin stain. Inclusion bodies may also be observed in red blood cells. However, only trained individuals should perform this procedure, as red blood cell nonviral inclusions are not uncommon findings in reptiles. Because of the low sensitivity of biopsy, false negative results are possible. One of the difficulties in characterizing IBD is that boas have been shown to harbor endogenous retroviruses. Because adenoviruses and reoviruses have also been isolated from snakes with neurologic disease, it is difficult to determine if these viruses also play a role in IBD. Postmortem examination should also include samples from the central nervous system. Encephalitis is a common pathologic finding. There is no effective treatment for IBD. Because this disease appears to be highly contagious, affected snakes should be culled and euthanized after antemortem confirmation. Preventing the introduction of O. natricis into a collection and quarantining new arrivals are strongly encouraged to reduce the likelihood of introducing IBD into an ophidian collection.

Parasitic Diseases Ticks are a common finding on wild-caught snakes. These parasites can be found anywhere on the snake but are frequently found around the eyes, oral cavity, and gular fold. Ticks can serve as vectors for disease and should be removed and disposed of properly. Special attention should be given to those tick species that are imported with reptiles from areas that harbor pathogens that could be potentially devastating to domestic livestock. The tick should be manually removed. It is important to grasp the tick at the mouthparts to prevent remnant material from causing a granuloma or abscess. The snake’s enclosure should also be thoroughly cleaned to remove larval stages of the ticks. Ophionyssus natricis is the common snake mite (Figure 7-26). This ectoparasite can be devastating in snake collections. Both larval and adult forms of the mite are parasitic. Snake mites can become reproductive within 2 to 2.5 weeks and live for 5 to 6 weeks. Chronic exposure can lead to the development of anemia. O. natricis also can serve as a vector for disease and can rapidly disseminate pathogens in a collection. Snake mites may play a role in the dissemination of inclusion body disease. A variety of miticides have been used to eliminate these parasites, including bathing the snake in water or applying mineral oil, topical and parenteral iver-

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A

B

Figure 7-26 Snake mites appear as red-brown “moving specks” on snakes (A). They can appear anywhere on the body. A mite infestation can be diagnosed by swabbing the red-brown moving specks with a mineral oil–coated cotton tipped applicator and reviewing the slide under a microscope (B).

mectin, organophosphates, topical permethrins, and topical pyrethrins. Treatment must be twofold, including both the snake and the environment. Provent-a-mite® (Pro Products, Mahopac, New York) has been found to be both effective and safe when the directions on the label are followed.18 Cryptosporidium serpentis is a significant protozoal parasite in snakes. Transmission of this parasite is via the fecal-oral route. Indirect exposure via inanimate objects can also occur, as these parasites have a thick-walled oocyst that is resistant to desiccation. Snakes with cryptosporidiosis may present as asymptomatic carriers or with severe gastroenteritis. The latter may present with a history of weight loss and regurgitation. On physical examination affected snakes may be dehydrated, have obvious muscle wasting, and a mid-body swelling. Because this parasite infects the mucus-secreting cells of the stomach, it can cause severe gastric hyperplasia. Acid-fast stains of feces or the regurgitated meal may be diagnostic. The parasite can also be identified on direct smears, but this is less common (Figure 7-27). Unfortunately, because this parasite can be transiently shed, multiple samples (3-7) may be required for diagnosis. Feeding a snake may increase the reproductive cycle of the parasite and can improve the likelihood of making a diagnosis. Survey and contrast radiographs may be used to differentiate a gastric foreign body from gastric hyperplasia. A commercially available immunofluorescent antibody assay (Merifluor, Meridian Biosciences, Inc., Cincinnati, Ohio), developed for use in humans to diagnose Giardia and Cryptosporidium parvum, may be used. This assay is more sensitive than an acid-fast fecal smear and is capable of characterizing fewer organisms than is an acid-fast fecal stain. To date, treatment of C. serpentis in snakes has been inconsistent. Antimicrobials have been found to reduce shedding, but they do not appear to eradicate the parasite. This organism does not appear to be zoonotic; however, strict sanitation and hygiene practices should be followed when working with infected animals. Infected animals should be culled to prevent the dissemination of the parasite into an ophidian collection.

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Figure 7-27 Cryptospordium serpentis oocyst (arrow) on a direct fecal smear.

Eimeria spp., Isospora spp., and Caryospora spp. are the most frequently encountered coccidian parasites in snakes. Coccidia have a direct life cycle. In higher vertebrates coccidial infections are generally self-limiting, but in snakes they generally are not. Coccidia infect primarily the intestinal and biliary epithelium, although infections of the renal epithelium also occur. Fibrosis and epithelial ulcerations are the most common histologic findings associated with coccidial infection. The clinical course of the disease and pathologic conditions is more severe in neonatal snakes. Diagnosis can be made by evaluating fecal material on a direct saline smear or fecal flotation. Treatment is generally accomplished using sulfonamides. The snake should be hydrated before it is administered sulfonamides. Most therapeutic plans recommend 5 to 7 days of treatment; however, this is generally insufficient, and treatment should continue until three negative fecal exams have been performed over a 15-day period.

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Entamoeba invadens is a protozoal parasite that can cause severe, protracted disease in snakes. The life cycle is direct; thus this parasite can spread rapidly through an ophidian collection. Transmission is via the fecal-oral route. Chelonians may serve as reservoirs for this parasite. Affected snakes may develop a hemorrhagic diarrhea, severe dehydration, and muscle wasting. Mortality can approach 100%. Ulcerative gastritis and colitis are common histopathologic findings. Diagnosis can be made on a direct smear with Lugol’s iodine or a fecal flotation. Metronidazole is the treatment of choice (25-50 mg/kg PO SID × 7-10 days). Supportive care, including fluid therapy, enteral support, and antibiotics for secondary bacterial infections, is also recommended. Ciliated and flagellated protozoa are frequent findings on a microscopic examination of snake feces. Many of these organisms are commensals and likely serve an important role in the maintenance of the microecology of the gastrointestinal tract. Treatment should be considered only if the snake is demonstrating clinical disease and has pathologic symptoms associated with the protozoa. Certain flagellates, including Giardia spp. and Trichomonas spp., can be pathogenic in snakes. Affected snakes may be dehydrated and have diarrhea. Diagnosis and treatment is similar to that described for E. invadens. Digenetic flukes are an infrequent finding in snakes. These flukes have an indirect life cycle, which generally includes a gastropod as the intermediate host. Styphylodoro, or renal flukes, generally parasitize the excretory system. Severe infestations can lead to significant renal pathology. Refifers, or lung flukes, may be found in the buccal cavity, pharynx, esophagus, or respiratory tract. The adult flukes generally cause mild ulcerations at their site of attachment. Because the life cycle of digenetic trematodes requires an intermediate host, the life cycle in captivity is self-limiting. Diagnosis can be made by performing a fecal or sputum flotation. Treatment can be accomplished by using praziquantel (5-8 mg/kg IM q10 days,

TABLE 7-2

2-3 treatments) and manually removing flukes that are found in the buccal cavity. Cestodes are not an uncommon finding in wild-caught snakes. The life cycle of the cestode also requires an intermediate host. Spirometra spp. and Bothridium spp. are pseudophyllideans that are found in snakes. The pleurocercoids of these cestodes are found usually in the subcutaneous (SC) tissues of the snake. On physical examination the pleurocercoids will appear as soft flocculent SC masses. Treatment can be accomplished by surgical excision of the parasite. Mesocestoides are another common cestode found in snakes. Because snakes serve as an intermediate host for this parasite, this cestode is generally found encysted in soft tissues. The snake generally serves as a definitive host for proteocephalidians. The adult cestodes are generally found in the small intestine. Identification of proglottids in fecal material or eggs from a fecal flotation is diagnostic. Treatment for those cestodes in which the reptile serves as a definitive host can be done using praziquantel (5-8 mg/kg IM q10 days, 2-3 treatments). Nematodes are the most frequently encountered endoparasites in snakes. The life cycle of nematodes may be direct or indirect. Clinical signs may vary with the density of parasites and the location of the parasites within the host. Affected snakes may be asymptomatic or have diarrhea, intestinal obstruction and volvulus, pneumonia, renal disease, or chronic muscle wasting. Diagnosis can be made by examining a direct fecal smear or fecal flotation. There are a variety of parasiticides that can be used to treat snakes (Table 7-2). Pentastomids are not an uncommon finding in wild-caught snakes. Affected animals may be asymptomatic or have generalized respiratory disease. Diagnosis can be made from examination of the feces (eggs) or sputum (eggs) and survey radiographs (adults). There are several recommendations for treatment, including ivermectin (0.2 mg/kg), fenbendazole (50-100 mg/kg), a combination therapy of ivermectin and fenbendazole, levamisole, and surgical/endoscopic removal of

Parasiticides Used to Treat Snakes

Parasiticide

Dose

Indication

Fenbendazole

Nematodes

Metronidazole

25-50 mg/kg PO 25 mg/kg SID 3 days, repeat 2 weeks if necessary 0.2 mg/kg IM, PO q 10 days, 2-4 times 5-10 mg/kg ICe24 once; repeat 2 weeks 25-50 mg/kg PO25

Permethrin

0.5%

Praziquantel

5-8 mg/kg PO, IM25 once; repeat 2 weeks 50 mg/kg PO, SID 7-21 days

Ivermectin Levamisole

Sulfa-dimethoxine

Ectoparasites, nematodes Lungworms Protozoa, anaerobic bacteria Ectoparasites

Comment

Do not use in indigo snakes or in debilitated snakes Narrow range of safety; do not use in debilitated snakes 2300-325 mOsm/L). Based on this, the majority of the isotonic fluids (e.g., 0.9% saline: 308 mOsmol/L; Normasol: 294 mOsmol/L) used to replenish fluid in mammals would be considered hypertonic or hypotonic depending on the individual snake’s status. Calculating a snake’s osmolarity using an osmometer is strongly recommended; however, when this can’t be done, estimating the osmolarity using the following formula is recommended: 2(Sodium + potassium). Assessing dehydration in snakes is more difficult than in mammals. Snake skin is not as naturally elastic as mammalian skin. To best assess the elasticity (or natural lack thereof) of snake skin, it is important to routinely examine clinically hydrated animals to develop an understanding of “normal” (Figures 7-29). The lateral body wall is the best area to assess skin elasticity in a snake. The mucous membranes of a snake should be moist. Dry, ropy mucus in the oral cavity is generally an indication of dehydration. Snake mucous membranes are typically pale pink in color, and the capillary refill time should be less than 2 seconds. An increased capillary refill time is suggestive of dehydration. The positioning of the eyes can also be

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used to assess snake hydration. Snakes that are severely dehydrated (>8%-10%) develop sunken eyes as a result of a loss of moisture in the retro-orbital fat pads. Blood work is often the best method for assessing the hydration status of a snake, and it is for this reason that veterinarians should recommend obtaining and testing a sample when an animal is clinically normal for comparison. A packed cell volume, sodium, chloride, total protein, and plasma osmolarity can be used to assess hydration and measure the response to treatment. There are several routes for replenishing fluids in snakes, including per os (PO), SC, ICe, and IV. For animals that are mildly dehydrated (5%) can be given fluids via the coelomic cavity or intravenously. ICe fluids should be given in the caudal third of the body. The snake should be placed in dorsal recumbency and the needle inserted between the ventral scales (Figure 7-31). Positioning the animal using this method will minimize the likelihood of introducing the fluids into the lung(s). IV catheters can be placed into a jugular vein. Jugular catheter placement generally requires a surgical cut-down. The heart is used as the primary landmark for locating the jugular veins. Place the snake in dorsal recumbency. Identify the heart. Once the heart is identified, count 9 to 10

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Figure 7-32 An IV catheter can be placed into the jugular vein of a snake to ensure rapid delivery of fluids and other therapeutics. The procedure requires a cut-down.

ventral scales cranially. The surgical site should be disinfected using standard techniques. A 0.5- to 1.0-cm incision should be made between the first and second rows of the dorsal scales. Blunt dissection of the medial ridge of the ribs will reveal the jugular vein. An appropriately sized catheter should be inserted into the vein, an injection port attached, and the incision closed (Figure 7-32). Fluids can be administered via a bolus or continuous rate infusion. Maintenance fluid rates for reptiles are much lower (1030 ml/kg/day) than those recommended for birds (100 ml/kg/ day) or mammals (80-100 ml/kg/day). When developing a fluid replenishment plan, it is important to consider both the deficit and maintenance requirements of the snake. Animals should be reassessed daily to ensure that the fluid deficit is being replaced and that the animal is not being overhydrated. Serial blood testing (e.g., packed cell volume, electrolytes, total protein, and osmolarity) can be used to assess the patient’s response to therapy.

Antimicrobial Drugs Antimicrobial drugs are likely one of the most common therapeutics administered to snakes. The dogma that all diseases are bacterial infections until proven otherwise is likely the culprit. Of course, antibiotics and antifungals do serve important purposes for captive reptiles. When deciding whether to include an antimicrobial into a therapeutic plan for a snake, it is important to consider the pharmacology of the compound. I consider a series of questions when selecting an antimicrobial. 1. Is the drug bactericidal or bacteriostatic? Determining whether a drug is cidal or static is important because most of the bacterial pathogens isolated from snakes are Gram negative. A rapid kill of Gram-negative bacteria could lead to the release of endotoxins, resulting in the death of the host.

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TABLE 7-3

Antimicrobial Drugs Used to Treat Snakes

Drug

Dose

Comments

Amikacin

3 mg/kg IM q72h

Not to be used in dehydrated snakes or those with renal disease

Carbenicillin Ceftazidime Chlorhexidine Ciprofloxacin Clotrimazole Doxycycline Enrofloxacin Itraconazole Ketaconazole Silver sulfadiazine Trimethoprim-sulfadimethoxine

400 mg/kg IM q24h26 20 mg/kg IM q72h27 Topical 0.05-0.1% 10 mg/kg PO q24-48h Topical 10 mg/kg PO q24h 5-10 mg/kg IM once, PO 5-10 mg/kg PO SID 25 mg/kg PO SID24 Topical 20 mg/kg PO, IM q24h

Apply to external topical wounds Topical application fungal dermatitis

Topical application fungal/bacterial dermatitis Not to be used in dehydrated snakes or those with renal disease

IM, intramuscular; PO, per os; q, every; SID, once a day.

2. What is the spectrum of the antibiotic? Does it cover Gram-positive bacteria? Gram-negative? Both? Aerobes versus anaerobes? 3. Will the drug get to the desired tissues? If an infection is associated with the respiratory tract, and a selected drug does not penetrate the respiratory tissues, then it is of no value. 4. What is the mechanism of action of the antibiotic? Knowledge of how an antibiotic works is important when considering methods of resistance. Certain resistance factors occur as a result of mutations while others are plasmid generated. 5. How is the drug metabolized and excreted? If a drug must be metabolized by the liver to become active, and the snake has liver disease, the likelihood that the drug will perform as desired is decreased. 6. What are the side effects? Being able to identify the side effects associated with a drug are important, so that the compound can be discontinued before it causes irreparable harm. 7. How often does the drug have to be administered? Corticosterone is a natural hormone produced by snakes in response to stress. Elevated levels of corticosterone can lead to immune system suppression and altered metabolism. By minimizing handling time (e.g., number of treatments), veterinarians can reduce the likelihood of stimulating excessive corticosterone secretion in their snake patients. By answering these questions veterinarians will be sure to select an antibiotic based on desired effect rather than a “best guess” approach. Table 7-3 lists antimicrobial drugs commonly used to treat snakes.

Antiparasitic Drugs Antiparasitic drugs are commonly used to treat snakes. Many veterinarians and herpetoculturists treat animals regardless of a confirmed diagnosis. This is often done because the parasites

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Emergency Drugs Used to Treat Snakes

Drug

Dose

Indication

Atropine

0.5 mg/kg IV, IT, IO, IM

Diazepam Doxapram

0.5-1.0 mg/kg IM, IV 20 mg/kg IV, IO, IM

Epinephrine

0.5-1.0 mg/kg IV, (1 : 1000) IO, IT

Bradycardia, CPR, reduced secretions Seizures CPR, respiratory stimulant Asystole, CPR

CPR, cardiopulmonary resuscitation; IM, intramuscular; IT, intratracheal; IV, intravascular.

may be shed periodically, and are thus difficult to identify, or because of a perceived problem. There are potential disadvantages to this treatment method. Regular treatment could lead to the development of resistance factors that limit the effectiveness of a drug. Antiparasitics are not without their negative side effects. Certain species of snakes may be more sensitive to certain anthelminthics. Routine treatment can lead to toxic side effects (e.g., hepatopathy, anaphylaxis, or death). I prefer to treat animals based on a confirmed diagnosis. The administration of fenbendazole to a snake with cestodes is a waste of money and may provide a false sense of security for the owner. (See Table 7-2 for a list of antiparasitics used to treat snakes.)

Emergency Drugs Veterinarians working with snakes may find themselves in situations where they need to provide emergency drugs. Cardiac depression (bradycardia) and/or cardiorespiratory failure during an anesthetic event are the most common problems that I encounter. Table 7-4 provides a list of emergency drugs used for snakes.

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Miscellaneous Drugs In general, drugs used to manage disease processes in nonreptilian vertebrates can be used for snakes. The vast majority of the therapeutic dosages recommended for snakes are empirical or based on mammalian or avian recommendations. Veterinarians should inform their clients that the compounds used to treat their pets are being used off-label.

Nutritional Support Nutritional support should be considered for any patient with a history of anorexia. However, defining anorexia in a snake may be easier said than done. It is not uncommon for some snakes, especially larger bodied pythons or boas, to not eat for extended periods in captivity. This may be associated with a number of factors, including that snakes are often overfed in captivity, not provided much exercise, or maintained at temperatures that lower their metabolic rate. To determine whether a period of inappetence is associated with a medical problem or is the result of a normal behavioral pattern for the snake, it would be prudent for the veterinarian to examine the body condition of the animal and perform a plasma biochemistry assay. The epaxial muscles along the spine are an excellent location for assessing body condition. A snake should have well-defined epaxial muscles. A prominent spinal column is suggestive of muscle wasting. If the spinal column cannot be easily seen or palpated, the snake should be considered obese. Ultrasound can be used on the coelomic cavity to detect the presence or absence of the linear fat bodies. Snakes that are truly anorectic may have decreased electrolyte and glucose levels. Snakes that are judged to be in a negative energy balance should be provided supplemental calories. There are several methods that can be used for calculating the caloric requirements of a snake. Calorie replacement for snakes can be done using whole prey or a liquid enteral. Whole prey species can be used when the anorexia is short term and the gastrointestinal system is functioning normally. Mammalian and/or avian prey can be skinned before being offered to the snake to reduce the digestive load and expedite caloric uptake. Lubricating the prey with a small quantity of petroleum jelly or vegetable oil may expedite the passage of the meal. Take care not to introduce the lubricant into the glottis. Pinkie pumps are designed to macerate juvenile rodent prey items and may be used to simplify the force-feeding process. Animals with a history of long-term anorexia may benefit from a liquid enteral. Fluker Farms Carnivore Repta-Aid (Fluker Farms, Port Allen, Louisiana) is a powdered diet that can be mixed with water to provide essential replacement calories (Figure 7-33).

SURGERY Preoperative and Postoperative Considerations Surgical procedures in a snake should follow the same basic practices used for domestic mammals. A thorough physical

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Figure 7-33 Fluker Farms Repta-Aid (Fluker Farms, Port Allen, Louisiana) can be used to replace essential calories in a snake. The product is mixed with water and can be delivered via a feeding tube.

examination and baseline data—including a packed cell volume, total solids, complete blood count, plasma biochemistries, and clotting profile—should be performed to assess the anesthetic risk and determine the suitability of the patient for the surgery. In cases where this information cannot be obtained, the client should be made aware of the added risk involved, given the lack of knowledge regarding the animal’s physiologic condition. The surgical suite should be prepared before the patient is anesthetized. A heat source should be used to maintain the animal’s body temperature during the procedure. The veterinarian should attempt to maintain the animal at a temperature consistent with its optimal environmental temperature range. Convective forced air heating systems, such as the Bair hugger (Arizant Healthcare, Eden Prairie, Minnesota), recirculating water blankets, electric heating pads, and incandescent lights can be used to provide heat. It is important to monitor the animal’s temperature during the procedure using an esophageal or colonic thermometer. By monitoring the animal’s temperature during the procedure, it will be possible to alter the heating pattern during the procedure to maximize the efficiency of the animal’s metabolism and thus the animal’s ability to process the anesthetics. The surgical site should be disinfected with 0.5% to 1.0% chlorhexidine or povidone-iodine and sterile 0.9% saline. I recommend warming the fluids for this procedure to minimize the loss of heat that may occur. Alcohol should not be used as a disinfectant, as it can also lead to the loss of body heat.

Anesthesia Historically, clinical and surgical procedures performed on snakes were done using manual restraint or hypothermia. Fortunately, new anesthetics have become available that produce safe and effective anesthesia in snakes. Dissociative anesthetics,

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propofol, local anesthetics, opioids, and inhalant anesthetics are the most frequently used anesthetics in snakes. Most of the knowledge veterinarians have of these agents is based on mammalian, avian, and nonophidian reptiles, as few anesthetic research studies have been performed using snakes. When developing an anesthetic plan for an ophidian patient, veterinarians should become familiar with the physiologic needs of a particular species. The dissociative agents, including ketamine and tiletamine, are frequently used to anesthetize snakes for clinical procedures. Although the dissociative agents are characterized as anesthetics, they do not provide sufficient visceral analgesia in mammalian species and should not be used exclusively for invasive surgical procedures in snakes. The dissociative agents are readily available, are inexpensive, and can be administered intramuscularly. One of the primary disadvantages of using dissociative anesthetics is a prolonged recovery period. Even at low doses (5 mg/kg tiletamine), snakes can require 24 to 48 hours to fully recover. Doses of ketamine as high as 80 mg/kg have been reported for the provision of surgical anesthesia; however, the dissociative anesthetics neither result in effective muscle relaxation nor provide visceral analgesia and should not be used as stand-alone anesthetics for invasive procedures. The addition of an alpha-2 anesthetic may be appropriate. However, little research has documented the effects of these drugs on snakes. I have found 10 to 20 mg/kg ketamine or 3 to 6 mg/kg Telazol (tiletamine and zolazepam) (Ft. Dodge Laboratories, Ft. Dodge, IA) sufficient to manage fractious snakes and provide anesthesia for minor surgical procedures. Propofol (AstraZeneca, Wilmington, Delaware) is a nonbarbiturate anesthetic that can be used as a sole parenteral agent to provide surgical anesthesia. The effects of propofol are not cumulative, and thus rapid recovery (30-45 minutes) occurs in most cases. To obtain a desired effect, propofol should be administered intravenously. Propofol can also be administered in the jugular vein or heart. Because of the potential cardiac and respiratory depressant effects of propofol, the snake should be intubated and ventilated during the anesthetic procedure. Propofol at 5 to 10 mg/kg administered in the ventral coccygeal vein by slow injection has been used repeatedly to anesthetize venomous snakes. In general, 5 to 15 mg/kg IV will provide general anesthesia for 30 to 45 minutes in most snake species. When longer anesthetic periods are required, additional dosing can be given to effect. The inhalant anesthetics (e.g., isoflurane and sevoflurane) can be used to provide controlled anesthesia. The primary benefit of the inhalant anesthetics over the injectable compounds is that the veterinary surgeon has more direct control over the anesthetic. The snake should be intubated to ensure proper delivery of the anesthetic. The glottis is located on the floor of the mouth, caudal to the tongue. A noncuffed endotracheal tube should be used for the procedure. IV catheters can be used to intubate small snakes. Closely monitor the endotracheal catheter to ensure that it does not become plugged with mucus. Because snakes lack a diaphragm and can become apneic for extended periods of time, I recommend positivepressure ventilating the snake 4 to 6 times a minute. Recovery

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should be done in a warmed, dark environment to reduce unnecessary external stimuli. Local anesthetics (e.g., lidocaine, prilocaine) are often underutilized in reptile medicine. These anesthetics can be used to provide complete, local anesthesia for an animal. I generally use these compounds in combination with other anesthetics. For example, lidocaine may be used to provide anesthesia along an incision or at the base of a hemipenis for an amputation. Although there have been no studies done to determine the toxic dose for these compounds in snakes, I have found that dosing at less than 5 mg/kg is safe. Analgesics should be used whenever an animal may feel pain. Unfortunately, knowledge of pain perception in these animals is limited. Most veterinarians accept the notion that snakes respond to noxious stimuli by moving away from a negative stimulus (e.g., injection of a drug); it is unknown, however, whether the animal feels pain during such a response. Because nothing is known about pain perception in reptiles, I err on the protective side and administer analgesics even though it is also unknown whether the analgesics have any effect. Analgesics I use in my practice are butorphanol (0.51.0 mg/kg IM), carprofen (2-4 mg/kg IM, PO), and meloxicam (0.5 mg/kg PO).

Surgical Procedures The coeliotomy is routinely performed to pursue the biopsy of an abnormal mass, remove a foreign body (gastrotomy and enterotomy), or correct a dystocia. Because snake locomotion involves the direct contact of the ventral scales with a surface, surgical approaches should not be made along the ventrum through the large scales. An incision in that area may lead to excessive tension at the incision site and possible dehiscence. Approaches to the coelomic cavity of a snake should be made between the second and third rows of lateral scales to prevent contamination of the incision by the substrate. Closure of the coeliotomy should be made in two layers: the body wall and skin. The skin is considered the holding layer. An everting horizontal mattress pattern should be used to close the skin incision. An absorbable synthetic should be used to close the body wall, and nylon suture can be used to close the skin. Sutures should be removed approximately 4 to 6 weeks after surgery. Snakes occasionally ingest foreign bodies from their environments. Affected animals may become anorectic or may regurgitate. The general location of the foreign body often can be isolated by digital palpation. Survey and contrast radiographs can be done to confirm location. Although the approach to a gastrotomy or enterotomy in a snake is similar to that used in mammals, special care should be taken when manipulating the thin walled intestine of the snake. While a twolayer closure may be used to close the bowel in mammals, this is often not possible in small snakes. Once the bowel is closed, the coelomic cavity should be irrigated with warm sterile saline. Broad-spectrum antimicrobials are indicated if the coelomic cavity becomes contaminated during the procedure.

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sutures should remain for approximately 72 hours, and the animal should be monitored closely to ensure that it can evacuate feces and urine. If the hemipenis is necrotic, it must be amputated. The hemipenes can be removed at the base of the organ using circumferential and transfixation sutures. The snake can remain in a breeding program as long as it retains one functional hemipenis.

ZOONOSES

Figure 7-34 When removing eggs from the oviduct in a snake, it is important to make the incision in an avascular region. Stay sutures should be placed into the oviduct to allow the surgeon the ability to manipulate the incision and direct the eggs through the incision. The stay sutures also prevent the surgeon from losing the oviduct into the coelomic cavity.

Dystocias in snakes are generally postfollicular. Approach to the dystocia in snakes should follow standard surgical protocol. The value of the snake and the embryos or eggs should be considered before the surgery. If the snake is used exclusively for reproductive purposes, then a hysterectomy is not an option. When performing a cesarean section in a snake, the incision should be made into a relatively avascular region of the oviduct (Figure 7-34). Once the embryos or eggs are removed, the incision should be closed using a two-layer closure. Subspectacular abscesses can develop in snakes with obstructed nasolacrimal ducts, retained spectacles, and penetrating wounds. To facilitate draining the abscess, an incision into the spectacle is needed. A ventral triangular spectaculotomy is the preferred method of treatment. By removing a ventral section of the spectacle, the veterinarian can drain the abscess, irrigate the area, unblock the nasolacrimal duct, and treat the corneal surface. Medical treatment, including topical antibiotics and artificial tears, is required until the spectacle heals. During excitement or stress, male snakes may evaginate a hemipenis and cause damage to the structure. A damaged hemipenis should first be evaluated for tissue viability. If the tissue is viable, then it should be lavaged with copious quantities of warm sterile 0.9% saline and a mild disinfectant (0.5% chlorhexidine). Once cleaned, the structure should be replaced. If the hemipenis is edematous and cannot be replaced, a hypertonic solution (50% dextrose) may be applied topically to reduce the edema. Copious lavage with sterile saline should be done after the swelling has been reduced to remove any of the residual irritating dextrose. Two simple interrupted sutures may be placed through the anal plate and the scale caudal to the vent to prevent the hemipenis from reprolapsing. The

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Snakes can serve as reservoirs for a number of potentially zoonotic diseases. Salmonella spp. is routinely isolated from captive snakes. The prevalence of this bacterium is high in captive snakes because of a relative lack of quarantine and minimal sanitation. Young children (15,000 cells/ml) in rabbits; however, several changes may be associated with an acute infection, including a low total number of white blood cells (3-4 years of age) rabbits.4 These cases should be managed using the diagnostic plan described for uterine adenocarcinomas. Removing the affected mammary gland (local or radical) and the reproductive tract provides the best prognosis. Metastasis can be found in regional lymph nodes, lungs, or other organs. The incidence of mammary adenocarcinoma can be reduced significantly by performing ovariohysterectomies in rabbits before they are sexually mature. Some clinicians use different chemotherapeutics as adjunctive therapy to surgical treatment. Tamoxifen is a nonsteroidal, antiestrogen drug with antitumor effects that works by binding estrogen receptors.28 The overall value of this drug in rabbits is unknown for

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392 spayed animals. However, clinicians using the protocol believe, subjectively, that survival is extended. If tamoxifen is used, the patient should be routinely monitored for changes in behavior, tumor size, and liver or CBC changes. Although adenocarcinoma is the most common reported reproductive neoplastic condition in captive rabbits, uterine leiomyoma and leiomyosarcoma, vaginal squamous cell carcinoma, ovarian hemangioma, testicular seminoma, interstitial cell carcinoma, testicular adenocarcinoma, and teratoma have also been reported. Lymphosarcoma, as in other mammalian species, is a commonly diagnosed neoplasia in rabbits. Urinary neoplasia rarely occurs; however, nephroma, lymphosarcoma, renal carcinoma, and urinary bladder leiomyoma have been reported. Primary neoplasia of the lungs is very rare. Carcinomas have been found to be associated with the nasal turbinates, causing an unresponsive nasal discharge. Thymomas have been reported in both juvenile and adult rabbits. Affected animals often present with tachypnea, dyspnea, and, occasionally, bilateral exophthalmos resulting from the interference of the blood flow to the heart. A thymomectomy can be performed via a median sternotomy.4 Metastasis to the lungs is a common sequela for many different malignant neoplasias. Both teratoma and neurinoma affect the central nervous system of rabbits. Neoplastic conditions found to affect the digestive tract include adenocarcinoma, leiomyosarcoma, papilloma, and metastatic uterine adenocarcinoma. Surgical resection is the treatment of choice for those conditions. Bile duct carcinoma has been reported in rabbits; however, in these cases, surgery is not an option and metastatic disease carries a grave prognosis. Cutaneous neoplasias that have been reported in rabbits include the following: cutaneous lymphosarcoma, nonviral papilloma, basal cell carcinoma, squamous cell carcinoma, and sebaceous gland carcinoma. Malignant melanoma has been associated with ocular, abdominal, thoracic, and vertebral disease.31

Miscellaneous Diseases DERMATOLOGY Many different diseases or conditions, such as obesity, sore hocks, arthritis, poor diet, dental disease, urinary disease, and neurologic disease, can lead to secondary dermatitis. Moist dermatitis around the oral cavity is a common sequela to dental disease. Discomfort caused by tooth malocclusion can lead to drooling, and the chronic wetness associated with the drooling predisposes the animal to bacterial dermatitis. Skin fold pyoderma is a common problem in obese animals; the sites most commonly affected are the dewlap, perianal, and genital areas. Treatment for those conditions requires aggressive wound debridement. Fur from the affected area should be clipped to provide adequate exposure. The wound should be cleaned with a topical disinfectant (e.g., Betadine or chlorhexidine). To control bacterial infections, we recommend using a hyperosmotic solution (e.g., 50% dextrose). The wound should be liberally irrigated with the 50% dextrose solution. After 1 to 2 minutes, the wound should be irrigated with isotonic saline

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to remove the dextrose. Extended contact with the dextrose can dry out the healing tissue. The procedure should be repeated at least three times, twice daily. In addition to this topical treatment, obese animals should be placed on a calorierestrictive diet. This is best achieved by reducing the amount of pellets and providing (ad libitum) grass hay (e.g., timothy hay). Surgery may be required to remove excess skin folds. In cases of tooth malocclusion, the primary problem should be identified and corrected. Dermal abscesses are a common presentation in rabbits and can occur anywhere on the body. Abscesses in rabbits are generally more caseous in nature than in other mammals. The thick, caseous nature of rabbit abscesses limits their drainage, and the use of standard Penrose drains is usually futile. Treating abscesses with (only) systemic antibiotics is ineffective, as the nucleus of the abscess in unaffected by the treatment. To effectively treat rabbit abscesses, the entire abscess (including the capsule) must be removed. In cases where the abscess and capsule are removed, the prognosis is good; however, in most cases the capsule is inherently attached to vital organs or bone and is difficult to remove. When it is impossible to extract the entire capsule, as much of the capsule should be removed as possible to avoid contact between caseous material and unaffected tissue. Culture and antibiotic sensitivity testing of the purulent material or capsule is recommended. Abscesses that cannot be completely removed should be left open for daily cleansing and debridement. When choosing a topical antibiotic therapy, be sure to select antibiotics that will not induce bacterial dysbiosis when the rabbit grooms itself. Penicillin (80,000 IU/kg) or ampicillin (20 mg/kg), although not to be given orally because of the potential for dysbiosis, can be used parenterally to treat local skin abscesses. In our experience, the application of honey to the abscess as a hyperosmotic has produced moderate success. The honey, in addition to acting as an antimicrobial, encourages a rabbit to groom or lick at the area and promotes drainage.2 The honey also acidifies the area and promotes healing. Systemic oral antimicrobials and analgesics should also be considered when treating rabbit abscesses.29 Antibiotic-impregnated polymethylmethacrylate (PMMA) beads can also be used to manage rabbit abscesses. Heat-stable antibiotics must be used to make PMMA beads. Amikacin 1.25 g/20 g methylmethacrylate, cefazolin 2 g/20 g methylmethacrylate, cephalothin 2 g/20 g methylmethacrylate, gentamicin 1 g/20 g methylmethacrylate, or tobramycin 1 g/20 g methylmethacrylate are all appropriate for PMMA beads.30 Treating abscesses in rabbits can be frustrating and may require multiple procedures. In some cases, treatment is not successful, and amputation may be the only option. Owners should always be made aware of their pet’s prognosis before treatment is initiated. Systemic antibiotics are appropriate for generalized abscesses. Antimicrobial treatment is ideally based on culture and antibiotic sensitivity testing. While testing is pending, abscesses can be treated with benzathine/penicillin G procaine (40,000 IU/kg SC SID × 14 days, then q48h × 2 weeks); enrofloxacin (5 mg/kg PO BID); or metronidazole (30 mg/kg PO BID).30,35 The use of trime-

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thoprim sulfa-dimethoxine should be avoided when there is caseous material present because the antibiotic is inactivated by pus. “Sore hocks,” or ulcerative pododermatitis, is caused by an avascular necrosis of the plantar surface of the rear feet. This is a painful, progressive disease that can be difficult to treat. Predisposing factors for this disease include breed (e.g., Rex), housing (e.g., metal grate flooring, concrete, carpet), lack of exercise, and obesity. Rabbits housed on an inappropriate flooring surface alter their weight distribution and bear the brunt of their weight on their metatarsus and hock instead of on the claws and plantar surface of the feet.2 This shift in weight leads to the medical displacement of the superficial flexor tendon, which can exacerbate the condition.2 Treatment is aimed at relieving the pressure on the affected area. A nonabrasive, dry surface should be provided. Providing an affected animal time during the day on a grass lawn or deep bed of hay is helpful. A thick foam rubber pad, towels, or newspaper substrate can also be used to minimize the likelihood of exacerbating this disease. Skin wounds can be managed using the technique described previously. Surgical skin glue can be used to protect the wounds. A padded splint can be used to alleviate any pressure on the lesions. Opportunistic infection should be managed with appropriate antibiotics. Analgesics (e.g., nonsteroidal antiinflammatories, opioids) should be used to alleviate discomfort. Underlying problems, such as obesity and inappropriate substrate, should be corrected. Surgery is usually not performed because there is no skin for closure. Hypersensitivity reactions can occur immediately or days after an injection is given. Affected animals present with erythema and swelling around the face and extremities. Most animals heal without complication or with a need for topical therapy. Giving injections subcutaneously may reduce the occurrence of these reactions.2 In severe cases, such as may occur with intramuscular injections, self-mutilation may occur.21 Severe reactions can be managed with steroidal (prednisone, 0.5-1.0 mg/kg) or nonsteroidal (meloxicam, 0.3 mg/ kg) antiinflammatories. Alopecia is a common problem in captive rabbits and may result from self-mutilation, barbering, parasites, endocrine disease, or infectious causes. Self-mutilation is most common in rabbits after a traumatic experience or surgery. Barbering is the result of aggression between cagemates. If barbering is suspected, animals should be separated. Diagnosis of parasitic, infectious, or endocrine disease requires a complete diagnostic work-up. Treatment is dependent on the specific diagnosis. Rectal papillomas can present as cauliflower-like masses extending from the rectum. Most of these masses do not cause a problem; however, occasionally these masses can cause frank hemorrhage or mild diarrhea. Surgical resection of the mass is required for treatment. Rare dermatologic disorders of rabbits include sebaceous adenitis, nonpruritic scaling and alopecia, Ehlers-Danlos syndrome, and neoplasia (see Neoplasia). Infectious and parasitic diseases are frequently associated with dermatologic disease in rabbits (see Infectious Disease; Parasitic Disease).

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DISEASES OF THE SENSE ORGANS Ears represent a significant proportion of the surface area of a rabbit. Otitis externa is a common problem, and it is most often associated with a parasitic or bacterial predisposition. Parasitic infections were described previously (see Parasitic Disease).32 Lop breeds are predisposed to developing bacterial otitis because of the flexion of the cartilage at the base of their ears. Rabbits with otitis externa may present for shaking, pruritus, erythematous pinna, malodor, and possible auricular discharge. Because otitis externa can be painful, care should be taken when manipulating the ears; we prefer to treat affected rabbits with systemic antibiotics and antiinflammatories rather than treating the ears topically. Bacterial cultures should be collected before treatment. Otitis externa can advance to otitis media or otitis interna. Otitis interna should be ruled out in rabbits with a head tilt. Radiographs and/or CT scans can be used to assess the tympanic bullae. In severe otitis cases, a permanent osteotomy may be required.32 The most common causes of conjunctivitis in rabbits are bacterial infections (e.g., Pasteurella), trauma, and entropion. When evaluating rabbit eyes, the approach to the examination should be similar to that for other mammals. Unless trauma has been acknowledged in the history, a thorough work-up should be pursued, including a cytologic scraping and bacterial culture. For confirmed bacterial infections, topical antibiotics can be used. Treatment regimens may require three to four doses daily. Application of topical antibiotics three to four times a day in addition to removing the initial cause of conjunctivitis is recommended. Entropion is a common presentation in large breed and obese rabbits, and surgical resection is necessary for resolution.21 Epiphora can be associated with early signs of dental disease, causing the blockage of the nasolacrimal ducts, or it can occur secondarily to conjunctivitis or rhinitis. Any rabbit presenting with dacryocystitis, conjunctivitis, or epiphora should receive a thorough ophthalmic examination, in addition to skull radiographs, to evaluate the roots of the teeth. Because many cases of epiphora are the direct result of blocked or inflamed lacrimal ducts, the lacrimal punctum should be flushed with sterile saline to facilitate the removal of debris (Figure 14-17). The ducts can be flushed with antibiotics too if a bacterial infection is suspected. Anterior uveitis can occur as a result of trauma, bacterial disease, or E. cuniculi–induced lens disease. Corneal ulcers are also somewhat common in rabbits housed with other animals. A fluorescein stain should be used to confirm the presence of an ulcer. Retrobulbar abscesses can be problematic; diagnosis and treatment should be based on culture and antibiotic sensitivity testing and aggressive antibiotic therapy instituted. In many cases, enucleation is required. Because of the large retroorbital venous sinus, enucleation can be difficult.

NEUROLOGIC DISEASES Vertebral luxation or fracture is a common finding in rabbits with a history of hind end paresis or paralysis. Rabbits have a highly developed muscular system and a relatively delicate

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vertebral column, making them more prone to fractures. The most common cause of these injuries is associated with inappropriate restraint (Figure 14-18). A diagnosis can be confirmed from a history, physical examination, and vertebral radiographs. The prognosis for these cases depends upon the extent of the lesions. With vertebral luxation, medical management may be attempted; however, it is usually not successful. Prednisolone sodium succinate (0.5 mg/kg IV) can be used to control edema in the spinal canal. In addition to the steroid treatment, cage rest and the manual expression of the bladder can be done.21 Surgery can be attempted, but only by experienced surgeons.

Rabbits with torticollis, difficulty righting, or abnormal locomotion (e.g., flipping over) generally have a central nervous system disorder. The three most common causes of head tilt in rabbits are E. cuniculi infection, trauma, and P. multocida infection. Although less common, listeriosis and aberrant larval migrans can also cause torticollis. Diagnosis is generally based on a thorough history, physical examination, radiographs, CBC, serology, and CT scans. Treatment can be difficult and prolonged. Enrofloxacin can be used to manage P. multocida and fenbendazole/albendazole to manage E. cuniculi infections.21

DISEASES OF THE MUSCULOSKELETAL SYSTEM Splay leg or hip dysplasia is an apparent congenital condition seen in young rabbits. Affected rabbits present with an inability to abduct their legs, which prevents ambulation. Amputation is recommended if only one leg is affected.14 Although this condition is thought to be genetic, environmental factors may also have some effect. In a recent study, rabbits housed on slippery surfaces had an increased incidence of splay leg compared with those housed with appropriate traction.33 In severe cases, humane euthanasia is warranted.

DISEASES OF THE REPRODUCTIVE SYSTEM

Figure 14-17 Flushing the nasolacrimal duct.

Pyometra occasionally occurs in rabbits and is usually reported shortly after parturition. Affected rabbits may be asymptomatic or may show signs of anorexia, weight loss, vaginal discharge, hematuria, or infertility. Diagnosis is made based on history, physical exam, CBC, radiology, and ultrasound. Inflammatory leukograms are common. Survey radiographs

Figure 14-18 Vertebral injuries are common in rabbits that are not restrained properly. (Courtesy Dr. David Guzman.)

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and ultrasound often reveal a fluid-filled soft tissue structure in the caudoventral abdomen. Affected animals should be provided supportive care (e.g., fluids, calories) to correct any deficiencies. Systemic antibiotics are warranted. Ovariohysterectomy is the treatment of choice. Orchitis is relatively uncommon in captive rabbits because many animals are neutered at a young age. The disease is spread venereally, and can be caused by a bacterial infection. Clinical signs generally include enlarged testicles or epididymis and discomfort. Orchiectomy can be done in pet animals. Antibiotics may be used to treat breeding rabbits. The animal should be reevaluated before it is placed back into the breeding rotation. Pregnancy toxemia is a problem that occurs in does at the end of gestation during nesting; these animals, especially obese Dutch, Polish, and English breeds, fast or have reduced caloric intake.14 Clinical signs include weakness, depression, incoordination, coma, convulsions, or death. Fasting leads to ketoacidosis and shock. Treatment should be aimed at reversing ketoacidosis and providing both shock therapy and supportive care. Supportive care may include the provision of calcium gluconate, fluids, and nutritional support. Treatment is usually unsuccessful, so prevention is the key. Clients should attempt to reduce obesity in breeding does and monitor their caloric intake carefully during pregnancy. Cystic uterine hyperplasia is a problem in aged, intact does. Clinical signs include hematuria, anemia, reduced activity, a firm irregular uterus, cystic mammary glands, and cystic ovaries. Diagnosis is via clinical signs, radiology, and ultrasound. Ovariohysterectomy should be recommended to treat this condition.4 Pseudopregnancy can occur in does and lasts, on average, 16 to 18 days. Toward the end of the pseudopregnancy, does will go through normal nesting behavior (e.g., pulling out hair to line nests) and will have mammary development. Pseudopregnancy can resolve spontaneously or lead to pyometra or hydrometra.4 Ovariohysterectomy is the treatment of choice. Dystocia and retained fetuses occur commonly in obese does, in cases where the fetus is too large for the doe’s pelvic canal, or in does with uterine inertia. Clinical signs include nonproductive contractions and bloody to greenish-brown vaginal discharge. In nonobstructive dystocia, calcium gluconate and oxytocin may aid in the delivery of fetuses. In obstructive dystocia, a cesarean section is required. For prolonged dystocia events, the prognosis is guarded for both the doe and fetus. Lactating or pseudopregnant does are prone to developing mastitis. Clinical signs in affected animals are similar to those observed in other animals and include pyrexia, swelling, blue coloration of teats, inappetence, and depression. Bacteria, such as Staphylococcus spp. and Streptococcus spp., are usually the causative agent. Without treatment, mastitis can progress to septicemia and death. Treatment includes surgical drainage or mastectomy, warm compresses, and appropriate antibiotics. Penicillin (40,000 U/kg IM BID × 5 days) has been used with success.14 Other reproductive problems that occur in rabbits are abortion-reabsorption, decreased fertility, prolapsed vagina,

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endometrial venous aneurysm, hydrometra, uterine torsion, cryptorchidism, testicular neoplasia, venereal spirochetosis, and cystic mastitis.

DISEASES OF THE EXCRETORY SYSTEM The exact cause of urolithiasis in rabbits is unknown; however, diet, vitamin and mineral oversupplementation, and infections can be predisposing factors. Clinical signs are often nonspecific and include lethargy, inappetence, abdominal distension, urine scalding, perineal debris, and hematuria. A thickened bladder can sometimes be palpated on physical examination. A diagnosis can be made from radiographs and a urinalysis (e.g., crystals). A cystotomy is generally needed to treat urolithiasis. Reducing calcium in the diet may help decrease the likelihood of urolithiasis formation. Cystitis is a common problem in does. Diagnosis is generally made from a cytologic examination (e.g., inflammatory cells and bacteria) and bacterial culture and sensitivity. Opportunistic Gram-negative bacteria are the most common isolates. Treatment is ideally based on culture and antibiotic sensitivity results. Renal failure can occur in rabbits as a result of hypercalciuria (e.g., mineralization) or infectious agents (e.g., E. cuniculi). Clinical signs include polyuria and polydypsia, weight loss, inappetence, anemia, and lethargy. Diagnosis can be made based on radiology, chemistry panel, CBC, and urinalysis. An inverse calcium-to-phosphorus (Ca : P) ratio is generally indicative of renal failure. Survey radiographs may reveal mineral densities in the kidneys. Isosthenuria, in the face of an inverse Ca : P ratio, on a urinalysis can be used to confirm a diagnosis. Confirmation can be made via renal biopsy. Treatment should focus on monitoring fluid levels. Renal cysts, or renal polycystic kidney syndrome, have been reported in New Zealand white rabbits. Most affected rabbits have nonspecific clinical signs. Animals with clinical disease generally are lethargic and anorectic. Polycystic disease can be associated with hypercalcemia, hypercreatinemia, and arterial mineralization.34 E. cuniculi can cause renal disease in rabbits. Early in the disease course, the parasite causes segmental, granulomatous, interstitial nephritis, which can be seen as irregular depressions on the surface of the kidneys. However, interstitial fibrosis can occur late in the disease. Many times these changes do not cause a reduction in kidney function. (For more information about E. cuniculi, see Parasitic Disease.) “Red urine” is a common problem in rabbits. In these cases, veterinarians must determine whether a sample is red because of the presence of blood or porphyrin pigment. Blood indicates a breakdown in the epithelial barrier and is common with cystitis, uroliths, and neoplasia. The presence of porphyrin pigment is associated most with stress, dietary changes, or both. A thorough history will often provide insight into whether a recent stress event or dietary change occurred. A urinalysis should be done to determine if a color change in the urine is based on blood or pigment. The presence of red blood cells is diagnostic. Disease rule-outs for animals with blood in their urine include neoplasia (e.g., adenocarcinoma), endo-

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396 metrial aneurysms, urinary or reproductive tract polyps, cystitis, pyelonephritis, urolithiasis, disseminated intravascular coagulopathy, and lead toxicosis.

DISEASES OF THE GASTROINTESTINAL SYSTEM Dental disease, as previously mentioned, is common in pet rabbits. Clinical signs can vary, but generally include difficulty with prehension of food, anorexia, cachexia, dacryocystitis, abnormal facial swelling, and excessive salivation. Incisor malocclusion has been attributed to congenital factors, such as mandibular prognathism, and is seen at an increased rate in rabbits less than 1.5 kg or certain breeds (e.g., Netherland dwarf rabbits).2 Animals that suffer traumatic injuries to the jaws are also more susceptible to developing tooth malocclusion. Abnormal incisor growth can lead to the blockage of the nasolacrimal duct, resulting in epiphora. Incisor malocclusion can also lead to molar malocclusion, causing further dental complications. Trimming incisors is the primary method for correcting malocclusion, and this can be done using a highspeed Dremel tool (Dremel, Racine, WI) or dental drill (Figure 14-19). Incisors grow at a rate of 3 mm per week when the opposing incisor is present and 1 mm per day when there is no tooth opposition. Therefore, trimming should be done at approximately 3-week intervals when there is a malocclusion or no opposing incisor. Toenail clippers have been recommended for trimming incisors; we do not recommend this, as toenail clippers have been associated with tooth fracture. Where there is severe malocclusion, complete incisor extraction is recommended. This is a relatively easy procedure; however, it does require patience. Aggressive, rough manipulation of the teeth can lead to tooth fracture and, again, pain and discomfort. (This procedure is discussed in detail in Surgery.) If the entire root and germinal tissue are not removed, the teeth can regenerate, requiring additional surgery. Cheek teeth or molar malocclusion is characterized by crown elongation and the development of hooks and points. Coronal overgrowth (due to incisor overgrowth or poor occlu-

MANUAL OF EXOTIC PET PRACTICE

sion) requires not only that the spikes be removed but also that the crowns be reduced and the angulation be corrected for ideal occlusion. Maxillary cheek teeth and all but the first mandibular cheek teeth can be reduced to the level of the gingiva; however, this is not always necessary. In some cases, removing the points is sufficient. The gingiva naturally exposes the cranial aspect of the first mandibular cheek tooth. Ideally, when trimming or floating the cheek teeth, the procedure should be done with power equipment (e.g., flat fissure bur) to ensure rapid tooth removal without damaging the surrounding tissue. The use of rasps or clippers can create friction and rock the teeth, causing periapical and periodontal damage. For this reason, these tools are not recommended or should be used with extreme caution. Extraction of the cheek teeth is much more complicated than removal of the incisor and more traumatic to the rabbit. (Crossley DA, personal communication) When several extractions are required, it is best to perform multiple procedures to minimize the pain and discomfort to the animal. For example, extracting the teeth on one side of the jaw will allow the animal to still use the opposite side to process its food. (Cheek tooth extraction is discussed in detail in Surgery.) Incisor and cheek tooth root elongation can result in an overall deterioration of the tooth quality. Affected animals often present with epiphora, mandibular swelling, or nasal discharge. If left undiagnosed and untreated, root elongation can lead to abscess formation and osteomyelitis. Dental or skull radiographs can be used to evaluate the roots of the teeth, and if any are diseased, extraction should be considered. Mucoid enteropathy is a poorly understood condition that causes both constipation and diarrhea, especially in young rabbits. The etiology of this disease is unknown, but it may be associated with a reduction in cecal pH. Feeding rabbits a diet that is high in fiber appears to control cecal pH and reduce the likelihood of this disease. Because the etiology of this disease is unknown, treatment is based on providing supportive care, antibiotics against opportunistic bacteria, and probiotics. Rabbits are susceptible to a number of different infectious and parasitic diseases, many of which affect the gastrointestinal system. (A review of those diseases can be found in Infectious Disease.)

DISEASES OF THE RESPIRATORY SYSTEM

Figure 14-19 High-speed dental tools are recommended for trimming rabbit teeth.

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Upper respiratory tract disease (URTD) is a common presentation in rabbits. Affected rabbits often present with rhinitis and sinusitis. There are numerous potential causes of URTD in rabbits, including bacterial infections (e.g., P. multocida, B. bronchiseptica, and Staphylococcus spp.), foreign bodies, dental abscesses, and myxomatosis. Rabbits with URTD are often anorectic, as they cannot smell or locate their food. Serous to purulent nasal discharge is common. Epiphora is also a common finding. Many rabbits with clinical disease have snuffles. This is a generic term used to describe clinical signs, but it is attributed primarily to pasteurellosis. It is important to recognize that snuffles is nonspecific and may be associated with any of the previously mentioned causes. Rabbits with URTD often have “wet” forelegs as a result of the chronic nasal

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discharge. In severe cases of URTD, when a rabbit cannot move air through its nasal passages, it becomes dyspneic and breathes with an open mouth. Hematologic testing (e.g., CBC), culture and antibiotic sensitivity testing, radiographs, and endoscopy may be required to diagnose a specific cause of URTD. Treatment should be based on a specific diagnosis. Animals that are dehydrated and anorectic should be provided fluids and caloric support. In rabbits, lower respiratory tract disease is primarily associated with bacterial disease. Gram-negative bacteria (e.g., P. multocida, B. bronchiseptica, Klebsiella spp.) are the most common cause of pneumonia in rabbits. Although not as common, Gram-positive bacteria (Staphylococcus aureus) have also been associated with pneumonia. Viral diseases are not well-studied but likely play a limited role. Primary lung neoplasia can occur, but it is not common. Rabbits are more likely to develop secondary metastasis. Rabbits with pneumonia may present in either an acute or chronic disease state. Rabbits with acute pneumonia are often acutely depressed, lethargic, and anorectic. The rabbits may or may not have audible respiratory sounds (e.g., wheezing, crackling). On physical examination, the rabbit may be febrile, have nasal discharge, and have auscultable respiratory wheezing or crackling. With chronic pneumonic presentations, the rabbit may appear clinically normal (e.g., walking pneumonia) but may have abnormal lung sounds. Beyond the physical examination, a CBC, survey radiographs, tracheal wash with cytologic examination and culture and antibiotic sensitivity profile, and endoscopic examination can be used to diagnose the pneumonic condition and establish a prognosis. Pneumonia in rabbits generally carries a poor to grave prognosis. Treatment should be based on the diagnostic results. Fluoroquinolones and potentiated sulfonamides are excellent first-choice antibiotics, as they have excellent distribution to the lungs.

CARDIAC DISEASE Cardiac disease is a problem being encountered with increased frequency in captive rabbits. The primary reason for this is improved husbandry and the resultant increased longevity. Rabbits with cardiac disease may be dyspneic, tachypneic, exercise intolerant, lethargic, and depressed and have edematous extremities. A complete diagnostic work-up is required to characterize the status of the patient and provide the client a prognosis. Beyond a physical examination and careful cardiac auscultation, suspect cardiac cases should have an electrocardiogram done to determine if there are any abnormal arrhythmias, survey radiographs to evaluate heart and great vessel size, and an echocardiogram to assess the valves, muscle thickness, and cardiac function. A CT scan or MRI can also be done to evaluate the heart in more detail. Cardiomyopathy can occur in all breeds, but it occurs at a higher incidence in giant breeds.2 Affected animals often develop myocardial fibrosis. A number of different causal factors have been associated with cardiomyopathy in rabbits, including bacterial and corona-virus infections and vitamin E deficiency. Arteriosclerosis is also a common finding in rabbits fed a vitamin D and calcium-rich diet. Although a diagnosis

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of arteriosclerosis is most often made at necropsy, in some cases, a diagnosis can be made from survey radiographs. An increased mineralized density in the aorta is the most common radiographic finding. Congestive heart failure is common in middle-aged to geriatric rabbits. Rabbits with congestive heart failure should be stabilized and the excess burden on the heart controlled. Furosemide (1-4 mg/kg IV or IM) and 2% nitroglycerine ointment (1/8 strip on the ear) can be used to eliminate excess fluid and improve contractility.4 In cases of pleural effusion, a thoracocentesis may be performed. After the animal’s immediate condition is stabilized, further work should focus on diagnosing the cause of the congestive heart failure. Because rabbit cardiac drug doses are empirical and frequently inaccessible, doses for cats or ferrets can be used.4 It is important to monitor rabbits for adverse effects to cardiac drugs, as would be done in domestic pets.

HEAT STROKE Rabbits are extremely sensitive to elevated environmental temperature, especially when combined with high humidity. Because rabbits are unable to sweat, and panting is inefficient for them, they rely on the large surface area of the ears to serve as the site of evaporative cooling. Rabbits with heat stroke may present for lethargy, lateral recumbency, dyspnea, cyanosis, diarrhea, hyperthermia (>104° F), and seizures. Heat stroke carries a grave prognosis. Cooling a hyperthermic rabbit too quickly can lead to iatrogenic hypothermia, which can exacerbate the physiologic shock the animal experiences. In addition to cool water baths, 70% isopropyl alcohol can be applied to the ears and extremities to facilitate body cooling. An intravenous catheter should be placed, and fluids should be provided to maintain the intracellular and extracellular fluid balance. Antiinflammatories (e.g., meloxicam and carprofen) should be administered to minimize the catastrophic effects of the inflammatory response. Parenteral antibiotics are warranted to minimize the impact of opportunistic infections. Because some of the deleterious effects associated with heat stroke are not immediately apparent (e.g., cell death resulting from fluid shift), veterinarians should monitor animals closely for 7 to 14 days after the presentation.

THERAPEUTICS Many of the therapeutics used to treat dogs and cats can also be used to manage rabbit patients, which reduces the burden of maintaining an appropriate rabbit pharmacy for the veterinarian. However, in some cases, it may be necessary for the veterinarian to obtain rabbit-specific formulations, that are flavored (e.g., fruity) or more specific to the needs of the rabbit. Fortunately, there are a number of compounding pharmacies that can assist with designing specific formulations for these animals. The following section is an introduction to the therapeutics commonly used to treat rabbits in captivity. (For a more extensive list, see Carpenter, JW: Exotic Animal Formulary, St. Louis, 2005, WB Saunders.)

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Fluid Therapy Rabbits that are dehydrated require immediate attention. The redistribution of fluids that results from dehydration can lead to a number of internal changes. One of these changes, occurring when fluid is resorbed from the intestinal tract, can lead to reduced intestinal motility and catastrophic changes to the microflora. The maintenance fluid rate for rabbits is 80 to 100 ml/kg/ day. For rehydration the rate can be increased to 10 to 20 ml/ kg/hr over the first few hours.9 In addition to providing the maintenance fluids for an anorectic rabbit, the veterinarian should also estimate and correct the animal’s fluid deficit. Skin elasticity, mucous membrane moisture, capillary refill time, femoral pulse, positioning of the globe, and packed cell volume and total solids concentration are useful indicators for assessing dehydration. Slowed skin elasticity and tacky mucous membranes generally suggest an animal is 3% to 5% dehydrated. Those findings, in combination with a decreased capillary refill time, increased packed cell volume and total solids, and increased “thready” pulse, suggest 5% to 8% dehydration. All of these findings, in combination with the presence of sunken eyes, suggest an animal is more than 8% dehydrated. It is essential that the fluid deficit be replaced promptly (e.g., over 24 hours) to reestablish the fluid balance between the extraand intracellular spaces. When selecting fluids for replacement, it is important to consider the rabbit’s physiologic status. Animals with elevated osmolalities should be provided hypotonic fluids to prevent worsening the fluid dynamics. Normasol (293 mOsm/L), lactated Ringer’s (273 mOsm/L), and 0.9% saline (308 mOsm/ L) represent isotonic crystalloids that can be used for fluid replacement. Dextrose (253 mOsm/L) can also be used, especially if the animal is hypoglycemic or if there is a need to rapidly replace the intracellular space. Animals that are hypoproteinemic may require colloid fluid replacement. Colloids can be used to offset the loss of oncotic pressure that can occur with hypoalbuminemia. Rabbits that are anemic may also require a blood transfusion or oxyglobin, a synthetic hemoglobin replacement. All rabbit patients should be monitored closely during fluid replacement so that neither fluid overload nor insufficient fluid replacement occurs. Pulmonary edema, ascites, and/or anasarca may occur in animals provided excess fluids. Animals provided insufficient fluids will remain clinically dehydrated. In these cases, the fluid deficit should be reevaluated and the appropriate fluid volume provided. Fluids can be delivered per os (PO), subcutaneously (SC), intravenously (IV), intraosseously (IO), or intraperitoneally (IP). The route of administration should be based on the needs of the rabbit. If the animals is mildly dehydrated and has a functional gastrointestinal tract (e.g., no diarrhea), then fluids should be provided PO. This represents the natural route of fluid administration. For mildly dehydrated animals that will not tolerate PO fluids or have diarrhea, SC fluids are recommended. SC fluids can be administered on the dorsum between the scapulae or over the lateral body wall. Animals that are moderately to severely dehydrated should be given fluids IV

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or IO. The primary IV sites we use are the cephalic, lateral saphenous, or auricular veins. The site for catheter placement should be sterilely prepared before catheter placement. The largest bore catheter that can be fitted comfortably into the vein should be used. For most large breeds, a 20- or 22-gauge catheter can be used, whereas a 24- or 26-gauge catheter may be required for smaller breeds. A smaller bore catheter should be used for the auricular vein. The peripheral site for IO catheter placement is the femur. A surgical preparation using a disinfectant (e.g., Betadine or chlorhexidine) should be done to minimize contamination at the catheter site. A local anesthetic should be used to minimize the discomfort associated with the procedure. The trochanteric fossa is the landmark for catheter insertion. A spinal needle should be used for the procedure, as it has a stylet and will prevent the needle from being clogged with a bone core. The IO route is preferred in cases where the animal is severely dehydrated and the veins are collapsed. IP fluids are generally reserved for moderately dehydrated rabbits when IO or IV catheters are not available or for juvenile rabbits. To administer these fluids, place the rabbit in dorsal recumbency and insert the needle in the caudal abdomen. The needle should be inserted at a 20- to 30-degree angle to the body. Aspirating before delivering the fluids should ensure that the needle is not in the viscera.

Antimicrobial Therapy ANTIBACTERIAL AND ANTIFUNGAL AGENTS Antimicrobial therapy is ideally based on culture and antibiotic sensitivity testing. When selecting an antimicrobial for a rabbit, it is important to avoid those compounds that may predispose the rabbit to dysbiosis, such as lincomycin, ampicillin, amoxicillin, amoxicillin-clavulanic acid, cephalosporins, clindamycin, penicillins, and erythromycin. Because most infections in rabbits are associated with opportunistic Gramnegative bacteria, initial antibiotic selection should be based on broad coverage against these bacteria. Enrofloxacin and trimethoprim-sulfadiazine are two excellent first-choice antibiotics. For a complete list of antibiotic compounds, see Box 14-3. A list of common antifungal compounds can be found in Box 14-4.

Antiparasitic Therapy Rabbits are susceptible to a number of endo- and ectoparasites. The most common endoparasites are coccidia and nematodes. The drugs and doses commonly used to treat these parasites are found in Box 14-5. Ectoparasites, including mites and lice, are also common.

Nonsteroidal Antiinflammatories Nonsteroidal antiinflammatories (NSAIDs) are routinely used to manage inflammation and pain in rabbits. Some of their most common uses include treating musculoskeletal pain associated with trauma, arthritis, soft tissue pain from dental disease, and gastric stasis. Historically, flunixin meglumine was

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BOX 14-3

Antibiotic Agents Commonly Used to Treat Bacterial Infections in Rabbits

Chloramphenicol Ciprofloxacin

Enrofloxacin Metronidazole Penicillin Penicillin G, benzathine Penicillin G procaine Rifampin/clarithromycin Silver sulfadiazine Sulfadimethoxine Trimethoprim sulfadimethoxine

30 mg/kg PO BID 10-20 mg/kg PO BID Ophthalmic 0.3%: 1 drop BID-TID 5 mg/kg PO, SC, IM, IV BID35 20 mg/kg PO BID 40 mg/kg PO SID × 3 days 40,000-60,000 IU/kg SC, IM SID × 5-7 days 42,000-60,000 IU/kg SC, IM q48h 42,000-84,000 IU/kg SC, IM SID R-40 mg/kg; C-80 mg/kg PO BID Topical SID; safe if ingested 10-15 mg/kg PO BID 30 mg/kg PO BID

BOX 14-5

Antiparasitic Drugs Commonly Used to Eliminate Parasites in Rabbits

Drug Albendazole Ivermectin

Fenbendazole

Lime-sulfur dip (2%-3%) Sulfadimethoxine

Dosage 7.5-20 mg/kg PO SID 0.4 mg/kg SC q7days × 2-3 wk 20 mg/kg SID 10 mg/kg PO, repeat in 14 days Topically q7days × 4 wk 10-15 mg/kg PO BID

Clotrimazole Fluconazole Ketoconazole Lime-sulfur dip (2%-3%) Miconazole

Topical 25-43 mg/kg IV slowly BID 10-40 mg/kg PO SID × 14 days Topical q 5-7 days × 4 wk

Coccidia

Nonsteroidal Antiinflammatory Drugs Commonly Used to Manage Inflammation and Pain in Rabbits

Meloxicam

0.3 mg/kg PO SID 0.2 mg/kg IM or SC SID 1-2 mg/kg IM or SC BID; limit to 3 days to minimize gastric ulcer formation 2-4 mg/kg SC SID 1.5 mg/kg PO BID 3 mg/kg IM or SC BID

Carprofen Ketoprofen

BID, twice a day; IM, intramuscular; PO, per os; SC, subcutaneous; SID, once a day.

Topical SID × 2-4 wk

BID, twice a day; PO, per os; SC, subcutaneous; SID, once a day.

the only compound available for rabbits. This drug serves an important purpose in managing pain, but it has to be used judiciously because of the potential for gastric ulcer formation. The newer generation NSAIDs (e.g., carprofen and meloxicam) are more selective and less likely to cause complications. A list of commonly used NSAIDs and their doses can be found in Box 14-6.

Steroids The use of steroidal compounds is controversial in rabbits. In the past, steroids have been recommended for managing inflammation associated with neurologic disease (e.g., trauma or E. cuniculi infection) or shock. However, hydrocortisone has been found to cause significant immunosuppression in rabbits.36 Because of the high likelihood for immunosuppres-

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Ectoparasites

BOX 14-6

Flunixin meglumine

Antifungal Drugs Commonly Used to Treat Rabbits

E. cuniculi Nematodes

BID, twice a day; PO, per os; SC, subcutaneous; SID, once a day.

From Carpenter JW, Mashima TY, Rupiper DJ: Exotic Animal Formulary, ed 2, Philadelphia, 2001, WB Saunders. BID, twice a day; IM, intramuscular; IU, international unit; IV, intravenous; PO, per os; SC, subcutaneous; SID, once a day; TID, three times a day.

BOX 14-4

Parasite Nematodes; E. cuniculi Nematodes, ectoparasites

sion, steroids should be used judiciously. We generally reserve their use for the treatment of spinal disease (prednisone sodium succinate, 1 mg/kg IV) and dermatologic conditions (e.g., flea bite allergy or extreme hypersensitivity) (prednisone, 0.5-1 mg/kg SID to BID, taper dose).

Cardiac Drugs Long-term therapy for heart disease may require treatment with diuretics (furosemide, 1-4 mg/kg IM) and other compounds required for a particular cardiac disease presentation. The use of cat and ferret dosages for cardiac drugs is advocated. Monitoring for drug side effects should be done as in feline medicine.

Emergency Drugs Rabbits that develop respiratory or cardiac arrest should be treated immediately with the appropriate emergency drugs. Dopram may be used for respiratory arrest, whereas epineph-

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MANUAL OF EXOTIC PET PRACTICE

Emergency Drugs Commonly Used to Treat Rabbits

Epinephrine Furosemide Hetastarch Lactated Ringer’s solution Lidocaine

0.2 mg/kg IV or IT 1-4 mg/kg IM 20 ml/kg IV 10-20 ml/kg/hr 1-2 mg/kg IV

IM, intramuscular; IT, intratracheal; IV, intravenous.

rine should be administered during cardiac arrest. A list of commonly used emergency drugs and their doses is listed in Box 14-7. Rabbits have atropinases, so the use of atropine is not commonly recommended. Glycopyrrolate can be used to help manage bradycardia; however, it is important to consider that this compound can increase the viscosity of tracheal secretions and impede the airflow to the lungs. Intubating and ventilating animals can minimize this risk.

Transfusions In cases of severe anemia (PCV < 10%) or acute blood loss (20%-25%), a blood transfusion is recommended. The blood volume of a rabbit is estimated to be between 55 and 65 ml/ kg.37 Cross-matching for transfusion in rabbits is not necessary for a first-time recipient.37 The donor must be healthy and must not have a history of infectious disease or neoplasia. A healthy blood donor can have up to 10 ml/kg of blood collected. The jugular vein is the recommended site for blood collection from the donor rabbit. The blood should be collected with a citrate anticoagulant that is mixed with blood at a rate of 1 : 3.5 (citrate : donor blood). The blood transfusion should be done within 4 to 6 hours of collection at a rate of 6 to 12 ml/kg/hr.

Nutritional Support Anorexia in rabbits should be treated as an emergency. When rabbits stop eating, physiologic changes can occur in the gastrointestinal tract (e.g., mucosal atrophy) that alter the gastrointestinal permeability. From this, a cascade of events can occur (e.g., reduced motility, decreased water absorption, pH alterations, microbial flora changes) that lead the animal into a life-threatening situation. The first step in managing these cases should be to restimulate the gastrointestinal tract, and the second step is to diagnose the cause of the anorexia. Providing nutritional support is the best method for restimulating the gastrointestinal tract. Motility modifiers can also be used, but these should be given only when there is no concern for a gastric or small intestine foreign body (e.g., trichobezoar). The preferred method for administering nutritional support to mild or moderately ill animals is syringe feeding. In more severe cases, an orogastric tube or nasogastric tube should be

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placed. A 3.5- to 5.0-French pediatric feeding tube can be used as a nasogastric tube. Before placing the tube, it should be marked to estimate the distance between the nare and the stomach. Placement can be achieved by inserting the tube through the ventral medial nasal meatus and advancing it ventromedially. The rabbit’s head should be maintained in a natural position during the procedure to facilitate passage of the tube. Placement of the tube should be evaluated using survey radiographs. Once placed, the tube can be secured to the skin covering the nasal bones using suture. A pharyngostomy or gastrotomy tube can also be placed to facilitate feeding. Enteral diets used for dogs or cats are not appropriate for use in rabbits. The domestic pet enterals are based on the nutritional needs of omnivores (dogs) and carnivores (cats) and will not meet the needs of an herbivore (rabbit). Oxbow critical care for herbivores (Oxbow Pet Products, Murdock, NE) is our preferred enteral. High-fiber Ensure or Ensure plus (Ross Laboratories, Columbus, OH) can also be used, but, again, these are not considered ideal. Although less complete, a variety of other pelleted rations, leafy greens, and vegetable baby foods can also be used as enterals.9 Label directions should be followed closely to ensure that the rabbit receives the appropriate quantity of calories. In cases where homemade diets are used, calorie content of the enteral constituents should be estimated or determined using available resources in the literature or on the Internet. The provision of nutritional support should continue until the rabbit is free-feeding on its own.

SURGERY Preoperative and Postoperative Considerations Although fasting is not necessary in rabbits because they cannot vomit, eliminating access to food 1 to 2 hours before inducing anesthesia minimizes the amount of food in the oral cavity and improves the direct visualization of the larynx for intubation. We recommend performing presurgical blood work to determine the physiologic status of the rabbit patient. This information can be used to provide the client with a prognosis related to the anesthetic procedure. Any rabbit with preexisting respiratory disease should be thoroughly evaluated before it undergoes a procedure. Rabbits have a small lung capacity, and any pulmonary disease can have a negative impact on the anesthesia event. Postoperative considerations for rabbits include minimizing the likelihood for anorexia, providing analgesia and antibiotics (when necessary), and monitoring fluid therapy. Thermal support should be provided until the rabbit has recovered and can maintain a normal body temperature. Rabbits should be recovered in a quiet area. Excessive noise or commotion can lead to increased stress (e.g., increased cortisol), immunosuppression, and decreased healing. Animals should be offered their normal food for a prolonged stay and a comfortable substrate in their cages.

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HCl, medetomidine, acepromazine) and analgesics can also be used for induction or as maintenance anesthetics. Doses for these compounds can be found in Tables 14-2, 14-3, and 14-4. One compound that should be avoided is Telazol (Fort Dodge Animal Health, Fort Dodge, IA), as it has been shown to cause severe renal tubular necrosis in rabbits.38 We recommend intubating rabbits for any procedure that requires general anesthesia. When an animal is intubated, it gives the anesthetist more control over the patient. Rabbits that are not intubated and experience respiratory arrest are rarely recovered. Intubating rabbits can be difficult. These animals have long, narrow oral cavities, which restrict direct visualization of the larynx. Several techniques have been described for intubating rabbits. Blind intubation using a 2.0

Anesthesia Anesthetic selection should be based on the procedure being performed. For example, in a case where a localized abscess needs to be lanced, a local anesthetic (e.g., lidocaine, prilocaine) and an opioid can be used to perform the procedure. Many veterinarians anesthetize rabbits using only inhalant anesthesia. Rabbits can be induced using a face mask or induction chamber. We prefer to induce rabbits with additional compounds before we provide inhalants, because it more effectively relaxes the animal for intubation. A benzodiazepam (e.g., diazepam, midazolam) and opioid are routinely used to provide relaxation and analgesia before inducing the animal. Other commonly used veterinary anesthetics (e.g., ketamine

TABLE 14-2

Induction and General Anesthetics for Rabbits

Agent

Dose (mg/kg)

Route

Sedation

Analgesia

Acepromazine Diazepam/midazolam Fentanyl (can cause respiratory depression; can be partially reversed with buprenorphine) Glycopyrrolate

1 mg/kg 0.5-2 mg/kg 0.2-0.5 mg/kg

IM IV, IM IM

++ + to + + + to + + +

* * + to +++

0.01 mg/kg 0.1 mg/kg 10-50 mg/kg 0.1-0.5 mg/kg

IV SC, IM IM IM

++ to + ++ + to + + +

+ +

2-5 mg/kg

IM

+ to + +

+

3-6 mg/kg

IV

Ketamine Medetomidine (reversible with atipamizole) Xylazine (reversible with atipamizole 1 mg/kg or yohimbine 0.2 mg/kg IV) Propofol (light anesthesia; can cause apnea, use with caution in un-intubated patients)

From Harcourt-Brown F: Textbook of Rabbit Medicine, London, 2002, Elsevier; Flecknell P: Laboratory Animal Anaesthesia, ed 2, San Diego, 1996, Academic Press. IM, intramuscular; IV, intravenous; SC, subcutaneous. *, none; +, minimal; ++, moderate; +++, heavy.

TABLE 14-3

Analgesics Commonly Used for Captive Rabbits

Agents

Dose (mg/kg)

Route

Comments

Buprenorphine Oxymorphone Butorphanol

0.01-0.05 0.05-0.2 0.1-0.5

IV, SC IM, SC IM, SC, IV

Analgesia for 6-12 h; for acute soft tissue, GI, and urogenital pain q8-12h Analgesia lasts 2-4 h; can be used with acepromazine to deliver sedation

Fentanyl Carprofen

0.2-0.3 2-4 1.5 0.2 0.3 1-2 3

IM SC PO IM, SC PO IM, SC IM, SC

Meloxicam Flunixin meglumine Ketoprofen

q24h q12h q12h q12h q12h

From Harcourt-Brown F: Textbook of Rabbit Medicine, London, 2002, Elsevier; Heard DJ: Anesthesia, analgesia, and sedation of small mammals. In Queensberry KE, Carpenter JW, editors: Ferrets, Rabbits and Rodents: Clinical Medicine and Surgery, St Louis, 2004, WB Saunders; Paul-Murphy J, Ramer JC: Urgent care of the pet rabbit, Vet Clin North Am Exot Anim Pract 1(1):125-152, 1998. IM, intramuscular; IV, intravenous; PO, per os; SC, subcutaneous.

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MANUAL OF EXOTIC PET PRACTICE

Common Anesthetic Combinations Used to Anesthetize Rabbits

Agents

Dose (mg/kg)

Route

Duration/anesthesia

Medetomidine Ketamine Butorphanol Fentanyl Midazolam Ketamine Xylazine Ketamine Diazepam

0.2 mg/kg 10 mg/kg 0.05 mg/kg 0.3 ml/kg 0.5-2 mg/kg 20-35 mg/kg 3-5 mg/kg 20-35 mg/kg 1-5 mg/kg

SC, IM

Anesthesia 30-40 min; recovery 1-4 h

IM IV IM/IV

Anesthesia 20-40 min with good muscle relaxation; recovery 1-2 h Anesthesia 25-35 min; recovery 1-2 h

IM/IV

Anesthesia 20-30 min; recovery 11/2 h

From Harcourt-Brown F: Textbook of Rabbit Medicine, London, 2002, Elsevier; Flecknell P: Laboratory Animal Anesthesia, ed 2, San Diego, 1996, Academic Press. IM, intramuscular; IV, intravenous; PO, per os; SC, subcutaneous.

to 2.5 outside diameter (OD) endotracheal tube can be done, but it can be associated with laryngeal trauma.9 To perform the procedure, the head should be elevated and neck stretched. The endotracheal tube should be gently lowered into the trachea. This technique requires patience and skill. Practicing this technique on recently expired rabbits is one way of obtaining experience with this procedure. Our preferred method for intubating rabbits is via an endoscope.39 A 2.7-mm rigid endoscope can be used to directly visualize the larynx and ensure passage of the tube into the trachea. In those cases where laryngospasms limit access to the trachea, lidocaine can be applied topically. Rabbits should be monitored closely during a surgical procedure. An ultrasonic Doppler, ECG, and/or pulse oximeter can be used to monitor the heart rate and rhythm during the procedure. A respiratory monitor can be used to monitor respirations. We prefer to use a ventilator during rabbit procedures to ensure that the rabbit is receiving an appropriate dose of anesthesia and oxygen. The ventilator also ensures that the animal is respiring. A capnograph can be added in line to measure carbon dioxide levels and determine the rabbit’s respiratory efficiency. In addition to the capnograph, the pulse oximeter can provide additional blood gas information by providing an estimate of oxygen saturation levels. Rabbits should be provided thermal support during a procedure, and its body temperature should be measured throughout the procedure. Routine anesthetic monitoring during a procedure is essential to a veterinarian’s success and will limit the occurrence of anesthetic deaths.

Surgical Procedures The majority of the surgical procedures performed on rabbits are associated with the reproductive tract, gastrointestinal tract, and integument. Of course, a rabbit can develop a need for surgical procedures on other systems as well, but, in our experience, they are less common. When faced with the need to perform a surgery on a rabbit, the veterinarian must identify the specific anatomic features that will be affected by the procedures and familiarize himself or herself with any specific

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Figure 14-20 When shaving rabbit fur for a surgical procedure, be careful to avoid tearing the delicate skin.

indications on tissue handling methods. In cases where there are no specific descriptions for a surgical procedure in rabbits, veterinarians may rely on those surgical descriptions reported in canine and feline surgery. When preparing a rabbit for surgery, it is important to adhere to strict aseptic techniques. Because rabbit skin is thin and tears easily, care should be taken when clipping the fur. We generally clip the fur with a small, battery-powered clipper or shave the fur with a razor. Holding the skin tight during fur removal can help reduce the likelihood of tearing the skin (Figure 14-20). There are a variety of methods that can be used to aseptically prepare a surgical site. Chlorhexidine and Betadine are excellent disinfectants. We generally use physiologic saline instead of alcohol during the preparation because it does not act as a heat sink, lowering the rabbit’s body temperature. The surgical instruments used for domestic pet surgery can also be used for rabbit surgery. Because of the small size of some dwarf rabbit breeds, ophthalmic instruments can also be beneficial. Microscopic loupes are helpful when manipulating

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fine tissues. Radiosurgery can be used to provide hemostasis during a procedure and minimize tissue trauma.

REPRODUCTIVE SURGERY Orchiectomy or ovariohysterectomy should be recommended for rabbits when the pet owner has no interest in reproducing the animal. This prevents unwanted pregnancies in mixed gender populations, reduces the risk for certain neoplastic conditions, and can reduce the likelihood for certain behaviors (e.g., aggression in males). It is generally recommended that males be castrated after 3 months of age and females be spayed after 5 months. The procedure can be performed at an earlier age if deemed appropriate, such as may be the case in large, mixed colonies, but this may affect the growth of the animal (e.g., smaller size).

Orchiectomy The inguinal canals of a rabbit are open throughout the animal’s life. Therefore, a male rabbit can actively move its testicles between the scrotum and body cavity. Because the inguinal canals remain open, it is important to either perform a closed orchiectomy, or to close the inguinal canal after performing an open orchiectomy. If the inguinal canals are left open, it is possible for the animal to herniate omentum or bowel into the scrotal area. There are two approaches that can be taken when making an initial incision. Either a prescrotal incision can be made, similar to that made for the dog, or scrotal incisions can be made, similar to those recommended for the cat. When performing the scrotal approach, each testicle is removed through a separate incision. If the tunic is not incised, a closed procedure can be done. At least two sutures should be placed on the cord in case a suture fails. Once ligated, the cords can be replaced into the scrotum, and the scrotum closed with an absorbable suture using a subcuticular pattern. Tissue glue can also be used to complete the skin closure. For the prescrotal incision, each testicle must be retrograded out of the incision. The same technique described earlier can be used for the orchiectomy, or a suture can be placed around the tunic at the cranial aspect of the incision, an open castration done, the remnant cord directed cranially past the suture placement, and the suture ligature tied down to close the canal. Again, a subcuticular closure or tissue glue can be used to close the skin incision. The animal should be monitored postoperatively for any swelling or discharge in the area of the incision or scrotum.

Ovariohysterectomy An ovariohysterectomy should be recommended for any female rabbit not intended for breeding. It is recommended that the procedure be done when the animal is young (45 min) under anesthesia with limited monitoring ability. Regardless of the present limitations, veterinarians should be aware that these techniques are available and that reference material exists that depicts normal anatomy.9 As guinea pig owners continue to demand high-quality care for their pets, these imaging techniques will likely become more commonplace in small mammal practice for these patients.

Microbiology Microbiologic samples can be obtained for diagnosis of various guinea pig infections. Exudates from nasal or ocular secretions can be examined for abnormal flora. Some bacterial organisms (e.g., Bordetella bronchiseptica) are difficult to culture. Laboratories need to be advised as to the organisms in question so they can perform more specific diagnostic tests to obtain organism identification. Abscesses are another disease condition that may require culture for proper treatment. Because guinea pigs form caseous abscesses, the purulent debris itself is typically not useful for bacterial isolation. For the best chance of identifying organisms within an abscess, a portion of the abscess capsule should be submitted to the laboratory. Both aerobic and anaerobic bacterial cultures should be requested, especially if it is suspected that the abscess may have originated from a dental problem.

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Parasitology Roundworms and coccidia are seen in guinea pigs and can be identified by fecal flotation or fecal direct smear, as in other species. Cryptosporidiosis has also been reported.10 Identification of Cryptosporidium organisms usually requires acid-fast staining or immunofluorescent antibody testing in addition to fecal flotation. Ectoparasites (e.g., mites, lice, fleas) commonly infest guinea pigs. Visualization of the parasites and/or their waste, as well as skin scrapings, microscopic examination of hair follicles, and tape preparations can be useful in the identification of these parasites.

COMMON DISEASE PRESENTATIONS Gastrointestinal

Figure 17-13 Cranial-caudal view of a guinea pig skull. Note the lack of yellow enamel on the cranial surface of the incisors and the severe occlusal angle of the premolars and molars.

ENTEROTOXEMIA The term enterotoxemia refers to the overgrowth of toxinproducing bacteria in the GI tract, particularly Clostridium difficile. This can occur with stress, an abrupt change in diet, GI stasis, or inappropriate antibiotic administration. The GI flora of guinea pigs is predominantly Gram positive, and administration of antibiotics with a primarily Gram-positive spectrum (e.g., beta-lactam antibiotics, macrolides, lincosamides) can lead to the depletion of normal gut flora, allowing colonization by opportunistic bacteria (e.g., Gram-negative organisms, Clostridium spp.).

DENTAL DISEASE The dentition of the guinea pig is described as aradicular hypsodont, meaning all teeth grow throughout the animal’s life and are open-rooted.1 The dental formula of the guinea pig is I 1/1, C 0/0, P 1/1, M 3/3. The mandible of these animals is wider than the maxilla. The occlusal angle of the molars and premolars in the guinea pig is quite severe when compared with that of rabbits and other rodents (Figure 17-13). Guinea pigs lack the layer of yellow enamel present on the rostral surface of the incisors in most other rodents. Dental malocclusion is a common disease process in guinea pigs. The development of malocclusion can be the result of multiple etiologies or a combination of factors (e.g., improper diet, genetics, trauma). Incisor malocclusion alone is rare in guinea pigs, so a thorough oral examination is necessary to determine the presence of cheek teeth malocclusion. When the molars and premolars do not occlude properly, overgrowth of the crowns and reserve crowns takes place. With improper wear, sharp points can form on the buccal aspects of the maxillary cheek teeth and the lingual aspects of the mandibular cheek teeth. The mandibular cheek teeth can overgrow to such an extent that they entrap the tongue, making eating and drinking difficult (Figure 17-14). Clinical signs can include inappetence or dysphagia, decreased fecal output, ptyalism, poor coat quality, and leth-

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Figure 17-14 Guinea pig with malocclusion. Note the severe overgrowth of the mandibular premolars and molars, causing entrapment of the tongue. (Courtesy Michelle G. Hawkins, VMD, DABVP [Avian].)

argy. Abscesses commonly occur at the apical aspects of overgrown molars and premolars. Because the premolars and molars are elodont (open-rooted), it is possible that these teeth can overgrow and impinge on the nasolacrimal duct, causing ocular and nasal discharge. The reserve crowns can also overgrow into the nasal cavity, where oral bacteria may be seeded, causing rhinitis and sinusitis. Diagnosis of dental disease is based on physical and oral examination findings. Careful palpation of the ventral mandible and maxilla may reveal bony protuberances corresponding to overgrowth of the apical surfaces of the cheek teeth. Proper instrumentation is very important for adequate visualization of the oral cavity. Dental speculums and pouch dilators are invaluable aids in obtaining a good view of the molars and premolars (Figure 17-15). However, many abnormalities can

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Figure 17-15 Oral examination of an anesthetized guinea pig aided by the use of incisor dilators and pouch dilators.

Figure 17-17 Computed tomography image of a guinea pig with mild to moderate malocclusion of the premolars and molars. (Courtesy University of California–Davis.)

Figure 17-16 Examination of the oral cavity of an anesthetized guinea pig using a rigid endoscope.

be overlooked in a conscious animal, so anesthesia is often required to obtain a thorough oral examination. Visualization of the oral cavity and occlusal surfaces can be facilitated by the use of an endoscope (Figure 17-16). Most endoscopes available in veterinary practice have a degree of angulation of the field of view, allowing greater focal area. Abnormal oral examination findings may include an uneven occlusal surface or angle of the cheek teeth, formation of sharp points with or without associated ulceration of the oral mucosa, food impaction, and abnormal spaces (diastema) between teeth. Imaging, including routine radiographs, magnified skull radiographs, and computed tomography can be incorporated to better evaluate the extent and seriousness of the process (Figure 17-17). Radiographic studies should include dorsoventral, lateral, and right and left oblique views. Treatment is centered on restoring a normal occlusal plane to the teeth, a procedure that should be performed while the

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animal is under general anesthesia. Unstable animals should be provided with supportive care before the anesthetic event. A high-speed surgical dental handpiece can be used to restore a normal occlusal plane and to reduce any sharp points. Handheld trimmers are not acceptable, as they tend to crush teeth and can cause fractures and pulp exposure.1 Dremel tools are also not considered acceptable because the small size of the guinea pig oral cavity will not allow for appropriate trimming. With any dental instruments, care should be observed to minimize oral soft tissue trauma. When a guinea pig is diagnosed with dental malocclusion, it is important to convey to the owners that this will be a lifelong problem for their pets. With many cases, routine occlusal adjustments will be necessary for the remainder of the animal’s life.

TYZZER’S DISEASE (C. PILIFORME) Tyzzer’s disease is the common term for bacterial enteritis caused by Clostridium piliforme. Infection with this organism is more commonly described in small rodents such as mice and hamsters, although it has also been described in guinea pigs as well as other mammalian groups.11 Tyzzer’s disease often presents acutely, with signs such as diarrhea, depression, and poor coat quality, often progressing rapidly to death. Transmission is via the fecal-oral route. Guinea pigs can be infected with C. piliforme without showing clinical signs. Animals that are carrying the organism but showing no signs of illness have still been found to shed organisms.12 Animals that carry the organism and then become immunocompromised (e.g., stress, immunosuppressive drug therapy) often develop clinical disease. Clostridium piliforme is an intracellular bacterium that will not grow on routine culture media, so antemortem

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diagnosis is difficult. Tyzzer’s disease begins as an intestinal infection, but later spreads to the liver hematogenously, causing areas of necrosis.11,13 Positive identification of the organism on necropsy requires examination of hematoxylin and eosin or silver staining of affected tissues. Treatment should consist of fluid and nutritional support as well as appropriate antibiotic therapy. Unfortunately, the progression of the disease is rapid, making treatment unrewarding. Prevention is key, focusing on owners’ reducing housing stress, providing a proper diet, and maintaining a clean environment.

CRYPTOSPORIDIOSIS Cryptosporidiosis has been described as causing disease in guinea pigs in a laboratory setting.10 Diagnosis was made histologically from affected animals that lost weight, had diarrhea, and suffered an acute death. Cryptosporidium organisms were found in the brush border of the intestinal tract from the duodenum to cecum, with associated inflammation.10

CORONAVIRUS Suspected coronavirus infection has been described in young guinea pigs.14 Clinical signs in affected animals included anorexia, weight loss, and diarrhea.14 Approximately half of the affected animals in the reported outbreak recovered.14 Necropsies performed on animals that died or were euthanized showed lesions consistent with coronavirus infection resulting from an acute to subacute necrotizing enteritis involving primarily the distal ileum.14 Coronavirus-like virions were identified on transmission electron microscopy of feces collected from the lower GI tract at necropsy.14

SALMONELLOSIS Outbreaks of salmonellosis have occurred in guinea pig colonies. Some of the organisms that have been isolated include S. enteritidis, S. dublin, S. florida, S. poona, and S. bredeney.15,16 Salmonellosis has been described to affect guinea pigs in acute and chronic disease processes. With acute salmonellosis, guinea pigs often die after a brief illness characterized by nonspecific signs of illness and diarrhea.15 Usually only a few pathogenic abnormalities are found at necropsy, and confirmation of a diagnosis is dependent on culture of affected tissues. Chronic salmonellosis is a wasting disease, lasting several weeks. Splenomegaly, hepatomegaly, and mesenteric lymphadenopathy are common postmortem findings in affected animals.15 Guinea pigs that recover can remain chronic, intermittent shedders of Salmonella organisms.

YERSINIA PSEUDOTUBERCULOSIS Yersinia pseudotuberculosis causes several disease syndromes in guinea pigs.15,17 The most common presentation affects the mesenteric and colonic lymph nodes, infiltrating these lymph nodes with caseous nodules. Clinically, affected guinea pigs will lose weight, have diarrhea, and develop a generalized lymphadenopathy. Transmission of the disease can be either vertical or horizontal.

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Musculoskeletal HYPOVITAMINOSIS C As stated previously, guinea pigs require dietary supplementation of ascorbic acid because they lack the enzyme L-gulonolactone oxidase necessary for synthesis of this compound. Although diets formulated for guinea pigs are supplemented with ascorbic acid, it is important to remember that it breaks down very rapidly, usually within the first 90 days after production. If a guinea pig is not receiving vitamin C supplementation in its diet, the veterinarian can assume that the diet is deficient in this important nutrient. Ascorbic acid is a necessary component of collagen, and deficiencies are often noted as manifestations of abnormal collagen synthesis.13 Clinical signs of hypovitaminosis C can include lameness, hemorrhage, lethargy, anorexia, poor coat quality, and bruxism. Diagnosis of hypovitaminosis C is based largely on clinical signs and history. Radiographs will reveal changes to the costochondral junctions and widening of the epiphyses of long bones. Recommended treatment includes fluids and nutritional support, pain control, and parenteral vitamin C supplementation. Attention should also be paid to improvement of the guinea pig’s diet at home. In addition to causing skeletal and cartilage abnormalities, vitamin C deficiency has been known to reduce immune function. In vitro, vitamin C deficiency was demonstrated to cause a reduction in migration of macrophages in guinea pigs.18

Urogenital DYSTOCIA Dystocia most frequently occurs in primiparous sows that are bred after approximately 6 months of age. At this time, the symphysis between the pubic bones becomes fused and will not expand to allow the passage of fetuses. Cesarean section must be performed in these cases to save the sow and the young. Factors other than age that can predispose a sow to dystocia include large fetuses in relation to sow size, uterine inertia, and obesity.13

URINARY CALCULI Urolithiasis occurs commonly in pet guinea pigs, and the common clinical signs associated with the disease include stranguria and pollakiuria, vocalizing when urinating, and hematuria. The underlying cause(s) of this condition is not completely understood but is likely associated with a genetic predisposition and/or the presence of a high-calcium diet. Other less common underlying etiologies associated with urinary calculi formation include ureteral neoplasms (e.g., papilloma).19 Calculi are primarily composed of calcium carbonate, although magnesium ammonium phosphate hexahydrate and calcium phosphate calculi will also occur.20 Radiographic or ultrasonic imaging can be used to confirm the location of the urinary calculi, which may be present in the renal pelvis, ureters, urinary bladder, or urethra (Figure 17-18). Urinary tract calculi often require surgical removal. Ideally, once

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REPRODUCTIVE NEOPLASMS

Figure 17-18 Ultrasound image of a calculus in the right renal pelvis of a guinea pig. (Courtesy University of California– Davis.)

Neoplasia of the reproductive tract is not commonly reported in guinea pigs, but several tumor types have been described. Of the described reproductive neoplasms, the vast majority occur in female guinea pigs. Uterine leiomyoma (often associated with ovarian cysts)22 and leiomyosarcoma, ovarian teratoma, and granulosa cell tumor are reported neoplasms of the reproductive tract.21 Diagnosis, as in any other patient, should be based on the results of cytologic or histopathologic sampling. Benign neoplasms may be resolved with ovariohysterectomy, but further diagnostics to determine the extent of local invasion should be performed before surgery takes place. A thorough diagnostic work-up, including imaging techniques (e.g., abdominal ultrasound, thoracic radiographs), should be performed to look for metastases of malignant tumor types.

MYCOPLASMA CAVIAE removed, the calculus and/or a portion of the bladder wall should be cultured. It may also be helpful to analyze the composition of the stone to help determine ways of preventing recurrence.

OVARIAN CYSTS Ovarian cysts are a common finding in middle-aged to older female guinea pigs. This disorder has a reported prevalence of 76% in female guinea pigs aged 1.5 to 5 years.13,21 Multiple cysts can be present on one or both ovaries. Clinical signs related to cystic ovaries can include abdominal distention, lethargy, and anorexia related to the space-occupying nature of the cysts. Ovarian cysts that are actively producing hormones can produce bilaterally symmetrical alopecia of the flanks. Ovarian cysts can be quite large (up to several centimeters) and can often be identified on physical examination. Abdominal ultrasound is the most useful diagnostic tool for a definitive diagnosis. The most definitive treatment for cystic ovaries is ovariectomy or ovariohysterectomy, curing the current problem and preventing recurrence. In animals where surgery is not a viable option, ultrasound-guided percutaneous aspiration of ovarian cysts can be performed as a palliative treatment. However, without removal of the ovaries, the cysts will recur, requiring repeated procedures.

PREGNANCY TOXEMIA Pregnancy toxemia typically occurs in pregnant sows during the last 2 weeks of gestation. As with other species, pregnancy toxemia is the result of a negative energy balance and the metabolism of fat. Sows that experience pregnancy toxemia are typically overweight and become anorexic. Clinical signs include lethargy, dyspnea, and anorexia, usually progressing to death within a few days.13 Therapy of affected sows should center on providing nutritional support, correcting electrolyte imbalances, and preventing opportunistic infections. Prognosis is generally considered poor, as many sows fail to respond to treatment.

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Mycoplasma caviae has been isolated from the reproductive tracts of guinea pigs.15 These guinea pigs are often unaffected by the organism, but metritis has been suspected to be associated with infection. As with other species, Mycoplasma spp. is suspected to cause reproductive problems (e.g., abortion and decreased fertility).

Respiratory PNEUMONIA Respiratory disease is a common presenting complaint in guinea pig patients. Clinical signs can vary from sneezing and upper respiratory signs to severe dyspnea and death. Bordetella bronchiseptica is one of the most common respiratory bacterial agents associated with pneumonia in guinea pigs.15 Many guinea pigs are carriers of the organism, which will cause clinical disease if the animal is stressed. A thorough history, obtained from the patient’s owner, often reveals interactions with other species that are also subclinical carriers of B. bronchiseptica (e.g., rabbits, dogs).13 Clinical signs noted in guinea pigs infected by the Bordetella organism include nasal discharge, dehydration, tachypnea, and lethargy.23 Diagnosis should be based on clinical and radiographic signs and history. Confirmation of the diagnosis can be determined with enzyme-linked immunosorbent assay (ELISA) and indirect immunofluorescence diagnostic tests, or culture of exudates.24 Treatment/preventive options for B. bronchiseptica infections in guinea pigs include an autogenous bacterin vaccine as well as three commercially available vaccines for Bordetella spp. (porcine B. bronchiseptica and human B. pertussis), which were found to offer some protection against the development of bronchopneumonia in experimentally infected guinea pigs.25

STREPTOCOCCUS PNEUMONIAE Streptococcus pneumoniae can cause pleuropneumonia, pleuritis, and peritonitis in guinea pigs.26 Serologic detection of S. pneumoniae antibodies using ELISA have been described in

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previously published papers.26 On postmortem examination, lesions found to be associated with S. pneumoniae infection included pleuritis, pleural effusion, lung abscessation, otitis media, pericarditis, and others.20 Streptococcus pneumoniae was also identified from septic arthritis lesions in a group of guinea pigs.26 Five guinea pigs in a laboratory colony demonstrated multiple enlarged joints from which pure cultures of S. pneumoniae were isolated. Several other individuals in this colony had previously died with typical S. pneumoniae lesions (e.g., pleuritis, pericarditis) and lesions consistent with hypovitaminosis C. The group with septic arthritis also had scorbutic changes on necropsy.26

STREPTOBACILLUS MONILIFORMIS Streptobacillus moniliformis, the causative agent of rat-bite fever in humans, was isolated from a laboratory guinea pig with pneumonia. This organism is of particular importance because of its zoonotic potential.27

Figure 17-19 A guinea pig patient with severe Trixacarus caviae infestation. (Courtesy Stephen D. White, DVM, DACVD.)

ADENOVIRUS Adenovirus has been shown to cause necrotizing bronchopneumonia in guinea pig populations, although it can also produce a transient, subclinical infection.28,29 Clinical signs include depression and dyspnea, but guinea pigs often die acutely without clinical signs.30 Identification of adenovirus DNA from diseased lung tissue was achieved using a polymerase chain reaction technique in one study.28 Development of this PCR assay for identification of the virus has determined that the guinea pig adenovirus is distinct.28 Histopathologic examination of affected tissues of animals infected with adenovirus will reveal the presence of characteristic intranuclear inclusion bodies.29

Figure 17-20 Microscopic view of an egg of the louse Gliricola porcelli adhered to a shaft of hair. (Courtesy Stephen D. White, DVM, DACVD.)

BRONCHOGENIC PAPILLARY ADENOMA The most commonly reported neoplasm of the respiratory tract in guinea pigs is bronchogenic papillary adenoma. The prevalence of the tumor is as high as 30% in guinea pigs over the age of 3 years.13 Given the relatively more common occurrence of pneumonia in guinea pigs, bronchogenic papillary adenoma can often be misdiagnosed. For this reason, thoracic radiographs of guinea pigs with respiratory disease are highly recommended.

YERSINIA PSEUDOTUBERCULOSIS Yersinia pseudotuberculosis can cause septicemic pneumonia in guinea pigs.15 Death usually occurs rapidly, after the development of coughing and dyspnea. At necropsy, severe congestion of the lungs is found grossly and through histologic evaluation of the tissues.

Integument ECTOPARASITES Trixacarus caviae are sarcoptoid mites that commonly affect guinea pigs (Figure 17-19). Affected guinea pigs are intensely

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pruritic, sometimes to the extent of seizure development. As with other mite infestations, diagnosis is based on the identification of the parasites on skin scrapings. Once definitive diagnosis has been made, treatment with ivermectin and selamectin (6 mg/kg q2-4wk) is usually effective. Chirodiscoides caviae are fur mites diagnosed in guinea pigs. Because it rarely causes clinical disease, treatment is usually unnecessary.20,31 Gyropus ovalis and Gliricola porcelli are species of lice that are commonly identified in guinea pigs (Figure 17-20). Infested guinea pigs may be pruritic, but are usually unaffected.20 Guinea pigs severely infested with these mites may demonstrate poor coat quality and alopecia.

DERMATOPHYTES Patchy hair loss without associated pruritus may be attributed to dermatophytosis, most commonly Trichophyton mentagrophytes2 (Figure 17-21). Lesions are circular and scaled and usually occur on the face and head.20 The diagnosis of dermatophytosis is made by a positive fungal culture. Because of the zoonotic potential of these fungal organisms, care should be

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Figure 17-21 A guinea pig with a crusted area on the dorsal pinna consistent with dermatophytosis. A Trichophyton organism was cultured from the site.

MANUAL OF EXOTIC PET PRACTICE

Figure 17-22 Severe, ulcerative pododermatitis in a guinea pig. (Courtesy Michelle G. Hawkins, VMD, DABVP [Avian].)

taken when handling guinea pig patients suspected of having dermatophytosis.

CERVICAL LYMPHADENITIS Streptococcus zooepidemicus Lancefield’s group C is the causative agent of cervical lymphadenitis. This disease will cause severe swellings of the lymph nodes in the cervical region in guinea pigs. Affected guinea pigs will frequently exhibit no other clinical signs but may become septicemic, with lesions affecting the heart, lungs, kidney, and skin.20 The most effective treatment for cervical lymphadenitis is complete surgical excision of the affected lymph nodes, followed by appropriate antibiotic therapy based on culture and sensitivity testing. Lancing and draining the abscesses is often not curative, as the abscesses form thick capsules that harbor organisms, leading to recurrence. Yersinia pseudotuberculosis has also been shown to cause cervical lymphadenitis in guinea pigs.17 Although the affected guinea pigs are usually not ill, the concern is that rupture of the abscesses will release large amounts of this potentially zoonotic organism into the environment. Another potential causal agent of cervical lymphadenitis is Streptobacillus moniliformis.32 One report exists of a cervical mass, initially believed to be cervical lymphadenitis, which was histologically determined to be a thyroid papillary adenoma.33 The mass was apparently nonfunctional and the guinea pig appeared otherwise healthy, but surgical resection was curative.

PODODERMATITIS Guinea pigs housed in cages with wire flooring are predisposed to developing ulcerated lesions on the plantar surfaces of their feet. Mild lesions may appear as hyperemic, swollen areas of the weight-bearing surfaces. These lesions can progress to ulcerations with secondary infections (Figure 17-22). Vitamin C deficiency has also been considered a predisposing factor for

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Figure 17-23 Trichofolliculoma on the dorsum of a guinea pig.

the development of pododermatitis, as affected animals may be in pain and reluctant to move, resulting in the development of pressure sores.20 Ulcerated, infected lesions should be managed with appropriate antibiotics and antiinflammatory drugs, but the focus of treatment should be improving husbandry (e.g., providing appropriate flooring/bedding, vitamin C supplementation).

TRICHOFOLLICULOMA Trichofolliculomas are the most common cutaneous tumor seen in guinea pigs.20 These benign tumors often occur on the dorsum and are typically round and hairless (Figure 17-23).

CUTANEOUS VASCULAR MALFORMATION Another cutaneous abnormality that has been described in guinea pigs is cutaneous vascular malformation.34 The lesion described was a raised, ulcerated plaque on the animal’s flank, which bled intermittently. Ultimately, the cutaneous vascular malformation resulted in fatal hemorrhage. Histologically, the

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lesion was described as an expansile mass extending into the skeletal muscle and consisting of multiple vascular spaces of varying sizes. These vascular spaces were lined with endothelial cells.

Neoplasia Compared to the incidence of neoplasia in other mammalian species, the incidence of neoplasia in guinea pigs appears low or is underreported. However, there have been several reported cases of neoplasia in guinea pigs. As more guinea pig owners seek quality veterinary care for their pets, reports of neoplasia will increase.

LYMPHOMA Lymphoma is the most commonly reported neoplasia in guinea pigs.20 Clinical signs associated with guinea pig neoplasia include lymphadenopathy, splenomegaly, and hepatomegaly. Leukemic and aleukemic forms of guinea pig lymphoma have been identified.

THYROID CARCINOMA Thyroid carcinoma has been reported in an adult guinea pig that demonstrated multiple masses in the ventral cervical region.35 As there was no evidence of neoplastic disease elsewhere on postmortem examination, this was considered a primary tumor.

MESOTHELIOMA Mesothelioma has been reported in the abdomen of an adult guinea pig.36 The guinea pig had died of complications associated with pneumonia, and the mesothelioma was diagnosed through a necropsy examination. The mass consisted of diffuse nodules on the serosal surfaces of numerous organs in the abdominal cavity.

Miscellaneous HEAT STROKE Guinea pigs are native to cooler regions of South America and are therefore relatively intolerant of temperatures above 80° F, lower if the environment is also humid. Guinea pigs should be housed in well-ventilated enclosures at temperatures between 65° and 75° F to prevent heat stress. Clinical signs of heat stress include rapid, shallow respirations, lethargy, poor peripheral perfusion, and ptyalism.13 Treatment includes reducing the animal’s core body temperature with cool water baths or applying alcohol to the feet and ears; in addition, fluid therapy (either intravenous or subcutaneous) is recommended to improve perfusion. The prognosis for this condition is guarded.

in adults as well.37 In one outbreak, the typical conjunctival abnormalities were present with other, more significant, clinical signs, such as rhinitis, abortion, and pneumonia.38 A definitive diagnosis can be achieved through Giemsa staining, immunofluorescent antibody testing of conjunctival scrapings, and serologic testing.37

PROLIFERATIVE UROCYSTICA AND ADENOMA Guinea pigs have been reported to develop pathologic changes related to ingestion of Forssk fern.39,40 In these studies, guinea pigs fed fresh fern developed hematuria and hemorrhage of the bladder wall.40 One guinea pig fed dried fern developed proliferative urocystica and adenoma of the bladder, a finding sometimes considered precancerous in human patients. This guinea pig did not show clinical signs associated with the bladder abnormalities.39

RABIES Rabies virus infection is uncommon in rodent species, but it has been described and should be considered a differential diagnosis for an ill guinea pig with suspect contact to wildlife, especially raccoons. A recent report of a rabies in a privately owned guinea pig described abnormal behavior (biting the owner) 26 days after the guinea pig had possible interactions with a raccoon.41 In this guinea pig, rabies virus antigen was detected by immunofluorescent antibody testing in the sublingual salivary gland, tongue, and buccal tissues, implying that the guinea pig could have transmitted the virus via a bite wound.

THERAPEUTICS Fluids Sick guinea pigs are often anorexic and therefore dehydrated. Restoration of normal hydration status is crucial for the successful treatment of many disease processes. Replacement of fluid deficit and maintenance of normal hydration can be achieved by administering crystalloids substances through subcutaneous, intraperitoneal, intravenous, or intraosseous routes. Because of the difficulty in placing and maintaining catheters in peripheral veins and bones, the most common route for fluid administration is subcutaneous. Subcutaneous fluid administration is generally well tolerated. Fluids are administered under the skin of the cranial, dorsal thorax (Figure 17-24). Butterfly catheter needles are useful because they allow the patient to move around without pulling out the injection needle. Maintenance fluid rates for guinea pigs are 80-100 mL/kg/day.

INCLUSION-BODY CONJUNCTIVITIS

Feeding

Chlamydophila psittaci has been identified as a disease-causing agent in guinea pigs; it usually causes a mild, self-limiting conjunctivitis. This intracellular bacterial disease usually occurs in young guinea pigs 4 to 8 weeks of age but has been reported

Another important aspect of management for the sick guinea pig is nutritional support. Anorexic guinea pigs can experience a change in their normal GI flora in as little as 8 to 12 hours. This change of GI flora can lead to ileus, colic, overgrowth of

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Figure 17-24 Subcutaneous fluid administration to a guinea pig using a butterfly catheter.

Figure 17-25 Mask induction of a guinea pig. A Doppler has been placed on the right forelimb to monitor heart rate.

pathogenic bacteria, and enterotoxemia. Commercial products are available that are palatable and high in fiber. These products help to maintain gut motility (Oxbow Critical Care for Herbivores, Oxbow Hay Company, Murdock, NE). Patients will often eat directly from a dish or 60-ml catheter tip syringe. For patients that are more resistant to eating, a technique that is useful, in my experience, is to remove the plungers from 1-ml or 3-ml syringes and fill them individually using a catheter tip syringe. Although this method may seem tedious, it allows the delivery of small boluses of food to be incrementally dispensed.

For lengthy or potentially painful procedures, preanesthetic medications may be used before induction of the inhalant anesthetic agent. As a preanesthetic, an injectable combination, midazolam (0.2-0.5 mg/kg) and butorphanol (0.20.5 mg/kg), has been used with success. The administration of premedications will also decrease the stress of induction. Isoflurane and sevoflurane are both acceptable inhalant anesthetics for guinea pigs. The advantage of sevoflurane is that it does not appear to have as noxious a scent as isoflurane, thereby reducing breath holding during induction and producing a smoother anesthetic episode. After induction, the inhalant anesthetic may be switched to isoflurane if cost is an issue. Guinea pigs should not be fasted more than 2 to 3 hours before anesthesia. Because these animals are hindgut fermenters, withholding food for longer periods of time may disrupt GI flora. Careful monitoring during anesthesia is essential. The number of respirations must be visually monitored for depth and character. Heart rate and rhythm should also be monitored continuously using a pediatric stethoscope or Doppler unit, which can be placed on a peripheral artery. Changes in heart rate or respiratory rate can occur rapidly, so it is advantageous to have precalculated doses of emergency drugs (e.g., glycopyrrolate, epinephrine, atropine, dopram) drawn and available for use before anesthetic induction.

Antibiotics The GI tract of guinea pigs can be very sensitive to the effects of certain classes of antibiotics. The GI flora of guinea pigs is primarily Gram positive, and administration of antibiotics with a primarily Gram-positive spectrum can result in overgrowth of Gram-negative and anaerobic organisms. Enteral administration of penicillins (e.g., amoxicillin, ampicillin), macrolides (e.g., erythromycin, lincomycin), and first-generation cephalosporins can result in overgrowth of opportunistic pathogens.20 Ampicillin administered subcutaneously at doses of 8 and 10mg/kg three times a day resulted in mortality rates of 20% and 30%, respectively, in a group of guinea pigs. At necropsy, Clostridium difficile was cultured from the ceca of all fatalities.42 Twenty-five percent of guinea pigs administered 100 mg/kg of cefazolin, a first-generation cephalosporin, intramuscularly died of enterocolitis after several injections.43 All antibiotics should be used with caution in guinea pigs because of the possibility of disruption of the GI flora (Table 17-2).

SURGERY Anesthesia Guinea pigs can be induced using isoflurane or sevoflurane administered by mask or induction chamber (Figure 17-25).

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Intubation Intubation of guinea pigs is generally considered to be very difficult. The long, narrow oral cavity makes visualization of the glottis difficult. The soft tissues of the tongue and soft palate are continuous in the caudal oropharynx, leaving only a small aperture, the palatal ostium. Intubation must be performed through this opening in the mucosal tissues. Several techniques have been described for visualizing the glottis to assist with intubation. These usually involve alterations of laryngoscope blades so that they may be introduced into

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TABLE 17-2

Selected Drug Dosages, Dosing Frequencies, and Routes of Administration

Drug

Dose(s)44-46

Dose frequency

0.1-1.0 mg/kg 0.05 mg/kg 0.4-2.0 mg/kg 1-4 mg/kg 0.5-3.0 mg/kg 0.01-0.02 mg/kg 10-44 mg/kg 20-30 mg/kg(K)/1-2 mg/kg(D) 5-10 mg/kg(K)/0.5-1.0 mg/kg(M) 1 mg/kg 1-2 mg/kg 1-2 mg/kg

— q8-12h q4-12h q12-24h — — — — — q12-24h — q3h

PO, SC, IM, IV PO PO, SC, IM PO PO, SC, IM

20-50 mg/kg 5-20 mg/kg 5-15 mg/kg 25 mg/kg 15-30 mg/kg

q6-12h q12h q12h q12h q12h

PO PO PO

5-25 mg/kg 5 mg/kg 10-40 mg/kg

q24h q24h q24h

PO SC Topical PO PO, SC, IM Topical PO

20 mg/kg 0.2-0.4 mg/kg — 25 mg/kg 5-10 mg/kg 6 mg/kg 25-50 mg/kg

q24h q7-14days q7days q12h q10-14days q28days q24h

PO, PO PO, IV PO, SC, PO, PO PO,

5-10 mg/kg 0.1-0.5 mg/kg 5-7.5 mg/kg 0.003 mg/kg 2-10 mg/kg 50-100 ml/kg 0.2-1.0 mg/kg 25-100 mg/kg 10-100 mg/kg

q6-12h q8-12h — — q12h q24h q12h q8-12h q24h

Route(s) of administration

Analgesic/anesthetic agents Atropine Buprenorphine Butorphanol Carprofen Diazepam Glycopyrrolate Ketamine Ketamine/diazepam Ketamine/midazolam Ketoprofen Midazolam Nalbuphine

IM SC, SC, PO, IM SC, IM IM IM SC, IM IM

IV IM SC IM

IM

Antibiotic agents Chloramphenicol Ciprofloxacin Enrofloxacin Metronidazole Trimethoprim-sulfa Antifungal agents Griseofulvin Itraconazole Ketoconazole Antiparasitic agents Fenbendazole Ivermectin Lime sulfur dip Metronidazole Praziquantel Selamectin Sulfadimethoxine Miscellaneous agents Cimetidine Cisapride Diphenhydramine Epinephrine Furosemide Lactated Ringer’s Metoclopramide Sucralfate Vitamin C

SC, IM, IV SC SC IV SC, IM SC, IM

IM, intramuscular; IV, intravenous; PO, per os; SC, subcutaneous.

the oral cavity without traumatizing the delicate buccal mucosa.47,48 Another possible intubation technique utilizes a stethoscope altered so that an appropriately sized endotracheal tube can be affixed to the end. The endotracheal tube is then inserted into the caudal oropharynx. The anesthetist will then listen through the ear pieces of the stethoscope for expiration.

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The tube is then gently introduced into the trachea. This technique requires practice to perfect but, once the procedure has been performed several times, is a reliable way to intubate guinea pigs. Application of a small amount of lidocaine to the rima glottis can ease intubation. Rigid endoscopes and otoscopes, when available, are also very helpful in visualizing the glottis for intubation.

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Ovariohysterectomy Indications for an ovariohysterectomy procedure in guinea pigs include routine sterilization, dystocia, and reproductive tract disease (e.g., neoplasia, muco/hydro/pyometra, ovarian cysts). In general, the surgical approach and technique are similar to those used with domestic species. The patient is placed in dorsal recumbency, and a ventral midline incision is made through the linea alba. Care must be taken upon entering the abdomen to avoid incising the large cecum, which is often situated along the ventral body wall. The body of the uterus can be located dorsal to the bladder. The ovaries lie caudally and laterally to the kidneys and are 6 to 8 mm in length.49 The abdomen of the guinea pig is deep, and the ovaries can be difficult to exteriorize. Care must be taken to avoid tearing the short ovarian vessels. Once the ovaries are removed, the uterus is ligated and transected in a manner similar to that used with other mammals.

Ovariectomy Another method of sterilization that is reported in guinea pigs is ovariectomy. The incidence of uterine disease compared with ovarian disease in guinea pigs is quite low. The approach for removal of the ovaries is through small incisions of the dorsal-lateral body wall caudal to the last rib. The advantage of this approach is that the sensitive GI tract is not manipulated to reach the structures to be excised.

Orchiectomy Orchiectomy is typically performed to prevent reproduction, to decrease undesirable sexual behavior, and to treat reproductive tract disease. The orchiectomy procedure in guinea pigs is performed by making an incision through the scrotum over each testicle. A closed technique or open technique, whereby the vaginal tunic is incised, may be performed. Although herniation of abdominal contents through the inguinal canal does not commonly occur with open castration, it is still advisable to close the inguinal ring if this technique is used. The incisions may be closed with either an intradermal suture pattern or with tissue glue.

Suture Reaction Reactions to suture material are relatively common in guinea pigs after surgical procedures. These reactions can vary in severity from mild local irritation to abscessation. Suture materials that cause a large amount of inflammation, such as chromic gut, should never be used in guinea pigs. In general, monofilament suture materials that are degraded by hydrolysis are preferred.

REFERENCES 1. Verstraete FJM: Advances in diagnosis and treatment of small exotic mammal dental disease, Semin Av Exot Pet Med 12(1):37-48, 2003.

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2. Flecknell PA: Guinea pigs. In Beynon PH, Cooper JE, editors: Manual of Exotic Pets, Cheltenham, Gloucestershire, 1991, British Small Animal Veterinary Association. 3. Harkness JE: Biology and husbandry: guinea pigs. In Harkness JE, editor: A Practitioner’s Guide to Domestic Rodents, Lakewood, Colo, 1993, American Animal Hospital Association. 4. Quesenberry KE, Donnelly TM, Hillyer EV: Biology, husbandry, and clinical techniques. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 5. Carpenter JW: Hematologic and serum biochemical values of rodents. In Carpenter JW, editor: Exotic Animal Formulary, ed 3, St Louis, 2005, WB Saunders. 6. Campbell TW: Mammalian hematology: laboratory animals and miscellaneous species. In Thrall MA, editor: Veterinary Hematology and Clinical Chemistry, Baltimore, 2004, Lippincott Williams and Wilkins. 7. Eremin O, Coombs RRA, Ashby J et al: Natural cytotoxicity in the guinea-pig: the natural killer (NK) cell activity of the Kurloff cell, Immunol 41:367-378, 1980. 8. Revell PA: The Kurloff cell, Int Rev Cytol 51:275-314, 1977. 9. Silverman S, Tell LA: Radiology equipment and positioning techniques. In Silverman S, Tell LA, editors: Radiology of Rodents, Rabbits, and Ferrets: An Atlas of Normal Anatomy and Positioning, St Louis, 2006, WB Saunders. 10. Gibson SV, Wagner JE: Cryptosporidiosis in guinea pigs: a retrospective study. JAVMA 189(9):1033-1034, 1986. 11. Wobeser G: Tyzzer’s disease. In Williams ES, Barker IK, editors: Infectious Diseases of Wild Mammals, ed 3, Ames, Iowa, 2001, Blackwell. 12. Motzel SL, Riley LK: Subclinical infection and transmission of Tyzzer’s disease in rats, Lab Anim Sci 42:439-443, 1992. 13. O’Rourke DP: Disease problems of guinea pigs. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 14. Jaax GP, Jaax NK, Petrali JP et al: Coronavirus-like virions associated with a wasting syndrome in guinea pigs, Lab Anim Sci 40(4):375-378, 1990. 15. Rigby C: Natural infections of guinea pigs, Lab Anim 10:119-142, 1976. 16. Haberman RT, Williams FP: Salmonellosis in laboratory animals, J Natl Cancer Inst 20: 933-947, 1958. 17. Paterson JS: The guinea pig or cavy, The UFAW Handbook on the Care and Management of Laboratory Animals, ed 4, Edinburgh and London, 1972, Churchill Livingstone. 18. Ganguly R, Durieux MF, Waldman RH: Macrophage function in vitamin c-deficient guinea pigs, Am J Clin Nutr 29:762-765, 1976. 19. Steiger SM, Wenker C, Zeigler-Gohm D et al: Ureterolithiasis and papilloma formation in the ureter of a guinea pig, Vet Radiol Ultrasound 44(3):326-329, 2003. 20. Quesenberry KE: Guinea pigs, Vet Clin North Am Small Anim Pract 24(1):67-86, 1994. 21. Bishop CR: Reproductive medicine of rabbits and rodents. In Speer BL, editor: The Veterinary Clinics of North America, Exotic Animal Practice: Reproductive Medicine 5:3, Philadelphia, 2002, WB Saunders. 22. Greenacre CB: Spontaneous tumors of small mammals. In Graham JE, editor: The Veterinary Clinics of North America, Exotic Animal Practice: Oncology 7:3, Philadelphia, 2004, WB Saunders. 23. Trahan CJ, Stephenson EH, Ezzell JW et al: Airborne-induced experimental Bordetella bronchiseptica pneumonia in strain 13 guinea pigs, Lab Anim 21:226-232, 1987. 24. Wullenweber M, Boot R: Interlaboratory comparison of enzyme-linked immunosorbent assay (ELISA) and indirect immunofluorescence (IIF) for detection of Bordetella bronchoseptica antibodies in guinea pigs, Lab Anim Sci 28:335-339, 1994. 25. Matherne CM, Steffen EK, Wagner JE: Efficacy of commercial vaccines for protecting guinea pigs against Bordetella bronchiseptica pneumonia, Lab Anim Sci 37(2):191-194, 1987. 26. Matsubara J, Kamiyama T, Miyoshi H et al: Serodiagnosis of Streptococcus pneumoniae infection in guinea pigs by an enzyme-linked immunosorbent assay, Lab Anim 22:304-308, 1988. 27. Kirchner BK, Lake SG, Wightman SR: Isolation of Streptobacillus moniliformis from a guinea pig with granulomatous pneumonia, Lab Anim Sci 42(5):519-521, 1992. 28. Pring-Åkerblom P, Blažek K, Schramlová J et al: Polymerase chain reaction for detection of guinea pig adenovirus, J Vet Diagn Invest 9:232-236, 1997.

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29. Butz N, Ossent P, Homberger FR: Pathogenesis of guinea pig adenovirus infection: Lab Anim Sci 49(6):600-604, 1999. 30. Harris IE, Portas BH: Adenoviral bronchopneumonia of guinea pigs, Aust Vet J 62(9):317, 1985. 31. Lumeij JT, Cremers HJWM: Anorexia and Chirodiscoides caviae infection in a guinea pig (Cavia porcellus), Vet Rec 119:432, 1986. 32. ILAR: A guide to infectious diseases in guinea pigs, gerbils, hamsters, and rabbits, part II, diseases outlines, Inst Lab Anim Res News 17:ID7-ID15, 1974. 33. LaRegina MC, Wightman SR: Thyroid papillary adenoma in a guinea pig with signs of cervical lymphadenitis, JAVMA 175(9):969-971, 1979. 34. Osofsky A, De Cock HEV, Tell LA et al: Cutaneous vascular malformation in a guinea pig (Cavia porcellus), Vet Dermatol 15:47-52, 2004. 35. Zarrin K: Thyroid carcinoma of a guinea pig: a case report, Lab Anim 8:145-148, 1974. 36. Wilson TM, Brigman G: Abdominal mesothelioma in an aged guinea pig, Lab Anim Sci 32(2):175-176, 1982. 37. Deeb BJ, DiGiacomo RF, Wang SP: Guinea pig inclusion conjunctivitis (GPIC) in a commercial colony, Lab Anim 23:103-106, 1989. 38. Schmeer VN, Weiss R, Reinacher M et al: Verlauf einer Chlamydienbedingten “Meerschweinchen-Einschluß Körperchen-Könjunktivitis” in einer Versuchstierhaltung, Zeitschrift für Versuchsteirkunde 27:233-240, 1985.

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473 39. Somvanshi R, Sharma VK: Proliferative urocystica and adenoma in a guinea pig, J Comp Pathol 133:277-280, 2005. 40. Somvanshi R, Sharma VK: Preliminary studies on Christella dentate (Forssk) fern toxicity in guinea pigs, J Lab Med 5:6-15, 2004. 41. Eidson M, Matthews SD, Willsey AL et al: Rabies virus infection in a pet guinea pig and seven pet rabbits, JAVMA 227(6):932-935, 2005. 42. Young JD, Hurst WJ, White WJ et al: An evaluation of ampicillin pharmacokinetics and toxicity in guinea pigs, Lab Anim Sci 37(5):652-656, 1987. 43. Fritz PE, Hurst WJ, White WJ et al: Pharmacokinetics of cefazolin in guinea pigs, Lab Anim Sci 37(5):646-651, 1987. 44. Ness RD: Rodents. In Carpenter JW, editor: Exotic Animal Formulary, ed 3, Philadelphia, 2005, WB Saunders. 45. Morrisey JK, Carpenter JW: Formulary. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 46. Plumb DC: Veterinary Drug Handbook, ed 4, Ames, Iowa, 2002, Iowa State Press. 47. Blouin A, Cormier Y: Endotracheal intubation in guinea pigs by direct laryngoscopy, Lab Anim Sci 37(2):244-245, 1987. 48. Turner MA, Thomas P, Sheridan DJ: An improved method for direct laryngeal intubation in the guinea pig, Lab Anim 26:25-28, 1992. 49. Breazile JE, Brown EM: Anatomy. In Wagner JE, Manning PJ, editors: Biology of the Guinea Pig, New York, 1976, Academic Press.

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Shannon M. Riggs Mark A. Mitchell

C H A P T E R

1 8

CHINCHILLAS COMMON SPECIES KEPT IN CAPTIVITY There are two species of chinchilla that are kept in captivity: Chinchilla laniger (or lanigera) and C. brevicaudata.1,2 C. laniger is the species commonly kept in captivity in the United States. Originally imported into the United States to provide a commercial fur, these long-lived rodents make excellent pets.

BIOLOGY Unique Anatomy and Physiology Chinchillas are native to the semiarid mountainous areas of South America, particularly the countries of Peru, Argentina, Bolivia, and Chile.1,2 Most of the chinchillas kept in the United States are descendants of a small group of animals imported into California in the 1920s.1 Because of overhunting for its beautiful coat, the chinchilla is believed to be nearly extinct in the wild. Wild-type chinchillas have a silver-gray coat with black ticking. Today, there are a number of different coat color variations that have been developed as a result of the fur, show, and pet chinchilla trades. Because the captive chinchillas found in the United States originated from a very small number of animals, a genetic component is suspected in many of the commonly seen disease processes. Adult chinchillas usually weigh between 400 and 800 g, females being slightly larger than males. Relative to other rodent species, the life span of the chinchilla is long, sometimes nearing 20 years of age.1 Chinchillas are naturally nocturnal, but they can adapt to a more diurnal lifestyle. Chinchillas are unique-looking animals with a short body, large head, delicate limbs, large hairless ears, and a bushy tail.

Chinchillas have the longest gestation period of any commonly kept rodent species (110 days). Young, of which there are one to six (average two), are precocial. Chinchillas become sexually mature at 7 to 9 months of age.

HUSBANDRY Creating an Appropriate Habitat ENCLOSURE SIZE Chinchillas are active animals, requiring a relatively large enclosure. The enclosure should be large enough to provide an area for eating, sleeping, exercising, and eliminating wastes (a latrine). Cages with multiple levels are preferred, as they provide additional room for the animals to exercise. Chinchilla cages are generally constructed of wire. Any other materials being considered for a cage should be “chew proof.” When considering a wire cage, it is important to select a structure that will not entrap limbs. The floor of the cage should be solid. If the bottom of the cage is also wire, a section of solid flooring should be provided to prevent pododermatitis. Chinchillas tend to be somewhat timid, so it is essential to provide ample space for hiding. Hide boxes should be constructed of materials that can be cleaned easily (e.g., polyvinyl chloride pipes) or can be disposed of (e.g., cardboard boxes). Wooden hide boxes are not recommended, as they are difficult to disinfect and are commonly chewed on by chinchillas.

TEMPERATURE/HUMIDITY Because of their dense hair coat, chinchillas are not tolerant of temperatures greater than 80° F (26.7° C). That said, chinchillas should be housed indoors throughout the summer. In hot weather, owners should be encouraged to operate the home’s air conditioner to maintain an appropriate temperature for

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their chinchilla. If this is not possible, alternatives might include using electric fans or placing plastic bottles filled with ice in the chinchilla’s enclosure.

LIGHTING Chinchillas are naturally a nocturnal animal, although they easily adapt to a diurnal lifestyle. These animals should be provided a 12-hour photoperiod (12 hours light, 12 hours darkness). Although there is no published evidence for this, these animals likely benefit from natural unfiltered sunlight. If they are housed indoors, full-spectrum lighting may be beneficial.

SUBSTRATE A layer of soft bedding should be provided on the floor of the cage to absorb waste and decrease pressure on the plantar surfaces of the feet. Recycled paper products, shredded newspaper, and aspen shavings are the substrates of choice. Cedar and pine shavings should be avoided, as the aromatic oils in these substrates can act as contact and respiratory irritants. The enclosure should be cleaned thoroughly on a regular basis, at least 2 times per week. Unsanitary conditions can predispose the chinchilla to pododermatitis, respiratory problems, and other health problems.

ACCESSORIES Shelter Box Chinchillas should be provided shelter areas within their enclosure. Non-treated wooden boxes may be used, but may eventually need to be replaced if the animal chews on the box.

Dust Baths To maintain proper coat health, chinchillas must be provided with a dust bath 2 to 3 times per week for 10 to 20 minutes (Figure 18-1). The dust used for these animals is a volcanic ash, and commercial brands are readily available. It is important to remove the dust bath from the chinchilla’s enclosure after the bathing period. If the dust bath is not removed, chinchillas tend to bathe excessively, and the prolonged exposure to dust can cause ocular and respiratory irritation.

NUTRITION

Figure 18-1 A chinchilla enjoying a dust bath. Dust baths are essential for chinchillas’ skin maintenance and coat health. (Courtesy Michelle G. Hawkins, VMD, DABVP [Avian].)

treats on an occasional basis, but they should comprise less than 5% of the animal’s diet. Chinchillas should be provided ad lib access to water in a hanging water bottle.

PREVENTIVE MEDICINE Quarantine Newly acquired animals should be quarantined from established animals for a minimum of 30 days. This recommendation should be considered a minimal quarantine period, as certain diseases could remain latent for more than 30 days. All quarantined animals should be thoroughly examined and screened using baseline diagnostics (e.g., complete blood count, plasma biochemistries, and a fecal exam). During the quarantine period, the animal’s appetite, demeanor, and fecal output should be monitored closely.

Diet

Routine Exams

Chinchillas are native to an area of the world that contains sparse vegetation, primarily consisting of grasses. The recommended nutritional composition of a chinchilla diet is 16% to 20% protein, 2% to 5% fat, and 15% to 35% bulk fiber.3 An appropriate chinchilla diet should be comprised of a highquality hay (e.g., timothy, oat, or orchard grass), chinchilla pellets, and an assortment of dark leafy vegetables (e.g., romaine lettuce, mustard greens, collard greens). Some veterinarians recommend rabbit or guinea pig pellets for chinchillas; however, these pellets are shorter in length than those recommended for chinchillas and are more difficult for these animals to hold. Fruits, grains, and raisins may also be provided as

Chinchillas are not routinely vaccinated for infectious diseases in the United States. However, owners should be encouraged to have an annual examination performed on their pets, including a thorough oral examination and blood work. Being a prey species, chinchillas are adept at hiding illness, and a routine evaluation by a qualified veterinarian may help in detecting abnormalities early. Many health problems of chinchillas are directly related to improper husbandry. During a routine veterinary visit for chinchilla patients, owners should be asked to provide a detailed description of the animal’s environment, including cage size and type, substrate, frequency of cleaning, ambient

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476 temperature, and exercise time. Information pertaining to the animal’s diet, feeding frequency, and bowel movements is also important to obtain.

Disinfection and Sanitation Chinchilla enclosures, food bowls, water bottles, and cage accessories should be sanitized using a standard disinfection protocol. Sodium hypochlorite is the preferred disinfectant (dilute 5.25% solution 1 : 32). It is important to thoroughly rinse the enclosure and accessories before replacing them in the cage. The cage should be allowed to dry in a well-ventilated area.

RESTRAINT Manual Restraint Chinchillas are relatively docile and usually do not require aggressive restraint; however, they are capable of quick bursts of activity. A chinchilla that is dropped may injure itself, so appropriate restraint is necessary when examining or transporting these animals. In general, the more a chinchilla is restrained, the more it tends to struggle, so a “less is more” approach should be used. A chinchilla should be restrained by supporting the weight of the body with one hand under the thorax and grasping the base of the tail with the other hand (Figure 18-2). Chinchillas should never be scruffed or handled roughly. Rough handling can result in “fur slip” or the loss of a patch of fur at the site where the animal was grasped. This lost fur can take several months to grow back (see Dermatologic Conditions).

Chemical Restraint Chinchillas that are fractious or must be anesthetized can be given either parenteral or inhalant anesthetics. We prefer inhal-

Figure 18-2 Restraining a chinchilla for transport. The handler has control of both the pelvic and thoracic limbs to prevent the patient from jumping and injuring itself.

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ants for these animals, although the parenterals can be used as preanesthetics to limit the concentration of inhalant required for a procedure. Chinchillas can be induced via a face mask with isoflurane or sevoflurane (Abbott Laboratories, North Chicago, IL). In general, an induction rate of 3% to 5% isoflurane (per 1 L oxygen) or 5% to 7% sevoflurane (per 1 L oxygen) is required. Chinchillas, like other rodents, are difficult to intubate because of their long, narrow oral cavity. We prefer to intubate these animals using an endoscope. A rigid 2.7-mm endoscope can be used to visualize the airway and direct the endotracheal tube into place. The endotracheal tube size required for a chinchilla can vary, although 2.0 to 4.0 outside diameter tubes are generally acceptable. Once the chinchilla is intubated, the endotracheal tube should be secured and the animal maintained on the inhalant. In general, chinchillas can be maintained at 1.5% to 3.0% isoflurane (per 0.5-1 L oxygen) or 3.0% to 5.0% sevoflurane (per 1 L oxygen). Many of the parenteral anesthetics used for domestic pets can also be used for chinchillas. (For a complete list of parenteral agents please refer to Anesthesia; see also Table 18-6).

PERFORMING A PHYSICAL EXAMINATION A physical examination should include both a hands-off and a hands-on review of the animal. For the hands-off exam, particular attention should be paid to the respiratory rate and character, attitude, and posture. Animals with respiratory compromise or neurologic disease should be handled carefully during the hands-on examination. It is important to always measure the animal’s body temperature before performing the complete physical exam. A chinchilla’s body temperature is generally lower (96° to 99° F [35.5° to 37.2° C]) than that found in other domestic mammals. Because chinchillas have such a dense hair coat, their body temperature can increase dramatically over the course of a physical examination, and this should be taken into consideration. Otherwise, the physical exam should be approached in a thorough, systematic manner, as in any mammalian patient. Thoracic auscultation and abdominal palpation can be performed as in other patients. Auscultation of gut sounds is also an important part of the chinchilla physical exam. A healthy chinchilla should have 1 to 2 borborygmi per minute. A complete oral examination is another important facet of the chinchilla physical exam. This should be reserved for the end of the examination, as it can be rather stressful for the patient. As is the case for most rodent species, the cranial surface of the incisors is covered with a layer of yellow to orange enamel (Figure 18-3). The oral cavity of the chinchilla has a small opening and is very narrow and long. In most cases, an otoscope with cone or a human nasal speculum is required to fully examine the molars and caudal aspect of the oral cavity. In most cases, the oral exam can be performed on an alert animal; however, anesthesia may be required when the animal is difficult to restrain, appears to be experiencing pain, or requires a more detailed oral exam.

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BOX 18-1

Hematologic Parameters for Chinchillas

Hematologic Parameter PCV (%) RBC (×106 cells/μl) Hemoglobin (g/dl) WBC (×103 cells/μl) Heterophils (%) Lymphocytes (%) Monocytes (%) Eosinophils (%) Basophils (%) Platelets (×103 cells/μl)

Figure 18-3 Rostrocaudal view of a chinchilla skull. Note the normal orange coloration of the enamel on the rostral surface of the incisors.

Common Abnormalities Found on the Physical Examination As with any species, it is important to always perform a thorough physical examination to minimize the likelihood of misclassifying the disease state of a patient. Therefore, veterinarians working with an animal should avoid the trap of only examining the system(s) that a client mentions during the history and instead examine the entire animal. There are a number of potential abnormalities that may arise during a physical examination. In many cases, an animal may present with multiple, unrelated problems. The primary problems found on a chinchilla examination include teeth malocclusion, otitis externa, heat stroke, upper and lower respiratory disease (e.g., pneumonia), penile fur ring, pododermatitis, and fur slip.

DIAGNOSTIC TESTING Hematology Venipuncture in chinchillas can be difficult for the veterinarian and stressful for the patient. Many of the peripheral vessels that are accessible in dogs and cats are very small in chinchillas and do not allow for the collection of adequate volumes of blood for routine testing. Access to larger vessels, such as the jugular vein, requires somewhat aggressive restraint, which may not be appropriate in a sick or stressed animal. It is often preferable to perform venipuncture under sedation or anesthesia in these patients; however, veterinarians must carefully consider the benefits of obtaining diagnostic samples against the risk of anesthesia for each individual patient. When a large blood sample is required (e.g., for complete blood count and plasma biochemistry panel), the jugular, cranial vena cava, or femoral vein can be used. These sites are best accessed under anesthesia, as restraint is stressful and excessive struggling by the patient may cause laceration of these large vessels. A 25-gauge needle fastened to a 1-ml or 3-ml

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Reference Interval 30-55 5-10 9-15 6-16 25-50 10-70 0-5 0-5 0-2 300-600

Data from Quesenberry KE, Donnelly TM, Hillyer EV: Biology, husbandry, and clinical techniques of guinea pigs and chinchillas. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders; de Oliveria Silva T, Kruetz LC, Barcellos LJG et al: Reference values for chinchilla (Chinchilla laniger) blood cells and serum biochemical parameters, Ciência Rural, 35(3):602-606, 2005. PCV, packed cell volume; RBC, red blood cell count; WBC, white blood cell count.

syringe can be used to collect the sample. If small quantities of blood are required (e.g., blood glucose or PCV/TS [packed cell volume and total solids determination]), blood can be collected from the cephalic or lateral saphenous veins. When sampling these smaller veins, a 25- or 27-gauge needle fastened to a 1-ml syringe can be used. Insulin syringes may also be used. Using a smaller syringe will help minimize the likelihood of collapsing the blood vessel. When placing the sample into a blood storage tube, it is important to remove the needle before discharging the sample to prevent hemolysis. Hematologic testing is an important diagnostic tool in chinchilla medicine. Because these prey species can mask their illness, veterinarians must rely on hematologic and other diagnostic tests to fully assess their patients. Box 18-1 presents reference intervals for chinchillas. The most common leukocyte in the chinchilla is the lymphocyte. The second most common circulating granulocyte is the heterophil. Heterophils lack myeloperoxidase, the enzyme responsible for liquefying purulent material. Therefore, chinchilla abscesses tend to be very thick and caseous. The remaining leukocyte types—monocytes, eosinophils, and basophils—are normally present in very low numbers. During an inflammatory response in a chinchilla, the early stages are often characterized by a shift in the differential (e.g., increased heterophils, decreased lymphocytes) rather than an increase in absolute leukocyte count, so evaluation of the entire leukogram is essential. The platelet count can also serve as an important marker of inflammation in chinchillas. Large increases in the platelet count (>1,000,000/μl) may be seen without an increase in the total white blood cell count, indicating an inflammatory process. Plasma biochemistries can provide important information regarding the physiologic status of a chinchilla. As with any patient, it is important to interpret the results of a chemistry

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MANUAL OF EXOTIC PET PRACTICE

Plasma Biochemical Parameters for Chinchillas

Biochemical Parameter Sodium (mEq/L) Potassium (mEq/L) Chloride (mEq/L) Glucose (mg/dl) Blood urea nitrogen (mg/dl) Creatinine (mg/dl) Calcium (mg/dl) Phosphorous (mg/dl) Total protein (g/dl) Albumin (g/dl) Globulin (g/dl) Creatine kinase (IU/L) Aspartate transferase (IU/L) Alkaline phosphatase (IU/L) Bilirubin (mg/dl) Cholesterol (mg/dl)

Reference Interval 130-170 3-7 110-130 80-125 10-40 0.8-2.3 8-15 4-8 5-8 2.5-4.0 3.5-4.2 0-300 15-100 10-70 0.6-1.3 40-300

Figure 18-4 A right lateral radiograph of a chinchilla. Note the small size of the thorax and the large size of the abdominal cavity. (Courtesy University of California–Davis.)

Data from Quesenberry KE, Donnelly TM, Hillyer EV: Biology, husbandry, and clinical techniques of guinea pigs and chinchillas. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders; de Oliveria Silva T, Kruetz LC, Barcellos LJG et al: Reference values for chinchilla (Chinchilla laniger) blood cells and serum biochemical parameters, Ciência Rural, 35(3):602-606, 2005.

panel in conjunction with the anamnesis, physical exam findings, and the results of other diagnostic tests. Reference intervals for various chinchilla biochemical parameters can be found in Box 18-2.

Urinalysis A urinalysis is an important diagnostic test in chinchilla medicine and should be considered in any case with signs of upper or lower urinary tract disease. Urine samples may be obtained by free catch, floor catch, or cystocentesis. If urine is to be cultured, it should be collected by cystocentesis. The cystocentesis procedure for chinchillas is similar to that described for cats. The animal should be placed in dorsal recumbency for the procedure. Sedation may or may not be necessary. A 25gauge needle fastened to a 3-ml syringe can be used to collect the sample. The ventral abdomen should be disinfected before needle insertion using standard techniques. Ultrasound can also be used to guide the needle into the bladder. The urine of chinchillas is typically yellow to amber in color, although the color may be darker and more orange in color depending on the diet. Pigments (e.g., porphyrins) in the urine can sometimes be mistaken for hematuria and must be ruled out by the absence or presence of blood cells on a direct smear. Because chinchillas are herbivores, the pH of chinchilla urine should be alkaline (pH 8.0-9.0).

Diagnostic Imaging Whole body radiographs can provide a significant amount of information regarding the status of a chinchilla patient. A

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Figure 18-5 A ventrodorsal radiographic view of a chinchilla. Again, note the small size of the thorax and large size of the abdomen in this species. (Courtesy University of California– Davis.)

minimum of two radiographic views should be taken. Our preferences are for lateral and ventrodorsal (or dorsoventral) views (Figures 18-4 and 18-5). Care should be taken to extend the limbs when positioning the patient, to minimize rotation and the superimposition of the limbs over the abdomen or thorax. Chinchillas typically resent aggressive restraint, so sedation or anesthesia is helpful in obtaining diagnostic

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Figure 18-6 Right lateral radiographic view of a chinchilla skull. Note the severe elongation of the crowns and apicies of the premolars and molars as well as the abnormal occlusal angle of the incisors. The large tympanic bullae of the chinchilla are also easily seen (arrow). (Courtesy University of California– Davis.) Figure 18-7 Computed tomography image of a chinchilla skull in saggital section. There is mild malocclusion of the cheek teeth visible (arrow). (Courtesy University of California–Davis.)

TABLE 18-1

Guidelines for Radiographic Techniques for Selected Radiographic Studies in Chinchillas

Anatomic location

mA

kVp

Whole body Extremities Skull

5.0 6.0 6.0

44 54-56 48-52

Data from Silverman S, Tell LA: Radiology equipment and positioning techniques. In Silverman S, Tell LA, editors: Radiology of Rodents, Rabbits, and Ferrets: An Atlas of Normal Anatomy and Positioning, St Louis, 2005, WB Saunders. kVp, kilovolt peak; mA, milliampere.

radiographs as well as in reducing the stress on the patient. Table 18-1 is intended as a general guideline for the techniques used for chinchilla radiographic studies. Dental malocclusion is a common problem in chinchillas. Skull radiographs can be used to fully assess the degree of malocclusion and secondary bony involvement (Figure 18-6). When taking skull radiographs, it may be necessary to take 3 to 4 different views. In addition to the standard lateral and dorsoventral (or ventrodorsal) views, right and lateral oblique views can be used to more specifically localize lesions. Magnified views of the skull can be obtained by placing the patient on an elevated platform under the x-ray beam without changing the distance between the x-ray cassette and the beam. Ultrasound is another imaging modality that can be used to diagnose various disease processes in chinchillas. As with radiographs, sedation or anesthesia may be required to reduce patient stress, minimize movement, and improve the overall quality of images that are acquired. The approach to the chinchilla ultrasound exam is similar to that described for domestic species, although there are some exceptions. Chinchillas have a large cecum. If the cecum contains a large quantity of gas, it

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may limit the value of an abdominal ultrasound exam. Ultrasound can also be used to assist with the collection of various aspirates (e.g., cystocentesis) or biopsies (e.g., liver biopsy). Advanced imaging techniques, such as computed tomography (CT) and magnetic resonance imaging (MRI), can also be used to assist in the diagnosis of disease in chinchillas. CT imaging is particularly useful in diagnosing dental problems, such as apical abscesses, osteomyelitis, and minor malocclusions not readily seen on oral examination or radiographs (Figure 18-7). The primary limitations associated with these imaging modalities include limited availability to the general practitioner, the expense associated with the collection and interpretation of the image, and reduced image quality in comparison with that of larger patients. MRI has the added limitation of requiring a significant amount of time (often >45 min) to collect the image. Regardless of the present limitations associated with these advanced imaging modalities, veterinarians should be made aware that these techniques are available and that reference material exists that depicts normal anatomy for comparison.5 As chinchilla owners continue to demand high-quality care for their pets, these imaging techniques will likely become more commonplace in exotic small mammal practice.

Microbiology Chinchillas are susceptible to many of the same pathogens that affect domestic species. The primary pathogens isolated from chinchillas include Gram-negative bacilli (e.g., Bordetella spp., E. coli, Klebsiella spp., Salmonella spp., Pseudomonas spp.) and Gram-positive cocci (Staphylococcus aureus, Streptococcus spp.). Many of these organisms are opportunists and can be routinely

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480 isolated from healthy animals. Samples being submitted for bacterial culture should be collected using sterile swabs and submitted to a diagnostic laboratory capable of isolating and characterizing a range of organisms. The majority of the samples being submitted from chinchillas are for aerobic isolates and require no special attention regarding collection or submission. However, if an anaerobic infection is suspected, it is important to contact the local laboratory to obtain the appropriate materials to collect and submit the samples. A positive bacterial culture does not necessarily confirm that an organism is responsible for a disease, so it is important to always use the data obtained from the culture, in combination with information obtained from the clinical examination (e.g., elevated body temperature) and additional diagnostic tests (e.g., elevated complete blood count, biopsy, etc.), to confirm a disease. This is especially important in chinchillas, as they are susceptible to antibiotic-induced enteritis and should only be treated with (appropriate) antibiotics when a bacterial infection is highly suspected. Fungal infections are not common in chinchillas. However, if a fungal infection is suspected (e.g., no response to antibiotics, biopsy results), then standard techniques may be used to isolate the fungus.

Parasitology Chinchillas may become infested with a variety of endoparasites and ectoparasites. Fortunately, because these animals are bred in captivity, parasites are uncommon. Protozoa (e.g., coccidian) are the most commonly encountered parasites in captive chinchillas. Fecal flotation and/or direct fecal smears mixed with saline can be used to diagnose these infections. These same techniques can be used to diagnose other endoparasites, such as nematodes. Diagnosing ectoparasites in these animals can be done using the same techniques used in domestic animals. Fleas may be identified by closely inspecting the animal hair coat and combing through the fur. Most mites can be identified by using tape preparations or skin scrapes. All of these samples, once collected, should be evaluated under light microscopy.

COMMON DISEASE PRESENTATIONS Infectious BACTERIAL Enterotoxemia One of the most difficult presentations to manage in chinchilla medicine is enterotoxemia. In many of the cases, the animals are presented in severe distress. Lethargy, lateral recumbency, diarrhea, respiratory distress, and fever are all common findings in end-stage disease, whereas in other cases there may be no clinical course of disease and the patients die acutely. Clostridium perfringens is commonly associated with this disease.6,7 In one described outbreak, chinchillas between the ages of 2 and 4 months were most commonly affected.6 Most of the

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affected chinchillas died without showing clinical signs, but some showed signs of colic or diarrhea. This disease reinforces the importance of maintaining the microflora of the chinchilla intestine. The intestinal microflora of the chinchilla is a dynamic mixture of bacteria, protozoa, nematodes, and fungi. In clinically healthy animals, the microflora establish a balance. Alterations to this balance (e.g., administration of inappropriate antibiotics, stress, dietary changes) can result in the overgrowth of certain pathogenic organisms. A number of the bacteria of the microflora produce toxins as a natural defense mechanism against other bacteria and a host’s immune response. When these toxins are produced in excess, as is seen with the overgrowth of certain microbes, the host can suffer the consequences. Diagnosing enterotoxemia can be difficult and is generally a postmortem diagnosis. Serial antemortem fecal/rectal cultures may provide insight as to the causative agent. Hematologic results are often consistent with a toxic inflammatory response. Radiographs frequently reveal significant gas within the intestinal tract and an ileus. Treating these patients is difficult. Supportive care, including intravenous fluids and caloric support, should be instituted. Nonsteroidal antiinflammatories may provide some protection against the toxins. If a positive culture is made, an antibiotic sensitivity profile can be performed to determine the most appropriate antibiotic to use for treatment. Because treating enterotoxemia is difficult, it is best to focus on prevention. Judicious use of antibiotics by veterinarians and breeders, as well as providing a balanced, consistent diet, washing all fresh food products thoroughly, practicing strict sanitary practices, and minimizing environmental stress, are practices that can reduce the likelihood of enterotoxemia in a patient.

Enteritis Enteritis is a common disease in chinchillas and can occur from numerous underlying causes, including diet change or inappropriate diet, improper antibiotic use, stress, and various infectious organisms (e.g., coccidia, Giardia spp., Cryptosporidium spp., E. coli, Clostridium spp., Proteus spp., Pseudomonas spp., Staphylococcus spp., and Salmonella spp.).2,8,9 Clinical signs associated with enteritis are variable but can include abdominal pain, diarrhea, decreased fecal output, intussusception, intestinal impaction, and rectal prolapse.8,10 Treatment of enteritis should focus on identifying the underlying cause and treating the presenting signs. Further diagnostics that may be necessary to identify an underlying cause are radiographs, abdominal ultrasound, fecal examinations for parasites and bacteria, and fecal cultures. If a condition such as intestinal intussusception or impaction occurs and requires surgery, the patient should be stabilized first. Nutritional supplementation, fluid replenishment, pain management, and appropriate antibiotic therapy (ideally based on fecal culture and sensitivity) are facets of stabilization in these patients.

Salmonella Salmonella sp. is a Gram-negative rod from the family Enterobacteriaceae. All members of this genus are considered pathogenic and have been associated with disease in all of the

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different classes of vertebrates. In chinchillas, infections with this bacterium have been associated with acute death. In an epizootic in Australia involving five chinchilla farms, both Salmonella Enteritidis and Salmonella Sofia were isolated at necropsy.11 Affected animals were found to have gastritis, enteritis, splenomegaly, and hepatitis. Salmonella sp. was not isolated from any of the fecal samples collected from the affected herds following a course of treatment with enrofloxacin. Isolates were found to be resistant to sulfamethoxazole, but this was attributed to the fact that this drug was routinely added to the chinchillas’ feed. Salmonella sp. may be occasionally isolated from clinically healthy chinchillas. We do not recommend treating unaffected animals, as the risk of antibiotic resistance increases with exposure to antibiotics. In cases where a Salmonella sp. is isolated from a clinically diseased animal, the antibiotic selected for treatment should be based on the antibiotic sensitivity profile. If this profile is pending, fluoroquinolones are an excellent first choice antibiotic. Many of the Salmonella spp. isolated from chinchillas can be directly attributed to human environments. Because of this, it is important that clients thoroughly wash the fresh foods that they provide their pets and always practice strict hygiene when cleaning their pet’s environment.

tations are uncommon compared with other mammalian species. Dermatophytosis, however, is not uncommon. Trichophyton mentagrophytes is the most commonly isolated dermatophyte in chinchillas, although Microsporum canis and M. gypseum infections have also been reported.2,8,9 Dermatophyte lesions can occur on any surface of the body but are most often found on the head and face. The lesions associated with these parasites are similar to those described for domestic species and are typically scaly and alopecic. Diagnosis is based on the results of a dermatophyte culture. Treatment with oral antifungal medications and or topical antifungals is required to resolve the lesions.8,9 Eliminating the infections can require several weeks of treatment. Because of the zoonotic potential associated with these infections, owners should be educated about handling their chinchillas during the treatment period. We recommend that clients wear latex or nonlatex exam gloves when treating their pets and that children not handle the chinchilla during the treatment period. Clients should also be directed to contact their personal physician if they develop dermatologic conditions consistent with a dermatophyte infection.

Listeriosis

ENDOPARASITES

Chinchillas appear to be highly susceptible to infections caused by Listeria monocytogenes. Ranched chinchillas, which are often fed silage as a portion of their diet, are most commonly affected. This increased risk is primarily attributed to the fact that L. monocytogenes thrives in decaying organic matter. Signs of infection can originate from the gastrointestinal or central nervous systems. Clinical signs are usually vague but may include depression, anorexia, weight loss, and diarrhea.2 In one outbreak, 47 animals from seven ranches were diagnosed with listeriosis. Contaminated food was the suspected source. The course of disease in these animals was short, with the animals primarily displaying gastrointestinal disease and succumbing within 24 hours.12 L. ivanovii has also been described in a single chinchilla.13 This chinchilla died shortly after purchase. At necropsy, multifocal areas of necrosis and supparative inflammation were found in the liver. L. ivanovii was isolated from these lesions. These cases reaffirm the importance of providing these animals high-quality forage. Clients should request only fresh grass hays for their chinchillas. Grass hay with signs of mold or necrosis should be discarded. Diagnosing listeriosis using antemortem tests can be difficult. Hematology and radiographs are generally suggestive of an inflammatory response of an infectious nature and a corresponding enteritis. Cultures can be attempted, but veterinarians should contact the laboratory to ensure that the samples are collected and processed appropriately. Treatment for this disease is generally based on the provision of supportive care.

Several parasitic diseases have been reported as causes of gastrointestinal disease in chinchillas; however, most of the reports have been in farm-ranched chinchillas. The incidence of parasitism in pet chinchillas is relatively low and is most likely due to the fact that these animals are housed indoors. Giardia spp. are often found in low numbers in normal chinchillas.15 These parasites can cause problems when, combined with housing stress and poor hygiene, they develop into active infections with severe diarrhea and even death.16 One report of Cryptosporidium spp. infection has been reported in a young chinchilla.17 This chinchilla had a severe diarrhea that was not responsive to supportive therapy and ultimately succumbed to the infection. Diagnosing endoparasites of the intestinal tract can be done using standard techniques (see Diagnostic Testing: Parasitology).

FUNGAL Dermatophytes Because of the extremely dense nature of the chinchilla’s hair coat (up to 90,000 fibers per square inch),14 ectoparasite infes-

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Larval Migrans Baylisascaris procyonis, the raccoon roundworm, has been associated with larval migrans in a number of different vertebrates. The raccoon is the definitive host of this parasite. In nonraccoon hosts, this parasite has a predilection for ophthalmic and neural tissue. Once an animal is infected, the prognosis for the animal is guarded to grave. This parasite has been associated with disease in a group of farmed chinchillas.18 Affected animals were found to have clinical signs associated with neurologic disease, including ataxia, incoordination, and paralysis. Malacic tracts consistent with parasitic migration were identified in the brains of five animals presented for necropsy, and Baylisascaris procyonis larvae were identified in two of these animals. The chinchillas were exposed to the parasite after being fed hay that had been contaminated with raccoon feces. Although not described in pet chinchillas, B. procyonis

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should be included on any differential list for chinchillas displaying neurologic signs, especially those given access to the outdoors where they could potentially come in contact with raccoon droppings.

ECTOPARASITES Chinchillas are susceptible to various ectoparasites, including fleas, mites, and ticks. Ectoparasites are not common in pet animals, because they are generally housed in limited numbers and housed indoors. When chinchillas present with a history of crusting and scaling that is consistent with ectoparasitism, it is important for the veterinarian to pursue the case using standard diagnostics. Skin scrapes and tape preparations are the most common initial tests. The majority of ectoparasites found on chinchillas, excluding fleas, are responsive to ivermectin (0.2-0.4 mg/kg). The course of treatment will depend on the life cycle of the parasite. In most cases, 2 to 3 treatments every 14 days will eliminate most ectoparasites. During the ivermectin treatment, it is also important that the owner clean the animal’s enclosure daily to remove any adult organisms or eggs that fall into the substrate.

Neoplasia Neoplastic disease processes are relatively uncommon, or at least uncommonly described in the literature, in chinchillas. However, several tumor types have been documented as occurring in chinchillas, including neuroblastoma, carcinoma, lipoma, and hemangiosarcoma. Unfortunately, because many of these diagnoses are associated with farmed chinchillas from the 1950s, there are no detailed pathologic descriptions or even descriptions of the organ(s) involved.14 The Animal Medical Center in New York conducted a 5-year retrospective study of chinchillas and found only one neoplasm, a uterine lyomyosarcoma, among all of the chinchillas examined. The tumor was an incidental finding at necropsy.14 One case of malignant lymphoma in a chinchilla is described in the literature.19 This chinchilla displayed marked lymphadenopathy and evidence of metastases to the liver, spleen, and kidneys. Even though descriptions of neoplastic diseases are not well reported in chinchillas, as owners begin seeking more routine care for their pets and as appropriate care results in longer lives, veterinarians caring for these animals are more likely to encounter such geriatric diseases as neoplasia.

Miscellaneous GASTROINTESTINAL To reinforce the importance of the gastrointestinal tract to the inner workings of a chinchilla, one of us (Mitchell) often states that the “chinchilla is basically a bowel wrapped in fur.” Although this may seem rather simplistic, it is the gastrointestinal tract that serves as the major site of complications for these animals. Veterinarians working with these animals should always reinforce to their clients the importance of maintaining the function of the gastrointestinal tract to ensure the good health of their pet.

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M P

I

Figure 18-8 Open view of a chinchilla skull. (I, incisor; P, premolar; M, molar.)

DENTAL DISEASE The dentition of the chinchilla is aradicular hypsodont, meaning all teeth are continuously growing and open-rooted.20 This is an important concept, as any damage to the teeth or inherent (e.g., genetic) changes to the growth of the teeth can result in malocclusion. The dental formula of the chinchilla is similar to that found in the guinea pig: I 1/1, C 0/0, P 1/1, M 3/3 (Figure 18-8). As in other rodent species but differing from lagomorphs, the mandible is wider than the maxilla and has an occlusal angle that is lowest at the lingual surface. The normal occlusal angle of the chinchilla is very slight, only 5 to 10 degrees from flat. The enamel on the cranial surface of the incisors of chinchillas is normally orange in color and should not be mistaken for a pathologic condition (see Figure 18-3). The clinical signs associated with dental disease in chinchillas are similar to those seen in guinea pigs and rabbits, including decreased appetite or dysphagia, weight loss, ptyalism, decreased fecal output, and poor coat quality.21-24 Abscessation of the molar and premolar apices can also occur with the overgrowth of these teeth. Ocular and/or nasal discharge is another common clinical sign seen in chinchilla patients with dental disease. Because the premolars and molars are elodont (open-rooted), it is possible that the apical surfaces of these teeth can overgrow and impinge on the nasolacrimal duct, causing ocular discharge. The apicies can also overgrow into the nasal cavity, which could result in the seeding of the sinuses with bacteria from the oral cavity. Diagnosing dental disease in chinchillas requires a thorough evaluation of the animal’s history and close inspection of the animal’s oral cavity. Animals with a genetic history of malocclusion, or that are maintained on an inappropriate diet, should be examined regularly to prevent severe dental disease. Careful palpation of the ventral mandible and maxilla may reveal bony protuberances that correspond to the overgrowth of the apical surfaces of the cheek teeth. To thoroughly examine the oral cavity, it is important to have the appropriate

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Figure 18-9 Severe incisor malocclusion (arrow) in a chinchilla.

Figure 18-10 Oral examination of an anesthetized chinchilla, demonstrating a large buccal point on a left maxillary premolar (arrow). Note the use of incisor dilators and pouch dilators to improve visualization of the oral cavity.

equipment. Dental specula and pouch dilators are invaluable aids in obtaining an adequate view of the molars and premolars; however, many abnormalities can be overlooked in a conscious animal, and anesthesia is often required to obtain a thorough oral examination. Abnormal oral examination findings that are commonly seen in chinchillas may include an uneven occlusal surface or angle of the incisors and/or cheek teeth (Figure 18-9), formation of sharp points (Figure 18-10) with or without associated ulceration of the oral mucosa, food impaction, and abnormal spaces (diastema) between teeth. Imaging modalities, including whole body radiographs, magnified skull radiographs, and CT, can be incorporated to better evaluate the extent and seriousness of the process. Radiographic studies should include dorsoventral, lateral (Figure 18-11), and right and left oblique views.

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483

Figure 18-11 Lateral view of a chinchilla skull showing severe malocclusion of the cheek teeth (arrow), with overgrowth of the apices of the molars and premolars. (Courtesy University of California–Davis.)

Treatment of dental malocclusion in chinchillas centers on restoring a normal occlusal plane to the teeth. This is a procedure that must be performed under general anesthesia to allow adequate visualization of the oral cavity and to minimize stress to the patient. Patients that are dehydrated have altered gastrointestinal motility, or other potentially complicating symptoms and should be provided with supportive care before undergoing the anesthetic event. Utilizing the right tools is important to ensure the best outcome of treatment. A high-speed surgical dental handpiece can be used to restore a normal occlusal plane and to reduce any sharp points that have formed because of improper wear. Handheld trimmers, such as canine nail trimmers, are not acceptable, as they tend to crush teeth and can cause fractures and pulp exposure.23 Dremel tools are not acceptable either because the small size of the chinchilla’s oral cavity will not allow for appropriate trimming. With any dental instruments, care should be observed to minimize oral soft tissue trauma. Dental malocclusion is a disease that can be managed but rarely cured. The owners of chinchillas diagnosed with malocclusion must be made aware that this will be a lifelong problem for their pets; it will require repeated anesthetic procedures, which could result in significant cost. With many cases, routine occlusal adjustments will be necessary for the remainder of the animal’s life. One of the most significant differences seen with malocclusions in chinchillas—as compared with other rodent species and rabbits—is that it is not infrequent for a chinchilla’s oral exam to be normal while significant disease is present in the portions of the tooth that are not visible. For this reason, imaging, such as skull radiographs or CT, is especially useful in these patients to determine the extent of the disease as well as the presence of apical abscesses or osteomyelitis, which may complicate treatment. Chinchillas normally have very short

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clinical crowns, limiting the amount of correction that may be performed during occlusal adjustments. Because dental disease appears to have a heritable component, animals with dental disease should not be allowed to breed.21,22 A dietary component is also considered likely.23,25 Owners of chinchillas with malocclusion should be advised to increase the amount of roughage in the animal’s diet to increase the grinding motion of the teeth and increase wear.

decreased appetite. This condition can be resolved by removing the fur ring. Sedation or anesthesia may be required for those animals that cannot be restrained manually. If paraphimosis has occurred, a hypertonic solution (e.g., 50% dextrose) can be applied to the penis to reduce the edema and facilitate replacement of the organ. A sterile, water-based lubricant can also be used to aid in replacing the penis into the prepuce.

CHOKE/BLOAT

DERMATOLOGIC CONDITIONS Fur Slip

Like other rodent species, chinchillas do not have the ability to vomit. Because of this, small pieces of food can become lodged in the esophagus. When this occurs, it is generally referred to as “choke.” The most common food items that become lodged in the esophagus are small treat items (e.g., raisins and nuts) that are not thoroughly chewed. Choke can also occur in chinchillas with dental malocclusion and often results from the animal’s inability to chew their food properly. The clinical signs most commonly observed in a chinchilla with choke include retching, ptyalism, pawing at the mouth, anorexia, and respiratory distress.9 If left untreated, a chinchilla with choke will die. Diagnosing choke in chinchillas should follow the same standard protocol as for any gastrointestinal foreign body. Hematologic testing should be done to evaluate the patient’s physiologic status. Radiographs should be taken to determine the location of the foreign body. In many cases, the foreign material can be retrieved using an endoscope. Supportive care, including fluids and caloric support, should be provided to the patient to ensure that the gastrointestinal tract continues to function. However, caloric support should not be instituted until after the foreign material is removed or after a feeding tube has been placed that is distal to the foreign body. Bloat, often a common sequela to choke, occurs because the obstruction of the esophagus does not allow the animal to eructate and release the gas that accumulates in the stomach. Animals with bloat are often depressed, painful on palpation, and lethargic. A diagnosis of gastric tympany can be confirmed via radiographs. Removing the foreign material from the esophagus will resolve the bloat. In an emergency situation, the gas within the stomach can be relieved via aspiration through a needle. The prognosis for chinchillas with bloat is guarded to grave.

UROGENITAL Fur Ring Fur ring is a generic name used to describe a common condition in male chinchillas, in which the fur surrounding the base of the penis can accumulate and form a restrictive ring. This is often a subclinical problem, but it can progress into causing paraphimosis or inflammation of the penis. To prevent this from occurring, clients should be shown how to routinely examine the prepuce for the accumulation of fur and how to extract the penis from the prepuce to inspect it for signs of inflammation. Clinical signs associated with fur ring can include stranguria, excessive grooming of the prepucial area, lethargy, and

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Because chinchillas have evolved as a prey species, it was important for them to develop protective measures against predators. As it is common for predators to grab or bite the skin of a prey species when they attempt to capture them, it would seem logical that to have a mechanism to prevent this from occurring would increase the likelihood of survival for a species. Chinchillas have developed an adaptive process that allows for a clump of fur that is grabbed roughly to easily epilate. This is known as fur slip, and can be problematic in both ranched and pet populations. Large areas of fur can be easily epilated when a chinchilla is handled roughly or is fighting with another chinchilla. While the underlying skin is usually not damaged, regrowth of the missing fur can take up to 4 to 6 months.2 For ranched animals this is associated with a significant loss of money. In captive pet animals, this can lead to a decrease in aesthetics. Fortunately, fur slip can be avoided in captive situations by teaching support staff and clients how to properly handle a chinchilla. In cases where this is associated with combat between male chinchillas, it is important to separate animals to prevent contact.

Fur Chewing (Barbering) Chinchillas occasionally develop a tendency to overgroom or barber their own fur or the fur of a cagemate. This can result in an animal having an unkempt or uneven hair coat quality. The underlying cause for this negative behavior is not known, but there are a number of suspected causes, including boredom or housing stress, poor diet (e.g., low fiber), or hereditary factors.1,2 Histopathologic examination and testing of the adrenal and thyroid glands of fur-chewing chinchillas have demonstrated increased hormone activity in these glands that is consistent with stress-induced changes.26,27 Histopathologic changes consistent with Cushing’s disease (e.g., cutaneous calcinosis, comedone formation) have also been described in furchewing chinchillas, suggesting that stress and genetics may both play roles in the development of this problem.27

MUSCULOSKELETAL Fractures of the long bones of the pelvic and thoracic limbs are relatively common in chinchillas. Improper caging, such as wire flooring with spaces large enough for a limb to become entrapped, and improper handling are common causes. Chinchillas are likely more prone to fractures of the distal limbs because of the relatively long, delicate construction of the limbs in comparison with the stocky body (Figures 18-12 and 18-13).

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Figure 18-12 A ventrodorsal radiograph of the pelvic limbs of a chinchilla, demonstrating the delicate structure of the long bones, particularly the tibia. (Courtesy University of California– Davis.)

Figure 18-13 A ventrodorsal radiograph of the right forelimb of a chinchilla. (Courtesy University of California–Davis.)

The cortices of the long bones of chinchillas are rather thin and brittle and are covered with little soft tissue.14,28 For these reasons, fractures of the limbs are often comminuted and open, complicating repair. External coaptation (Figure 18-14) is typically not adequate for stabilization of these fractures, and surgical repair is often necessary. Because these bones are so thin and fragile, most orthopedic equipment available for use in domestic animals is too large, and the veterinarian must be creative in developing fixators using such components as small K-wires and hypodermic or spinal needles (Figure 18-15). Chinchillas are rather active animals, and strict cage rest is extremely important for a positive outcome of any fracture repair. Unfortunately, despite the type of fracture fixation, malunion fractures are common in chinchillas. Degree of commi-

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Figure 18-14 A lateral radiograph of the forelimb of a chinchilla demonstrating the use of external coaptation for stabilization of mid-diaphyseal fractures of the radius and ulna. (Courtesy University of California–Davis.)

Figure 8-15 Application of a Type-II external skeletal fixator using hypodermic needles as cross bars. (Courtesy Paul M. Gibbons, DVM, MS, DABVP [Avian].)

nution, contamination of the fracture site, and poor soft tissue coverage are all contributing factors. Because malunion occurs frequently, owners should be made aware of this possibility from the beginning of treatment. Fortunately, chinchillas typically adapt well to the loss of a limb, so amputation does not affect them negatively in the long run.

HEART DISEASE Heart murmurs are relatively common physical exam findings in chinchillas. Anecdotal reports suggest cardiomyopathy as a possible underlying cause.14 One report exists of a heart murmur in a young, male chinchilla that was found to be caused by a ventricular septal defect and tricuspid valve regurgitation on echocardiography,29 but cardiomyopathy has not

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486 been reported as a significant cause of cardiac disease in chinchillas. Chinchilla patients with cardiac murmurs should be treated as any domestic pet with a heart murmur. Thoracic radiographs, electrocardiography, and echocardiography may be necessary to achieve a diagnosis. As more chinchilla owners begin seeking a higher level of veterinary care for their pets, cardiac disease may become more commonly reported and better described.

LEAD TOXICOSIS There have been a few cases of lead toxicosis in chinchillas reported in the literature. Blood concentrations of lead higher than 15μg/dl are considered suspicious for lead toxicosis, and levels higher than 25μg/dl are considered definitive.14 Chinchillas affected with lead toxicosis display primarily neurologic signs, including seizures and blindness.8,30 These cases were successfully treated with calcium disodium edentate (25 mg/kg SC q6h × 5 days,30 30 mg/kg SC q12h × 5 days8). Interestingly, in the reported cases of lead toxicosis in chinchillas, the affected animals were housed in older apartments in urban neighborhoods. Because of the tendency of rodents to gnaw on different surfaces, lead toxicosis should be included on any differential list for animals with neurologic disease and exposure to lead-based paints.

HEATSTROKE Chinchillas are native to the cool mountainous regions of South America and are therefore rather intolerant to temperatures higher than 80° F.10,14,31 Ideally, chinchillas should be housed in well-ventilated enclosures at temperatures between 65° F and 75° F. Animals that are exposed to excess heat can develop life-threatening heatstroke. Clinical signs in affected animals include panting, lethargy, poor peripheral perfusion, and ptyalism.10 Treatment includes reducing the animal’s core body temperature with cool water baths or the application of alcohol to the feet and ears and fluid therapy (either intravenously or subcutaneously) to improve perfusion. It is important to cool the animal gradually, as a rapid decrease in body temperature can induce a life-threatening hypothermia. Nonsteroidal antiinflammatories and antibiotics are often recommended to minimize endotoxemia and overgrowth of opportunistic pathogens, respectively. The prognosis for this condition is guarded to grave.

DIABETES MELLITUS Diabetes mellitus has been reported in a single obese female chinchilla.32 Clinical signs in this chinchilla were similar to those seen in domestic animals and included polydipsia, polyuria, weight loss, lethargy, and bilateral cataracts.22,32 This chinchilla had a blood glucose of higher than 400 mg/dl, glucosuria, and ketonuria. Treatment with insulin, 2 to 12 IU daily, did not succeed in controlling the hyperglycemia, and the chinchilla was eventually euthanized. Histopathologically, the pancreas was found to have marked vacuolation of the islet cells, consistent with diabetes mellitus.

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THERAPEUTICS Fluids Chinchillas, like other rodents, are highly dependent on the function of their gastrointestinal tract to provide essential calories in the form of volatile fatty acids. Any changes to the function of the intestinal tract can be catastrophic to the animal. Unfortunately, when these animals become dehydrated, it is often the gastrointestinal tract that suffers. The shift of the fluid balance to maintain the function of the brain and heart often comes at the expense of the gastrointestinal tract. As the shift in the fluid balance occurs, gastrointestinal motility diminishes, microflora changes occur, and the animals reduce their dietary intake and production of calories. If left untreated, these patients die. To minimize the likelihood of this occurring, it is essential that these patients receive parenteral fluids. Replacement of a fluid deficit, and the maintenance of normal hydration, can be achieved by administering crystalloid substances via the subcutaneous, intravenous, intraperitoneal, or intraosseous routes. In cases of severe dehydration, the intravenous and intraosseous routes are preferred. For mild dehydration, or when the animal cannot be catheterized, the oral, subcutaneous, or intraperitoneal routes can be used. In those cases where the gastrointestinal tract is functioning normally, and the patient is only mildly dehydrated, replacement fluids can be delivered per os. The simplest method of delivering fluids per os is via a syringe. Some chinchillas will tolerate this, whereas other will not. Mixing the water with food or using fruit-flavored electrolyte solutions may increase success. Chinchillas can also be provided per os fluids via a feeding tube; however, this is often difficult to do in these animals because of their narrow, long oral cavity. In those cases where a chinchilla will not accept fluids via a syringe, we rely on the other techniques of providing fluids (e.g., subcutaneous fluids). Subcutaneous fluid administration is generally well tolerated by chinchillas. Fluids can be administered into the subcutaneous space of the dorsal or lateral thorax, similar to that described for domestic species. When grasping the skin to introduce the needle, care should be taken to avoid causing fur slip. Butterfly catheters can be used to deliver fluids via the subcutaneous space; these catheters allow the patient to move around without pulling out the injection needle. The quantity of fluids that can be administered in a single space should be based on the size of the animal and elasticity of the skin. Fluids should be administered only to a point where the skin is mildly stretched, not taut. In many cases, veterinarians should use their best judgement when determining a maximum volume to deliver. Veterinarians should remember that they can deliver the fluids into multiple subcutaneous sites or increase the frequency of fluid administration over the course of a day to diminish the volume that needs to be delivered at a single point in time. The primary disadvantage associated with the delivery of drugs via the subcutaneous route is that fluid absorption is not as rapid as with the other routes. Another disadvantage is that there are certain types of fluids that should not be deliv-

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ered via this route, such as 5% or higher dextrose. This type of fluid can be irritating to the subcutaneous space. However, we have delivered lower concentrations of dextrose (2.5%) via this route without concern. The intraperitoneal cavity is an underutilized route of fluid administration in chinchillas. We prefer this route for moderately to severely dehydrated juvenile animals and adult animals that have collapsed peripheral veins. Fluids administered via the intraperitoneal route are presumed to be absorbed via the serosal surfaces of the viscera and the peritoneal membrane. Because of the large surface area of the peritoneal membrane and visceral surfaces, larger boluses of fluids can be provided via this route. However, care should be taken to avoid inducing an ascites. This is best prevented by monitoring the hydration status of the animal closely during treatment and routinely inspecting the size (direct visualization) and fluid status (ultrasonography) of the abdomen. The chinchilla should be placed into dorsal recumbency for the procedure, as this will allow gravity to displace the organs away from the injection site. The injection site should be disinfected using standard protocols. The needle should be inserted at a 20- to 30-degree angle off the abdominal wall. A sharper angle will increase the likelihood of inserting the needle into the viscera. Veterinarians should always aspirate the syringe before delivering the fluids to ensure that the fluids are not going to be delivered into an organ. The fluids being inserted into the abdomen should always be prepared at body temperature, as the provision of cool fluids into the abdomen can be painful to the animal and require additional energy from the animal to maintain its core temperature. When chinchillas are severely dehydrated and the peripheral vessels are collapsed, fluids can be administered via the intraosseous route. The proximal femur and tibia are the preferred sites for catheter placement. We suggest clipping the fur surrounding the proximal femur to increase the likelihood of visualizing the insertion landmarks. The injection site should be disinfected using standard sterile techniques to limit the likelihood of inducing an osteomyelitis. For a catheter being placed into the proximal femur, the space between the neck of the femur and the greater trochanter should be used as the

TABLE 18-2

insertion landmarks. For catheters being placed into the proximal tibia, the tibial crest is the insertion landmark. A spinal needle with a stylet should be used for the procedure to limit the likelihood of coring the needle. We secure the catheters into place using butterfly tape and suture, and we generally leave the catheters in place for 72 to 96 hours. Intravenous catheters remain the preferred route of delivering fluids in severely dehydrated chinchillas. The primary vessels used for intravenous fluids are the cephalic, lateral and medial saphenous, and jugular veins. To ensure the patency of the smaller peripheral vessels, we tend to use only the jugular veins for venipuncture. The peripheral vessels in these animals tend to be very moveable under the skin in comparison with those in other domestic mammals. We find that the likelihood of inserting an intravenous catheter increases significantly when the animal is sedated. Once a catheter is placed, it should be secured using standard bandaging techniques. In some cases, a distasteful substance may need to be placed on the bandage to prevent the chinchilla from extracting the catheter. An Elizabethan collar constructed out of exposed radiograph film may also be used to prevent a chinchilla from tampering with its catheter.

Antimicrobial Therapy Because many of the diseases diagnosed in chinchillas have a bacterial origin, the use of antibiotics in the captive management of this species is common. It is important to remember, however, that the indigenous microflora of these animals is predominantly Gram positive and that certain antibiotics can cause severe changes (antibiotic induced dysbiosis) in the bacterial population of the chinchilla gastrointestinal tract. Enteral administration of penicillins (e.g., amoxicillin, ampicillin), macrolides (e.g., erythromycin, lincomycin), and firstgeneration cephalosporins are most commonly associated with these changes.8 A list of common antibiotics used in these animals can be found in Table 18-2. As fungal disease can also occur in these animals, in some cases as a direct result of the misuse of antibiotics, it is important to be familiar with dosages for these compounds too (see Table 18-2).

Common Antimicrobials Used in Chinchilla Medicine

Drug

Route(s) of administration

Dosage(s)33-35

Dose frequency

Chloramphenicol Ciprofloxacin Enrofloxacin Metronidazole* Trimethoprim-sulfa

PO, SC, IM, IV PO PO, SC, IM PO PO, SC, IM

30-50 mg/kg 5-20 mg/kg 5-15 mg/kg 10-20 mg/kg 15-30 mg/kg

q12h q12h q12h q12h q12h

PO PO PO

25 mg/kg 10-40 mg/kg 5 mg/kg

q24h q24h q24h

Anti-fungal agents Griseofulvin Ketoconazole Itraconazole

*Use with caution. IM, intramuscular; IV, intravenous; PO, per os; SC, subcutaneous.

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Antiparasitic Drugs Commonly Used in Chinchilla Medicine

Drug

Route(s) of administration

Dosage(s)33-35

Dose frequency

Fenbendazole Ivermectin Lime-sulfur dip Metronidazole* Praziquantel Sulfadimethoxine

PO SC Topical PO PO, SC, IM PO

20 mg/kg 0.2-0.4 mg/kg — 10-30 mg/kg 5-10 mg/kg 25-50 mg/kg

q24h q7-14days q7days q12h q10-14days q24h

*Use with caution. IM, intramuscular; PO, per os; SC, subcutaneous.

TABLE 18-4

Common Emergency Drugs Used in Chinchilla Medicine

Drug

Route(s) of administration

Dose(s)33-35

Dose frequency

Epinephrine Dopram Furosemide Atropine

IV, IT, IO, IC IV, IT, IO, IC PO, SC, IM, IV (low dose) IM, IT, IC

0.2 mg/kg 20 mg/kg 2-10 mg/kg 0.1-0.2 mg/kg

prn prn q12h prn

IC, intracardiac; IO, intraosseous; IT, intratracheal; IV, intravenous, prn, as needed; SC, subcutaneous.

Antiparasitic Therapy Chinchillas that present with parasites should be treated promptly to limit the spread of disease between conspecifics. In a previous section of this chapter we reviewed the most common parasites identified in captive chinchillas. Table 18-3 represents a list of the compounds that can be used to treat these parasites.

Emergency Drugs Because of their ability to mask illness, chinchillas routinely present to veterinary clinics in emergency situations. It is also not uncommon for these animals to become emergencies during their stay in a hospital (e.g., enterotoxemia) or as a result of an anesthetic event. Although the prognosis for these cases is guarded to grave, attempts to provide emergency care should be made. A list of common emergency drugs used in chinchilla medicine can be found in Table 18-4.

Nutritional Support Nutritional support should be considered for any chinchilla that is not eating. Anorexic chinchillas can experience a significant change in their gastrointestinal microflora in as little as 8 to 12 hours. This change in the microflora can lead to ileus, colic, overgrowth of pathogenic bacteria, and enterotoxemia. To minimize the likelihood of the gastrointestinal tract from slowing or stopping, it is important to provide these animals calories. Oxbow Critical Care for herbivores (Oxbow Hay Company, Murdock, NE) is a commercial product that we have used with good success. This enteral provides essential

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Figure 18-16 Syringe feeding a chinchilla using 1-ml syringes.

calories and contains bacteria that can be used to refaunate the gastrointestinal microflora. It is important to follow the manufacturer’s feeding recommendations to ensure that the desired quantity of calories is provided. Patients will often eat the enteral directly from a dish or can be force-fed via a 60-ml catheter tip syringe. For patients that are more resistant to eating, a technique that we have used is to remove the plungers from 1-ml or 3-ml syringes and fill them individually using a catheter tip syringe (Figure 18-16). Although this method may seem tedious, it allows the delivery of small boluses of food to be incrementally dispensed.

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TABLE 18-5

Common Therapeutic Agents Used to Treat Chinchillas

Drug

Route(s) of administration

Dosage(s)33-35

Dose frequency

Cimetidine Cisapride Diphenhydramine Furosemide Lactated Ringer’s Metoclopramide Sucralfate

PO, PO PO, PO, SC, PO, PO

5-10 mg/kg 0.1-0.5 mg/kg 1-2 mg/kg 2-10 mg/kg 50-100 ml/kg 0.2-1.0 mg/kg 25-100 mg/kg

q6-12h q8-12h q12h q12h q24h q12h q8-12h

SC, IM, IV SC SC IV SC, IM

IM, intramuscular; IV, intravenous; PO, per os; SC, subcutaneous.

Miscellaneous Drugs Chinchillas, like domestic pets, can present to a veterinary hospital for a variety of different disease conditions. Because of this, it is important to consider treating these animals with the same compounds used to manage domestic pets. Unfortunately, the drugs used in small animal medicine are untested in chinchillas. To limit liability, it is important for the veterinarian to explain and document to their clients the limitations associated with the use of any therapeutic when treating their chinchilla patients. In our opinion, however, it is better to treat and accept a smaller risk than to not treat and accept certain patient loss. A list of common therapeutics used, at least anecdotally, in chinchilla medicine can be found in Table 18-5.

SURGERY Preoperative and Postoperative Considerations CATHETERIZATION As with venipuncture, catheterization for fluid or drug administration in chinchillas can be a challenge. Peripheral vessels typically used for catheterization in dogs and cats, such as the cephalic and lateral saphenous veins, are very small in chinchillas. Because of their small size, 24- to 26-gauge intravenous catheters are considered the most useful in these patients (Figure 18-17). However, small-gauge catheters tend to bend when piercing the skin. Catheter failure can be prevented by making a small puncture in the skin using a 22-gauge needle before placing the catheter. Care should be taken to avoid lacerating the vein before introducing the catheter.36 In some cases, sedating the chinchilla may reduce the stress and ease catheter of placement. Intraosseous catheters can be used as an alternative to intravenous catheters. The most common sites for intraosseous catheterization in the chinchilla are the proximal femur and the proximal tibia.36 A spinal needle with a stylet is preferred. Because intraosseous catheterization is a painful procedure, patients should be placed under general anesthesia and topical anesthetic (e.g., lidocaine) used to reduce the pain at the insertion site. If the patient is debilitated and general anesthesia is a risk, then the local is generally sufficient to place the catheter. The catheter insertion site should be disinfected using standard

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Figure 18-17 Placement of a 26-gauge intravenous catheter into the cephalic vein of a chinchilla.

aseptic techniques to minimize the risk associated with introducing pathogens. After placement, the catheter should be secured with a butterfly (tape) placed around the catheter hub and then sutured to the skin.

INTUBATION Intubating chinchillas is generally considered a difficult procedure. The long, narrow oral cavity of these animals makes the visualization of the glottis difficult. As in dogs and cats, laryngoscopes may be used to visualize the glottis, but extreme care should be taken to avoid damaging the soft tissues of the oral cavity or even fracturing the mandible. Even with the use of a laryngoscope, direct visualization of the glottis is very difficult. Another technique that may be used for intubating chinchillas is to affix an endotracheal tube to a stethoscope and place the tube via auditory cues. To place the tube, the anesthetist should listen through the ear pieces of the stethoscope for when the animal expires, and then gently insert the tube into the trachea. This technique requires practice to perfect, but once the procedure has been performed several times, can be a reliable method for intubating chinchillas. Application of a small amount of lidocaine to the rima glottis can also ease intubation by decreasing laryngospasm. One of the most productive methods of intubating chinchillas is to use a rigid

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endoscope. We have found that a 2.7-mm rigid endoscope can be used to visualize both the glottis and the endotracheal tube as it is passed into the airway.

Anesthesia Anesthesia should be provided to any chinchilla undergoing a painful or stressful procedure. Inhalant anesthetics are by far the most common way of anesthetizing these patients. Chinchillas can be induced using isoflurane or sevoflurane administered by face mask or induction chamber (Figure 18-18). Induction is generally done at 3% to 5% isoflurane (per 1 L oxygen) or 5% to 7% sevoflurane (per 1 L oxygen). Once anesthetized, patients can be maintained at 1% to 3% isoflurane and 2% to 4% sevoflurane, respectively. Sevoflurane is generally better tolerated by chinchillas, because they tend to object less to its odor than to the odor of isoflurane. This also reduces the amount of breath-holding that occurs during induction, which expedites induction and reduces the potential for complications. Chinchillas should not be fasted more

than 2 to 3 hours before anesthesia. Disruption of intestinal microflora can occur if food is withheld for longer periods of time, potentially resulting in negative sequelae such as enterotoxemia. Water does not need to be withdrawn before an anesthetic event. Preanesthetic agents can be used to reduce the anxiety associated with a procedure and decrease the amount of inhalant required for induction and maintenance of anesthesia. A combination of intramuscular midazolam (0.2-0.5 mg/kg) and butorphanol (0.2-0.5 mg/kg) has been used with good success. A list of other anesthetic and analgesic agents can be found in Table 18-6. Careful monitoring of the patient during anesthesia is essential. Respirations should be visually monitored for depth and character. Heart rate and rhythm should also be monitored continuously using a pediatric stethoscope or Doppler unit, placing the transducer on a peripheral artery (Figure 18-19). Changes in heart rate or respiratory rate can occur rapidly, so it is advantageous to have pre-calculated doses of emergency drugs (e.g., glycopyrrolate, epinephrine, atropine, doxopram) drawn and available for use prior to anesthetic induction. Monitoring a chinchilla’s body temperature is also very important during an anesthetic procedure. In chinchillas, as in other small animals, hypothermia often occurs when anesthetized animals are not provided thermal support. Supplemental heat should always be provided through water-circulating heating pads, heat lamps, or forced air blankets. Body temperature should be monitored closely to ensure that the patient is not overheated, which could lead to hyperthermia and death if not corrected.

Common Surgical Procedures

Figure 18-18 Inducing a chinchilla using a gas anesthetic agent.

TABLE 18-6

Chinchillas may present for a variety of potential surgical procedures, including fractures, trichobezoar/foreign body, and cystic calculi. The majority of these procedures can be performed using the same basic practices described for domestic mammals. The only primary difference between chinchilla surgery and that for domestic pets is that chinchilla tissues are

Common Analgesic and Anesthetic Agents Used in Chinchilla Medicine

Drug

Route(s) of administration

Dosage(s)33-35

Dose frequency

Atropine Buprenorphine Butorphanol Carprofen Glycopyrrolate Ketamine Ketamine/diazepam Ketamine/midazolam Ketoprofen Midazolam

IM SC, SC, SC SC, IM IM IM SC, IM

0.1-0.2 mg/kg 0.05 mg/kg 0.2-2.0 mg/kg 4 mg/kg 0.01-0.02 mg/kg 20-40 mg/kg 20-40 mg/kg(K)/1-2 mg/kg(D) 5-10 mg/kg(K)/0.5-1.0 mg/kg(M) 1 mg/kg 1-2 mg/kg

— q8-12h q4h q24h — — — — q12-24h —

IV IM IM

IM

IM, intramuscular; IV, intravenous; SC, subcutaneous.

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open technique, whereby the vaginal tunic is incised, may be performed. Although herniation of abdominal contents through the inguinal canal does not commonly occur with open castration, it is still advisable to close the inguinal ring if this technique is used. The incisions may be closed with either an intradermal suture pattern or with tissue glue.

ZOONOSES

Figure 18-19 An anesthetized chinchilla. Oxygen and anesthetic gas are being provided via a face mask. An intravenous catheter is being used to deliver fluids via the cephalic vein. The heart rate is being monitored via a Doppler unit (arrow).

generally more fragile. Therefore, when working with chinchillas, it is important to take extra care when manipulating the tissues (e.g., intestines, bones, bladder).

Reproductive Surgery OVARIOHYSTERECTOMY The indications for ovariohysterectomy in chinchillas are similar to those in domestic species, and include routine sterilization, dystocia, and reproductive tract disease (e.g., neoplasia, muco/hydro/pyometra). The surgical approach and techniques for this procedure in chinchillas are also similar to that of domestic species. The patient should be placed in dorsal recumbency and a ventral midline incision made through the linea alba. Care must be taken upon entering the abdomen to avoid incising the large cecum, which is often situated along the ventral body wall. Using forceps to lift the linea alba when incising the body wall can help reduce the risk of complication. The body of the uterus can be located dorsal to the urinary bladder. Manual expression of the urinary bladder before beginning the surgery will increase the visualization of the uterine body. The ovaries lie caudally and laterally to the kidneys and are 6 to 8 mm in length.37 The ovarian vessels are short, which does not allow for exteriorization of the ovaries for ligation of vessels.38 Care must be taken to avoid tearing these vessels when removing the ovaries. Once the ovaries are removed, the uterus is ligated and transected in a manner similar to that used with other mammals.

ORCHIECTOMY An orchiectomy is typically performed to prevent reproduction, decrease undesirable sexual behavior, and reduce the likelihood for reproductive tract disease. The orchiectomy procedure in chinchillas is performed by making an incision through the scrotum over each testicle. A closed technique or

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Many of the opportunistic bacterial pathogens (e.g., E. coli, Salmonella spp.), fungi (e.g., dermatophytes), and protozoal infections (e.g., Giardia spp., Cryptosporidium parvum) found in chinchillas can be infective to humans. To minimize the likelihood of transmitting these agents, young children should not handle these animals during their treatment period. When adults handle these animals to apply treatment, they should wear protective gloves. Gloves should also be worn when cleaning the animal’s enclosure. Overall, chinchillas represent a low potential zoonotic risk for pet owners; however, clients should always be advised to follow standard disinfection and hygiene practices when working with these animals.

REFERENCES 1. Merry CJ: An introduction to chinchillas, Vet Tech 11(5):315-322, 1990. 2. Webb RA: Chinchillas. In Beynon PH, Cooper JE, editors: Manual of Exotic Pets, Cheltenham, Gloucestershire, 1991, British Small Animal Veterinary Association. 3. Quesenberry KE, Donnelly TM, Hillyer EV: Biology, husbandry, and clinical techniques of guinea pigs and chinchillas. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 4. de Oliveria Silva T, Kruetz LC, Barcellos LJG et al: Reference values for chinchilla (Chinchilla laniger) blood cells and serum biochemical parameters, Ciência Rural, 35(3):602-606, 2005. 5. Silverman S, Tell LA: Radiology equipment and positioning techniques. In Silverman S, Tell LA, editors: Radiology of Rodents, Rabbits, and Ferrets: An Atlas of Normal Anatomy and Positioning, St Louis, 2005, WB Saunders. 6. Moore RW, Greenlee HH: Enterotoxaemia in chinchillas, Lab Anim 9:153-154, 1975. 7. Bartoszcze M, Nowakowski M, Roszkowski J et al: Chinchilla deaths due to Clostridium perfringens a enterotoxin, Vet Rec 126:341, 1990. 8. Hoefer HL: Chinchillas. Vet Clin North Am Small Anim Pract 24(1):113120, 1994. 9. Kraft H: Diseases of Chinchillas, Neptune City, NJ, 1987, TFH. 10. Jenkins JR: Husbandry and common diseases of the chinchilla (Chinchilla laniger), J Small Exot Anim Med 2(1):15-17, 1992. 11. Naglic´ T, Šeol B, Bedekovic´ Ž et al: Outbreak of Salmonella enteritidis and isolation of Salmonella sofia in chinchillas (Chinchilla laniger), Vet Rec 152:719-720, 2003. 12. Finley GG, Long JR: An epizootic of listeriosis in chinchillas, Can Vet J 18(6):164-167, 1977. 13. Kimpe A, Decostere A, Hermans K et al: Isolation of Listeria ivanovii from a septicaemic chinchilla (Chinchilla lanigera), Vet Rec 154:791-792, 2004. 14. Donnelly TM: Disease problems of chinchillas. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 15. Eidman S: Studies on the etiology and pathogenesis of fur damage in the chinchilla. Hannover, Germany, Tierarztliche Hochschule 61:163, 1992. 16. Newberne PM: An outbreak of bacterial gastro-enteritis in the South American chinchilla, North Am Vet, 34:187-188, 191, 1953. 17. Yamini B, Raju NR: Gastroenteritis associated with a Cryptosporidium sp. in a chinchilla, JAVMA 189:1158-1159, 1986.

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492 18. Sanford SE: Cerebrospinal nematodiasis caused by Baylisascaris procyonis in chinchillas, J Vet Diagn Invest 3:77-79, 1991. 19. Newberne PM, Siebold HR: Malignant lymphoma in a chinchilla, Vet Med 48:428-429, 1953. 20. Osofsky A, Verstraete FJM: Dentistry in pet rodents, Compend Contin Educ Dent Jan:61-74, 2006. 21. Crossley DA: Dental disease in chinchillas in the UK, J Small Anim Pract 42:12-19, 2001. 22. Richardson VCG: Chinchillas: systems and diseases. In Richardson VCG, editor: Diseases of Small Domestic Rodents, Oxford, UK, 2003, Blackwell. 23. Verstraete FJM: Advances in diagnosis and treatment of small exotic mammal dental disease, Semin Av Exotic Pet Med 12(1):37-48, 2003. 24. Legendre LFJ: Malocclusions in guinea pigs, chinchillas, and rabbits, Can Vet J 43:385-390, 2002. 25. Crossley DA, Jackson A, Yates J et al: Use of computed tomography to investigate cheek tooth abnormalities in chinchillas (Chinchilla lanigera), J Small Anim Pract 39:385-389, 1998. 26. Varonjack WJ, Johnson HD: Relationship of thyroid and adrenal function to “fur-chewing” in the chinchilla, Comp Biochem Physiol 45:115120, 1973. 27. Tišljar M, Janic´ D, Grabarevic´ Ž et al: Stress-induced Cushing’s syndrome in fur-chewing chinchillas, ACTA Vet Hung 50(2):133-142, 2002. 28. Williams J: Orthopedic radiography in exotic animal practice. In Tully TN, editor: Vet Clin North Am Exot Anim Pract 5(1):1-22, 2002.

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29. Hoefer HL, Crossley DA: Chinchillas. In Meredith A, Redrobe S, editors: BSAVA Manual of Exotic Pets, ed 4, Quedgeley, Gloucester, 2005, British Small Animal Veterinary Association. 30. Morgan RV, Moore FM, Pearce LK et al: Clinical and laboratory findings in small companion animals with lead poisoning: 347 cases (1977-1986), JAVMA 199:93-97, 1991. 31. Cousens PJ: The chinchilla in veterinary practice, J Small Anim Pract 4:199-205, 1963. 32. Marlow C: Diabetes in a chinchilla, Vet Record 136:595-596, 1995 (letter). 33. Plumb DC: Veterinary Drug Handbook, ed 5, Ames, Iowa, 2005, Blackwell. 34. Morrisey JK, Carpenter JW: Formulary. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 35. Carpenter JW: Exotic Animal Formulary, ed 3, St Louis, 2005, WB Saunders. 36. Briscoe JA: Techniques for emergency airway and vascular access in special species, Semin Av Exot Pet Med 13(3):118-131, 2004. 37. Bennett RA, Mullen HS: Soft tissue surgery. In Quesenberry KE, Carpenter JW, editors: Ferrets, Rabbits, and Rodents: Clinical Medicine and Surgery, ed 2, St Louis, 2003, WB Saunders. 38. Jenkins JR: Surgical sterilization in small mammals. In Bennett RA, editor: Vet Clin North Am Exot Anim Pract 3(3):617-627, 2000.

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Maya Bewig Mark A. Mitchell

C H A P T E R

1 9

WILDLIFE

IMPORTANCE OF VETERINARIAN PARTICIPATION IN WILDLIFE CARE As humans continue to encroach on previously undisturbed and pristine wild habitats, the frequency of wildlife-human encounters has increased. These increased encounters have led to an increased need for veterinary participation in managing sick and injured animals. For much of the public, veterinarians are still held in reverence because of the James Herriott stories. We believe that it is important for veterinarians to maintain this image. This does not necessarily suggest that veterinarians should consider themselves experts on all animal species, but that as professionals, they should develop an infrastructure of resources that enables them to provide medical and surgical care for those animals in need. In addition, because many of these animals can serve as reservoirs for zoonotic diseases, veterinarians can play an important role in minimizing the risk of disease transmission between wildlife and concerned citizens. For some veterinarians, their role may require them to maintain a current list of veterinarians or wildlife rehabilitators capable of managing an injured wildlife patient and refer clients to these professionals. For others with a more invested interest, it may mean that they will pursue available resources to develop the knowledge and skill for managing these animals in their hospital. The last thing clients, or prospective clients, want to hear from veterinarians is “We don’t work with those animals.” Instead, if veterinarians can, at a minimum, direct clients to the most appropriate resource, veterinarians maintain (and justify) their image among the professions.

DECIDING WHETHER OR NOT TO TREAT WILDLIFE There are many financial, ethical, and emotional issues for veterinarians to consider when deciding whether to accept wildlife cases to their practice. Wildlife is not owned and therefore do not come with paying caretakers. In many cases, the hospital will be expected to absorb the cost of treatment, although avenues for monetary compensation, including grants and public donations, do exist. Accepting wildlife cases is often perceived by (prospective) clients as a positive reinforcement of a veterinarian’s compassion toward animals and can serve, directly or indirectly, as a method of increasing a veterinarian’s domestic and exotic pet caseload. One ethical consideration to make with these cases is deciding when intervention may interfere with a natural process occurring in a population. In some cases, euthanasia will certainly end suffering, whereas in others treatment may interfere with both natural selection and the disease status in a population. However, because the lives of humans have become so enmeshed with populations of wild animals, it is unavoidable that some, if not most, of the injuries or disease processes occurring in wildlife populations are the direct result of human contact. In these cases, there may be a strong moral (ethical) obligation to repair damage caused by the human species. An early and rational assessment of the likely return to function for a patient is imperative to conserve the available resources for treatment and prevent undue suffering in the patient. Animals who are considered poor candidates for release or placement should be considered for euthanasia as soon as an assessment is made. Release candidates should be able to function appropriately within their natural

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494 habitat (including reproduction) and with conspecifics; otherwise, a disservice is rendered to both the animal and its population. Certain factors should be weighed before deciding to accept wildlife cases to a practice. These factors should be considered with every case as well. The potential costs include those welfare expenses associated with captivity, treatment, release, and failure to reestablish the animal in the wild, as well as the welfare risks to conspecifics and other species through the possible introduction of infection or competition for resources and the upset in natural selection (e.g., treating animals that have increased susceptibility to disease may inadvertently select for less fit animals). The potential benefits associated with working with these animals include the emotional pleasure humans derive from helping a “lesser” species, the potential to educate the public, and the opportunities this type of medicine provides for monitoring threats to wildlife and human populations. Assessing the balance or inherent value between welfare costs and benefits is often difficult. In addition to the unpredictability of responses between individual cases to treatment, there are few robust yardsticks for use in judging, for example, whether any stress or pain associated with captivity and treatment does or does not outweigh the potential welfare benefit to the animal. The task is not made easier by the scarcity of information on outcomes of wildlife rehabilitation cases, although more studies are being done. Veterinarians have an important role in these ethical assessments and in helping to design protocols that will best manage the welfare of these animals in both captive and wild settings.

KNOWING THE REGULATIONS Veterinarians working with wildlife should become familiar with the regulations addressing the handling, transport, and treatment of these animals. These regulations have been enacted at the federal and state levels to ensure the protection of these animals against individuals that would otherwise exploit them (e.g., meat or fur). Historically, veterinarians were perceived as “good Samaritans,” and it was just accepted that veterinarians would or could provide any medical and surgical care that a particular wildlife patient required. Unfortunately, some veterinarians, like many unlicensed wildlife rehabilitators, will attempt to complete the rehabilitation process for their wildlife patients, even though they cannot meet their patient’s husbandry, nutritional, or posttreatment rehabilitation needs. Currently, federal and state governments are committed to maintaining veterinarians as an integral component to the care of injured wildlife and consider veterinarians to be the experts for providing medical and surgical care for these animals. However, these agencies also realize that most veterinarians do not have the time or facilities to fully rehabilitate injured wildlife. Because of this, federal and state agencies require veterinarians (without state and/or federal rehabilitation permits) to transfer these animals to licensed rehabilitators to complete the rehabilitation process. It is expected that a veterinarian will transfer any wildlife patient to a licensed wildlife rehabilitator within 24 hours of stabilizing the patient. A stabilized patient is one that is no longer in dire need of

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veterinary medical care. Patients that require medical treatment that can be administered via an experienced (trained) rehabilitator should be transferred. The federal government is primarily invested in the protection of migratory birds and all other species that are listed federally as species of special concern, threatened, or endangered. State agencies also require rehabilitators to maintain permits to cover these same species, as well as mammals and other species (reptiles, amphibians, and fish) considered threatened or endangered at the state level. Veterinarians interested in obtaining federal permits can find information regarding the permits on the United States Fish and Wildlife Service website (www.fws.gov). Veterinarians interested in obtaining state rehabilitation licensure should contact their state wildlife and fisheries service.

BECOMING MORE INVOLVED Wildlife medicine is in its infancy. Because of this, much of what veterinarians do is based on anecdotal or subjective recommendations. With this knowledge, it is important for those veterinarians working with these animals to document their work and disseminate it to others. This information can be disseminated through presentations at local, state, or national wildlife rehabilitation meetings, or published in appropriate bulletins or journals. In our opinion, the knowledge base regarding wildlife medicine, more so than any other aspect of veterinary medicine, would benefit from more active participation by veterinarians.

COMMON SPECIES PRESENTATIONS Veterinarians that publicize an interest in accepting wildlife cases may be surprised by the diversity of cases being presented to their hospitals. In our practices, amphibians, reptiles, birds, and mammals are all routinely presented. Such a diverse caseload requires veterinarians to be adaptive to the different needs of these animals, both in their medical and surgical knowledge and in their ability to meet the rehabilitation needs (e.g., diet, environment) of the patient. Many veterinarians will limit the types of cases presented to their hospitals. Some of the veterinarians we consult with only accept chelonians, whereas others may accept only avian cases. It is important that veterinarians inform wildlife rehabilitators and clients of particular species they exclude from their practices. It would also be helpful to have a list of other veterinarians on hand that do accept those species, so that those individuals can be referred to an appropriate facility.

PREPARING FOR WILDLIFE IN VETERINARY HOSPITALS Staff Preparedness When veterinarians accept wildlife cases into their hospitals, it is important that they have buy-in from their staff. Wildlife cases can require a significant amount of time and effort,

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especially orphaned animals. Therefore, it is often the veterinary staff that will have the most contact with the wildlife cases. Veterinarians that accept these types of cases on a moderate to large scale would certainly benefit from providing at least some of their staff an opportunity to obtain continuing education in wildlife rehabilitation. Many of these opportunities can be obtained from local or state meetings, although the large national meetings (e.g., the annual conferences of the National Wildlife Rehabilitators Association and the International Wildlife Rehabilitators Association) provide the most diverse training opportunities. Membership in these organizations is also highly recommended, as they publish regular bulletins or journals and provide a real resource of potential personal contacts. It is important that veterinarians train their staff to avoid becoming emotionally attached to their wildlife patients. In our opinion, this is the single most controversial issue that can arise in a hospital. If staff members (and veterinarians) become emotionally attached to their cases, they are less likely to maintain an objective outlook on their cases. It is important to always approach these cases with a triage mentality (Figure 19-1). The time spent treating these patients should also be limited to necessary contact only. Animals that become habituated to humans in captivity may be at greater risk of reinjury or death after release if they lose their fear of humans. For orphaned animals, imprinting is a major concern. Imprinting occurs when an animal recognizes a human as the “parent” animal. Imprinting can potentially lead to dangerous encounters for humans and wildlife, especially with mesopredators, carnivores, and raptors. For this reason, it is important that only trained staff manage orphaned animals. When accepting wildlife patients into a hospital, veterinarians must maintain a complete record for each patient. We have found that staff can prove invaluable for this task. The following information should be collected for each patient: the species, age (if known or adult/juvenile), gender (if known),

date of presentation, location and date found, type of injury, medical treatment provided (if any), diet and husbandry provided (if any), and name and contact information of the person(s) presenting the animal. Each patient should also be thoroughly evaluated daily and the data recorded using a standard format (SOAP: subjective, objective, assessment, plan). Maintaining this information in a computer database will allow for routine review and analysis of the data. This information can then be used to develop reference ranges for a particular species or area and can be published and presented to a wider audience.

Equipment Most of the medical and surgical equipment required for wildlife patients is already available in veterinary hospitals that treat domestic species. However, if a practice is primarily based on domestic species, there may be some specialty equipment that needs to be obtained. A list of different equipment we consider important can be found in Box 19-1.

BOX 19-1

Specialty Equipment Required to Manage Wildlife at a Veterinary Hospital

Incubators Incubators are thermostat-controlled microhabitats for small mammals, birds, reptiles, and amphibians. These enclosures should be capable of providing a range of environmental temperatures (80°-100° F). Oxygen Cage Both orphaned animals that aspirate during feeding and avian species with respiratory disease may benefit from placement in an oxygen cage. Ophthalmic Loupe A loupe can be used to provide better definition to small structures (e.g., blood vessels). Endoscopic Equipment A 2.7-mm rigid endoscope can be used to perform exploratory coeliotomies in birds. Sexing birds can also be done using this equipment. Radiosurgery A radiosurgery unit can be used to control hemostasis. This can be especially important in small patients. Restraint Materials Leather gloves are required to restrain raptors and mesopredators. Body wraps can be used for birds. A rabies pole or pole-syringe can be used to handle mesopredators.

Figure 19-1 This barred owl presented with a degloving injury over the entire antebrachium. Although the animal was alert, responsive, and eating, it was euthanized because of the extensive tissue necrosis associated with the wound.

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Digital Scale A scale or scales capable of measuring the variety of animals being presented to the facility are needed. We suggest a scale capable of measuring 1-g increments.

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HOUSING Veterinarians that accept wildlife cases into their practices must be prepared to house a variety of species. It is always best to house wildlife separately from domestic species, to minimize the likelihood of interspecies disease dissemination; however, when this is not possible, all attempts should be made to minimize transmission (e.g., wear gloves, clean cages with separate sponges). The following are some recommendations for housing different wildlife species.

Avian PASSERINES The most difficult aspects of providing housing for small birds is preventing escape and providing security. Using a room that can be darkened will facilitate capture of escapees. Mesh or metal used to cover a cage must have a small diameter to prevent escape. It can also be important to have the enclosure open into a room that will allow for ease of capture (e.g., low ceilings, few hiding spots) should the bird escape the primary cage. The setup of feeding and water stations will vary greatly depending on the species.

PIGEONS AND DOVES For short-term housing, any container, such as a dog or cat kennel, is sufficient. Because pigeons and doves are reservoirs for various avian pathogens, it is always best to isolate them from other species to limit the potential for disease transmission. For longer periods in captivity, these birds may be best housed in an aviary.

GAMEBIRDS Principles of short-term and long-term housing are the same for these birds as was described for pigeons and doves, although the size of the cage may need to be increased for larger specimens. No perching materials need to be provided for shortterm housing but should be provided if the bird is kept for a longer period of time to allow roosting at night.1 Substrate should be easily disinfected, to prevent foot disease (e.g., Astroturf is a good substrate).

WATERFOWL For short-term waterfowl care, the principles are the same as with those previously mentioned, with the exception of providing water in a container large enough for the bird to stand in. For long-term care, adequate housing will require the addition of a large pool or pond, which may not be feasible in a general practice setting.

WADERS Short-term housing for waders should be tall enough to enable the animal to stand naturally. The cage should also be kept dark and quiet with moist, nonslip flooring (e.g., moist

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sand or newspaper).2 Water should be provided in a shallow container. For long-term housing, a shallow pond or pool and suitable natural cover should be provided. As with waterfowl, this may be difficult for general practitioners to provide.

SEABIRDS Seabirds have special requirements in captivity that may make rehabilitation difficult, especially for beginners. Some species of seabird are gregarious and do better when housed in groups. Seabirds are sensitive to sudden noises and should be kept in a quiet environment. A soft substrate that is easy to clean, such as Astroturf, rubber mats, or deep layers of wood shavings, should be used. Animals left on firm substrates (metal flooring or concrete) can develop pododermatitis. Straw or hay bedding should never be used, as they can be a source of fungal spores. It is important to provide ample ventilation to minimize the risk of respiratory disease. For stable patients in medium- to long-term captivity, a water source (e.g., pool or pond) must be provided. This water source should be cleaned regularly to prevent autoinfection.

RAPTORS All of the general principles for housing apply to raptors. In addition, it is extremely important to provide an environment that will prevent any self-induced trauma. Raptors are prone to damaging their feathers, wings, talons, and cere in captivity. Injuries to any of these body parts can delay the rehabilitation process and should be avoided at all costs. The best cage materials to use for housing raptors are those that do not bend or damage the feathers. Because most veterinary hospitals maintain chain-link fence kennels or stainless steel cages, housing for raptors may be limited. Covering the cage with a soft mesh can help diminish the amount of feather damage done to hospitalized raptors. Tailguards made out of old radiograph film can be used to reduce tail feather damage. Perches can also be used to help limit the amount of damage done to the tail feathers. The perch should be positioned so that the tail feathers do not touch the base of the cage and the flight feathers have minimal contact with the sides of the cage. For extended hospitalization, raptors should be housed in an appropriately sized aviary. Whereas groups of some species can be housed together, mixing raptors is not recommended. As with shortterm housing, care must be taken that the animal cannot injure itself. An easily disinfected substrate, such as Astroturf or gravel, can be used for the aviary. Multiple perches should be provided. If the bird is not a solid flier, the perches should be maintained close to the ground. Covering the perches with a rough surface (e.g., Astroturf) will minimize the likelihood of a bird developing pododermatitis. Multiple diameter and textured perches should be placed in the cage; however, the diameter should not be so small that the talons puncture the plantar surface of the foot.3 The perches should be cleaned regularly to minimize the likelihood of fecal contamination. As with other species, hay or straw should not be used because of the potential for exposure to Aspergillus spp. spores. Certain species, such as owls, will benefit from shelter.

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Mammals SMALL RODENTS Housing for small rodents must be constructed to withstand aggressive chewing and escape. For adult rodents, wood, some plastics, and aluminum caging materials are not recommended. Galvanized or stainless steel are more appropriate, as they can handle chewing and are easier to disinfect. For short-term housing, adult rodents can be housed in stainless steel hospital kennels. It is important, of course, that the width of the bars be narrow enough to prevent escape. Within the enclosure, a nest box should be provided, which can be made from wood or cardboard. Recycled newspaper is a suitable substrate. Tree limbs and other natural substances are also recommended and will provide the animals a surface to chew on. Orphaned animals or nonchewing species (e.g., bats) can be housed in clip-top plastic containers. It is important to drill a sufficient number of holes in the container to provide ventilation. Newspaper or towels can be used as a substrate for these cages. For long-term rehabilitation, larger portable cages are recommended. Galvanized wire is probably the least expensive material for constructing these cages. It is best to secure these cages to protect against predation.

LAGOMORPHS Lagomorphs are extremely sensitive and prone to stress, so maintaining a quiet, predator-free environment is essential. Care must be taken to make the enclosure escape-proof. Covering the cage will increase the feeling of security for the animal. Attention to the flooring is also important, as these animals like to dig. A hay bedding or false floor can be used to simulate a burrow. The cage types described for rodents can also be used for lagomorphs.

MESOPREDATORS Mesopredators can be housed in standard domestic animals stainless steel cages; however, this is not without its limitations. Because these cages have swing-doors, it can be difficult to work with these animals if they are fractious. For animals that are difficult to manage, anesthesia is highly recommended. A pole-syringe capable of being inserted between the cage bars can be used to safely deliver an appropriate anesthetic. Shifting these animals to outdoor cages is highly recommended. Chainlink cages can be used to house these animals, but they must be constructed to be escape-proof. Whether inside or outside, the cage should be provided with a shelter box and an area for a latrine.

DEER For short-term hospitalization, fawns can be kept in an isolation area or any area that is away from the noise of humans and other animals. Large deer can be kept for short periods of time in a stable or outhouse with a deep layer of straw. As for the size of the enclosure, a maximum of 1.5 × 1.8 m and 2 × 3 m for fawns and adult deer, respectively, is recommended.4 The enclosure should not have windows, and ventilation should be provided through slats at the top of the walls or

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ceiling. A door that is divided into a stable-door arrangement, with the top portion only 0.5 m high, is preferred. It is best if the door can be swung inward, so that it can be used as a restraint device for the deer. The top portion of the door should be opened for access only from above. Deer will leap toward light when the door is opened, making other arrangements dangerous. Deer who will be kept in captivity for longer periods of time should be transferred to an experienced rehabilitator who has appropriate pens.

Reptiles and Amphibians The housing needs of reptiles and amphibians will vary greatly depending on species. Some examples of reptile groupings by environment include those that are aquatic, semiaquatic, arboreal, or terrestrial. Of the terrestrial reptiles, subgroups include those that are fossorial (burrowing), thigmotactic (prefer to wedge themselves into rocky crevices when not basking), and those that live on the surface of the soil. Reptiles can also be grouped according to the time of day that they are active: diurnal, nocturnal, and crepuscular (active during dusk or dawn). Species habits must be known before appropriate housing can be provided. Reptiles kept in inappropriate environments can develop a variety of conditions that will delay the rehabilitation process. For example, terrestrial species kept in moist cages may develop integumentary disease, whereas aquatic species housed in dry habitats will become dehydrated. Arboreal reptiles must be provided with branches, reptiles adapted to saline or brackish water should be furnished with water of the appropriate salinity, fossorial species must be provided substrate to burrow in, and secretive reptiles must be provided hiding places. Water should be provided in the form in which the animal is used to imbibing it. Many lizards drink from rain or dewdrops on foliage and will not drink from dishes. Tortoises should be provided with a shallow bowl of water. For additional information regarding the specific husbandry needs of reptiles, see the appropriate chapters in this text.

QUARANTINE Quarantine is an important consideration when practicing wildlife medicine in a domestic species veterinary hospital. Unfortunately, most veterinary hospitals are not built with quarantine in mind. Because wildlife can harbor various bacterial, fungal, viral, and parasitic diseases that can be transmitted to domestic pets, it is important to minimize and restrict both direct and indirect contact between these animals. Wildlife should be housed in a separate room from domestic species. A room with its own ventilation system is preferred. The traffic through the wildlife ward should be one-way. A foot bath with a disinfectant (e.g., sodium hypochlorite) should be placed outside the doorway and used after exiting the room to minimize the likelihood of tracking infectious diseases throughout the hospital. The disinfectant should be changed daily, or as often as needed, to minimize the amount of organic debris in

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498 the solution. Organic debris renders many disinfectants useless. Laboratory coats, or preferably jumpsuits that totally cover clothing, should be placed in the wildlife ward and worn when working with the animals. These clothing materials should be taken out of the room only for washing. Any materials being removed from the ward should be placed in a garbage bag and carried through the hospital in these bags to minimize the likelihood of disseminating disease through the hospital. A hand-washing station should be placed within the room. Signs should be posted on the wildlife ward door to alert staff and clients about the presence of wildlife and the need to maintain a strict quarantine protocol.

ZOONOSES Zoonotic diseases should always be a concern for those individuals working with wildlife, as many of these animals can harbor a variety of bacterial, viral, fungal and parasitic zoonotic agents (Box 19-2). Because of this, children and individuals with compromised immune systems should not be allowed to work with wildlife. Staff should not be allowed to eat, drink, or smoke near any wildlife patients, and food and drink items should not be allowed near any of the places where diagnostic samples are held. We strongly recommend wearing examination gloves when handling wildlife. Wearing gloves will help minimize the likelihood of introducing pathogens into cuts or abrasions on practitioners’ and staff’s hands. For those cases where an aerosolized pathogen is suspected, a protective mask and eyeglasses should be worn. Individuals who develop an illness after working with wildlife should be examined by a health care specialist immediately. Zoonoses and Communicable Diseases Common to Man and Animals (ed 3), coedited by P. N. Acha and B. Szyfres and published by the Pan American Health Organization (Washington DC), is an excellent reference for obtaining additional information on zoonotic diseases associated with wildlife.

EMERGENCY CARE General Concepts When wildlife cases are presented to a veterinary hospital, they should be considered an emergency. Most of these patients will have injuries that are hours to days old that have limited their ability to obtain food, water, or shelter. Because of these limitations, many of these animals will be dehydrated, cachectic, and possibly hypo- or hyperthermic (seasonally dependent). To increase the likelihood of success with these cases, it is vital that the veterinarian stabilize these patients before pursuing a case diagnosis.

Cardiopulmonary Resuscitation In some emergency situations an animal may present with or develop cardiac or respiratory failure during a course of treatment. Veterinary staff should determine beforehand the level of response they will provide in these cases. If the clinic adheres

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BOX 19-2

A Partial List of Common Zoonotic Diseases Associated with Wildlife

Bacterial Chlamydophila psittaci Salmonella spp. Campylobacter spp. Escherichia coli Staphylococcus aureus Bacillus anthracis Streptococcus spp. Mycobacterium spp. Leptospira interrogans Clostridium spp. Francisella tularensis Yersinia pestis Yersinia pseudotuberculosis Fungal Aspergillus spp. Histoplasma spp. Blastomyces spp. Viral Rabies West Nile virus St. Louis encephalitis Eastern equine encephalitis Influenza Parasitic Giardia spp. Cryptosporidium parvum Baylisascaris procyonis Ancylostoma spp. Sarcoptes spp. Encephalocytozoon cuniculi

to a strict triage protocol, then most cases of cardiac or respiratory failure will not be treated. If the clinic does attempt to resuscitate these animals, then the staff should be prepared for a low success rate (10%

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always be warmed to the patient’s physiologic temperature before being administered. Identifying which fluid is most appropriate will be based on the type of dehydration. Most wildlife cases present with isotonic dehydration, so a balanced fluid (e.g., 0.9% saline, lactated Ringer’s solution) can be used. Calculating the patient’s osmolality is the best method for estimating the type of dehydration. Laboratories using osmometers can calculate this number; however, if submitting a sample is not possible, then an estimate can be made using the following mathematical formula: OsM = 2(sodium + potassium) + urea + glucose. For reptiles and birds, uric acid can be used as a substitute for urea; however, it may underestimate the true osmolality. A fluid deficit in a mammalian patient can be corrected within 24 to 36 hours, whereas in birds and reptiles, it may take 48 and 96 hours, respectively. Fluids can be delivered to wildlife using the same basic techniques described for domestic pets. Animals that are only mildly dehydrated and have a functional gastrointestinal tract can have fluids delivered per os. Subcutaneous fluids can be used for those patients that are mildly dehydrated but have some gastrointestinal disease (e.g., diarrhea, vomiting). For moderately dehydrated reptile and mammalian patients, fluids can be given intracoelomically or intraperitoneally, respectively. Because birds have air sacs, they should never be given fluids intracoelomically. For moderate to severe dehydration, fluids should be given intravenously (Figure 19-4) or intraosseously.

Fracture and Wound Management Fractures should be stabilized to minimize the risk of further injury to the patient and to control pain. Depending on the site and nature of the fracture, various emergency bandaging methods can be employed. We generally immobilize wing fractures with a figure-of-eight splint (Figure 19-5) or a body wrap. The figure-of-eight splint is appropriate for fractures distal to

Estimating Dehydration in Wildlife Clinical signs Some loss of skin turgor Skin slow to return after tenting (2-3 sec) Some tackiness in mucous membranes Capillary refill time (3 sec) Sunken eyes (loss of retroorbital fat pads)

Figure 19-4 Intravenous fluids are highly recommended for those patients that are severely dehydrated. In this white-tailed deer fawn, a jugular catheter was placed.

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Figure 19-5 This screech owl presented with a radial fracture. Because the fracture was reduced, the wing was splinted using a figure-of-eight bandage.

the elbow, and a body wrap is recommended when the fracture includes the humerus. Vet-wrap (3M Corp., St. Paul, MN), or another comparable bandage material, can be used. For small birds (e.g., hummingbirds), placing the wings in a normal position and taping the primaries where they cross can provide an adequate splint. When immobilizing the wings of a bird, it is important to change the bandage and provide physical therapy every 3 to 4 days. This will prevent excessive contraction of the patagial tendon and atrophy of the muscles. We prefer to replace the bandage under anesthesia, as this minimizes the likelihood of the animal reinjuring itself and controls the pain associated with the procedure. In birds, fractures of the legs can be immobilized with modified syringe-case splints, tape splints, a ball bandage, or a Robert Jones–type splint. Syringe case splints are best suited for fractures distal to the stifle. Tape splints can be used to stabilize toes or the distal leg of a passerine or other lightweight bird. We generally use white, porous tape for these splints. For mammals, limbs can be splinted using the standard techniques described for domestic mammals. Fractures of the reptile limb can be immobilized by taping the limb to the animal’s body. In lizards, a forelimb can be secured against the body wall using porous tape while the rear leg can be immobilized against the tail. In chelonians, the limb can be reduced into a normal position within the shell and taped into place. When splinting any limb, it is important to always immobilize the joint above and below the fracture site. Wound management should be initiated to limit the amount of fluid or moisture lost from the break in the integument and to limit wound contamination. There are a variety of methods that can be used to manage a skin wound in a wildlife patient. The first and most important step to managing a wound is to remove any necrotic tissue. If necrotic tissue is left behind, it will delay the healing process. Sharp dissection of the wound should be done to remove all of the necrotic tissue. Wound disinfection can be done by irrigating the

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wound with dilute chlorhexidine, Betadine, or 50% dextrose. We have used the hyperosmotic dextrose, with excellent results. Others report similar results with honey. After disinfection, the wound should be rinsed with a warmed sterile saline. Once disinfected, the wound should be protected against additional contamination. Silver sulfadiazine, or another topical antimicrobial, can be applied to the wound to protect it against opportunistic pathogens and to maintain the hydration of the exposed tissues. Bandage material can be applied to the wound if additional protection is required. A wet-to-dry bandage can be applied to any wound that requires decontamination. Depending on the size of the wound, either 2″ × 2″ or 4″ × 4″ gauze pads can be used for the bandage. The first 4 to 5 gauze pads should be irrigated with sterile saline or a disinfectant, such as chlorhexidine. The gauze pads should be wrung out before being applied to the wound. An additional 4 to 5 dry gauze pads should be placed on top of the moistened pads. All of the gauze should be secured to the wound with Vet-wrap. The bandage should be changed daily. Any necrotic tissue should be removed in between bandage changes. While uncovered, the wound should be disinfected using the technique described previously. The wet-to-dry bandage should be used only until the wound is considered disinfected. Prolonged use of this bandage can lead to tissue desiccation.

THERAPEUTICS Once fluid therapy is initiated, chemotherapeutics can be given. It is important to wait on certain drugs until after fluid therapy has been instituted. For example, the administration of steroidal antiinflammatories could have a negative response if given to a dehydrated patient. A variety of chemotherapeutics are available to the veterinarian, including antiinflammatories (steroidal and nonsteroidal), analgesics, antibiotics, antifungals, antivirals, anesthetics, antiemetics, chelating agents, vitamins, and minerals. A list of common drugs used to treat wildlife can be found in Table 19-3.

CAPTURE AND TRANSPORT Most injured wildlife cases will be transported to a veterinary clinic by the public, but there may be times when veterinarians are in a situation that requires them or their staff to capture an animal or give counsel on appropriate capture and transport to a member of the public. The techniques vary according to the species being approached, but some generalities apply. If observing the animal before capture, it is best to evaluate its posture and attitude from a distance (e.g., alert, depressed, lethargic). This information can be used to develop a capture strategy. Animals that are depressed and lethargic should be approached cautiously, as they may be more susceptible to sudden death or capture myopathy. When developing a capture strategy, it is also imperative to consider the humans involved with the procedure, to ensure no harm comes to those involved. Before capture, identify the major “weapons” an animal will use to defend itself. Raptors, for example, may use their talons and beak as weapons.

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Drugs Commonly Used to Treat Wildlife

Drug

Dose

Comments

0.5 mg/kg 0.5-1.0 mg/kg 20 mg/kg 0.5-1.0 mg/kg

IV, IO, IT IV, IO IV, IO, IT 1 : 1,000; IV, IO, IT

3-5 mg/kg 15-20 mg/kg 2.5-10 mg/kg 10-30 mg/kg 100 mg/kg 10-20 mg/kg 20-40 mg/kg 100 mg/kg 15-30 mg/kg 20 mg/kg 10 mg/kg 15 mg/kg 15 mg/kg 5-10 mg/kg 25-100 mg/kg 5-10 mg/kg 10-15 mg/kg 5-10 mg/kg 200 mg/kg 20 mg/kg 15-30 mg/kg 50-75 mg/kg 5-25 mg/kg

Reptiles: IM, q72h Birds: IM, q24h Mammals: IM, q12-24h Reptiles: PO, IM, q12-24h Birds, PO, q12h Mammals: PO, q12h; NOT recommended for rodents or lagomorphs Reptiles: PO, q12h Birds, PO, q12h Mammals: PO, q12h; NOT recommended for rodents or lagomorphs Reptiles: IM, q72h Reptiles: PO, q24h Birds: PO, q12h Mammals: PO, q12h Reptiles: PO, q24h Birds: PO, q24h Reptiles: PO, q24h Birds: PO, q12h Mammals: PO, q12h Birds: PO, q12-24h Mammals: PO, q12h; use with caution in lagomorphs and rodents Reptiles: PO, q24h Birds: PO, q12-24h Mammals: PO, q12-24h

5-10 mg/kg 2.5-10 mg/kg 2.5-5 mg/kg 25-50 mg/kg 20-50 mg/kg 20-50 mg/kg 100,000 IU/kg 100,000-300, 000 IU/kg

Reptiles: PO, q24-48h; monitor for neurologic disease Birds: PO, q24h; monitor for neurologic disease Mammals: PO, q24h; monitor for neurologic disease Reptiles: PO, q24h Birds: PO, q12-24h Mammals: PO, q24h Reptiles: PO, q24h Birds: PO, q12h

25-100 mg/kg 5-10 mg/kg 0.2-0.4 mg/kg 25-50 mg/kg 20-40 mg/kg 15-25 mg/kg 5-8 mg/kg 20-40 mg/kg 5-10 mg/kg

Avian, reptiles: PO, once, repeat 10-14 days Mammals: PO, q24h × 3-5 days All wildlife: IM, PO; Do not use in chelonians Reptiles: PO, q24-48h Birds: PO, q12-24h Mammals: PO, q12-24h Reptiles: IM, repeat 10-14 days Birds: PO, q12-24h Mammals: IM, repeat 10-14 days

Emergency drugs Atropine Diazepam Doxapram Epinephrine Antibiotics Amikacin

Amoxicillin

Cephalexin

Ceftazidime Ciprofloxacin

Doxycycline Enrofloxacin

Tetracycline Trimethoprim-sulfadiazine

Antifungals Itraconazole

Ketoconazole

Nystatin Antiparasitics Fenbendazole Ivermectin Metronidazole

Praziquantel

IM, Intramuscular; IO, intraosseous; IT, intratracheal; IV, intravenous; PO, per os.

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An appropriate-sized, darkened cardboard box (with ventilation holes cut in) or pet carrier that opens from the top can be used to transport wildlife. The cardboard box is an excellent transport carrier because it can be discarded after use; the pet carrier can be disinfected between uses. Homemade wooden boxes may also be used but are more difficult to disinfect. It is important to include a nonslip floor covering in the transport container to ensure proper footing in the transport box. Noncovered cardboard surfaces have been associated with the development of splay leg and bilateral partial paresis in some bird species.5 When transporting birds, it is imperative that the box be large enough that the bird can turn without damaging its feathers. When flight or tail feathers are damaged, it can increase the rehabilitation time for a patient.

extended behind the bird. Care must be taken that the head is always sufficiently restrained.

SEABIRDS

Small and medium-sized birds, including songbirds, pigeons, doves, and crows, pose few physical risks to humans, although the larger birds in this group can deliver painful bites or scratches. These animals are best captured using a small diameter hole net or cloth. A cloth sack may be used if no box is available for transport.

The main danger posed by seabirds is from their beaks, which they may use to bite or stab at the face. Some seabirds also have sharp claws, which are capable of causing deep lacerations. As with waders, safety goggles should be worn and the bird never restrained close to the face. If the species in hand has internal nares, be careful not to hold the beak closed because it can affect normal respiration. These birds will also attempt to return to the water when they sense danger, so care must be taken to block this route of escape. A net is usually necessary for capture, but a towel or blanket may also be used. The beak can be kept closed by using tape or a rubber band. The head of these animals should be restrained and covered at all times, to prevent injury to the handler and reduce the stress on the animal. A towel can be wrapped around the body of these birds to restrain the wings; however, the towel should not be held so tight that it affects respiration (e.g., keel excursions). Because seabirds commonly regurgitate, it is important to remove any beak restraint before transporting the animal, to prevent aspiration.5 The transportation carrier must be large enough to prevent damage to the flight and tail feathers.

GAMEBIRDS

RAPTORS

Gamebirds are notorious for biting and scratching. These birds are generally good runners and will often run toward the thickest cover available, making capture difficult. Because gamebirds are generally high-strung, they may be injured by the inexperienced handler. Several people may be needed to flush a bird under cover. A net or towel is preferred for capturing these animals. Once the bird is captured, it is best to keep the box covered (darkened) to minimize the stress associated with transport.

Although the beaks of raptors may look formidable, the biggest threat associated with these birds is their talons. Thick leather gloves, such as welding gloves, should be worn when handling medium to larger-sized specimens. Small raptors may be handled using outdoor work gloves (e.g., cotton or leather). When approached, most raptors roll backward and present their talons. When presented with a bird in this position, it is possible to offer them a towel or blanket to grasp before restraining the legs. The legs should always be grasped as close to the body wall as possible, as this will reduce the likelihood of causing a tibiotarsal fracture or injuries associated with selftaloning. The head of the raptor should be grasped using the same techniques to restrain other birds, with the index finger and thumb positioned under the mandible (Figure 19-6). For animals that are free-ranging, a bal chatri trap may be used. These traps can be built out of wire mesh (e.g., chicken wire) and fishing line. The wire mesh is first used to construct a wire cage to hold a live prey species (e.g., pigeon or rodent). A second piece of chicken wire is then placed over the prey cage, thus preventing injury to the prey species. The fishing line is then cut into small pieces (6-8 cm) and tied into small nooses. These nooses are secured over the entire surface of the trap. When the raptor attacks the trap, it becomes entangled in the trap and cannot lift off the ground with the trap.

Avian Species SMALL AND MEDIUM-SIZED BIRDS

WATERFOWL Waterfowl, including ducks, geese, and swans, can deliver very painful bites and scratches. In addition, swans can also cause injury by striking with their wings. During the capture process, it is important to prevent these animals from reentering the water. Some form of water transportation and expert assistance may be necessary if the birds are in the water. Specialized equipment, such as swan hooks and bags, may also aid in capture and transport. The easiest method for transporting these animals is to place them in a transport carrier or blanket or sack.

WADERS Many of the wading species have razor-sharp pointed beaks, and they will use them to stab at the faces of any perceived predators (including humans). Appropriate eye protection, such as safety goggles, should always be worn when handling these birds. Capture is similar to that of waterfowl. Transport boxes should be tall enough that the bird can stand comfortably. If such a box is not available, short trips can be made with the bird wrapped in a towel or blanket and the legs

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Mammals Unconscious mammals must be approached and handled with caution, as they may suddenly regain consciousness. Mammals should only be carried in an appropriate transport container and should never be captured or handled with

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in a net and then pinned using a soft-headed broom. Once pinned safely, they can be scruffed. If this method does not work, a quick-release rabies pole can be used. It is preferable to use one with a rubber end to prevent fracturing an animal’s teeth as is bites the pole.7 Inexperienced individuals should not attempt to capture these animals, as they are likely to injure both the animal and themselves. A live animal trap, plastic domestic pet travel carrier, or sturdy plastic trash can with a tight-fitting lid can be used to transport mesopredators.

DEER

Figure 19-6 The talons and head of a raptor should be securely restrained to prevent injury to the handler.

unprotected hands. For capture, a towel or blanket may be placed over the animal to reduce its vision. Large and potentially dangerous mammals should always be sedated before being transported.

RODENTS These animals frequently bite when handled. Leather gloves should always be worn by the individual capturing the animal to reduce the risk of injury and disease transmission. Small wild mammals may be caught with small nets or by covering them with a towel or blanket. Transport boxes for these animals should be escape-proof. Because of their ability to chew through cardboard, sturdy plastic or fine-meshed wire carriers are strongly recommended.

LAGOMORPHS Although considered docile by many, lagomorphs may bite and scratch during capture and handling. Leather gloves should be worn to reduce the risk of injury. Rabbits and hares are more susceptible to the effects of stress than most other species. The short-term effects associated with stress may be manifested in sudden death due to heart failure or fatal oliguria.6 Over longer periods of time, stress may be manifested as reduced gut motility and the disrupted carbohydrate metabolism, leading to diarrhea, hepatic lipidosis, liver failure, and death. In these cases, the cecal microflora are also disrupted, leading to enterotoxemia and gut stasis. Lagomorphs are also prone to spinal injury during capture and handling.6 Blankets, towels, or nets may be used to capture animals that are free-ranging.

MESOPREDATORS: OPOSSUMS, RACCOONS, AND FOXES Mesopredators have very sharp teeth and can deliver a painful bite, even through leather gloves. Animals that are severely ill or in shock may appear tame but can still inflict a serious injury. It is important to know that opossums can climb upward when held by their tail. Mesopredators may be caught

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Yearling and adult deer can pose a special danger. These animals are capable of gouging with their antlers or kicking with their feet. Only trained professionals should attempt to capture an injured subadult or adult deer. A deer that is severely injured or in shock may appear tame when approached; however, they are still capable of quick bursts of speed and energy and can injure an inexperienced handler. Before approaching an injured deer, it should be evaluated from a distance to determine its general status. The equipment generally used to capture deer include dark towels or drapes for covering the head, thick blankets, soft ropes, soft cargo or freight netting, and possibly a dart gun or pole-syringe.4 Anesthetics appropriate for sedating deer and a euthanasia solution should be kept on hand. If the deer is trapped in fencing or some other physical structure, it should be sedated remotely to prevent further injury to the deer. Expert assistance should also be sought for transport of injured deer. Individual deer should never be transported loose within a trailer without restraint. Large deer can best be transported wrapped up in cargo nets with the eyes blindfolded and the feet hobbled. Small deer can be wrapped in blankets. Trussed-up deer can be transported in a vehicle or trailer. Fawns should be captured by hand and restrained manually. The deer should be approached from behind, and if recumbent, covered with a blanket, net, or coat. Many fawns do not display any resistance to capture. Once restrained, the animal should be blindfolded to reduce its stress levels. Once masked, these deer will lie quietly if they are undisturbed. The animal’s legs can be tied together at the level of the metacarpi and metatarsi (cannons) using a soft rope; however, hobbling an animal in this way must be done carefully so that the soft ropes are not too tight. To prevent further struggling, the deer should then be trussed up in a blanket or cargo net and secured with additional ropes. Some deer are very vocal when caught, injured, or handled.

Reptiles and Amphibians Expert assistance should be sought before attempting to capture and transport venomous snakes. For nonvenomous snakes and other reptiles, a cloth or canvas bag placed in a solid box makes an ideal transportation container. Amphibians can be moved in a similar manner, although the transport bag should be kept moist to prevent desiccation. This can be achieved by placing damp vegetation, such as sphagnum moss, in the bag or frequently misting the bag with dechlorinated water.

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RESTRAINT AND HANDLING Manual Once the animal is transported to the clinic, the veterinary staff should be prepared to restrain and handle the animal for the physical examination and any diagnostic testing procedures. It is essential that the staff be trained properly to minimize the likelihood of injury to the human handler and patient. Restraint and handling is a stressful time for the patient, often leading to elevated cortisol or corticosterone production. Over time, this physiologic stress could have detrimental effects on the patient, including a reduction in metabolism, immune function, and normal behaviors.

GENERAL AVIAN For most simple procedures, physical restraint alone can be used. Most species are easier to catch and handle in low light conditions and can be calmed by covering their head. At our hospitals, a towel is often used to support the body, restrain the wings, and cover the head. If the animal is not stable enough for chemical anesthesia during radiography, tape and a plexiglass board can be used to restrain the animals. In all cases of restraint, special care must be taken to ensure that the animal can have normal keel excursions that allow for adequate ventilation, as birds lack a diaphragm. Most birds will attempt to beat their wings during restraint, and this must be prevented for several reasons: They (1) risk damaging their flight and tail feathers, (2) can fracture the bones of their wings, (3) can worsen any injury, and (4) risk expending valuable energy in the process. When the wings are immobilized, they should be placed into their normal resting position.

Passerines The simplest and most effective method of restraint is to place the bird in one hand with the dorsum of the bird in the palm and the neck held gently between the forefinger and second finger. It is especially important in the handling of small birds that undue pressure is not exerted that may inhibit respiration.

Pigeons and doves Smaller species can be held with the keel resting in the palm and the head toward the wrist, using the fingers to secure the feet, tail, and wings.

Gamebirds Smaller gamebirds may be held using the same technique described previously for small passerines; however, because larger gamebirds are both muscular and powerful, they must be held using two hands. When handling these animals, care must be taken to prevent them from losing excessive amounts of feathers, as they are “loosely feathered” and feather loss may delay release. Wrapping these birds in a towel will minimize wing-flapping and feather loss. Sufficient space should be maintained between the legs, as these animals can injure themselves by kicking their legs. Male gamebirds may have danger-

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ous spurs on their legs, and care should be taken to secure the legs firmly to avoid injury to the handler or bird.

Waterfowl Smaller waterfowl can be restrained by folding the wings against the body and then holding both the wings and the body in both hands. A wing hold can also be used by grasping the wings near the point of the shoulders, with the index finger between the wings and the other hand supporting the sternum. The wing hold can be used only for short periods of time; otherwise, brachial plexus paralysis may result. Larger waterfowl can be restrained by tucking the body under the arm and against the restrainer’s body. The bird’s legs can also be tucked up against the body and the beak held with the free hand. For larger waterfowl, multiple individuals may be needed to properly restrain a patient. Both the handler and the animal may incur serious wounds by the calloused carpal joints of waterfowl. Appropriately sized towels or commercial body wraps can be used to limit wing flapping. The ends of the towel can be wrapped diagonally across the bird’s body, leaving the head and neck exposed. The towel can be held in place with tape or bandage material. In those species without external nares, it is important to allow the beak to remain slightly open to ensure adequate ventilation.

Waders A variety of challenges are present in the restraint and handling of waders. The smaller species are very delicate, and damage to the extremities and beak tips can easily be incurred. If waders are restrained with their legs flexed, permanent paralysis may occur due to impeded circulation. Some waders, such as herons and egrets, have very sharp beaks, and will make lightening-fast stabbing movements at the face and eyes of their captors. Safety goggles should be worn at all times when handling these birds. The head should be secured first. As with other birds, placement of the index finger and thumb under the mandible will provide adequate head restraint. The body can be restrained using the techniques described for larger waterfowl. Rubber tubing can be used to keep the beak closed during examinations, but it should never be left on while the bird is unattended, as regurgitation and aspiration may occur.2 Impaling a cork on the end of the beak has also been used as a safety precaution, but we do not recommend it.

Seabirds Seabirds can also deliver dangerous blows to the face and body with their beak and should therefore be restrained using the techniques described for waders.

Raptors Raptors can pose a serious injury risk to any handler, experienced or inexperienced. Therefore, it is important to always remain vigilant around these animals. Raptors will use two primary defenses against their human handlers: their talons and their beak. To protect against injury, leather gloves should always be worn. When approached, raptors generally roll into a dorsal recumbent position and present their talons. The legs

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506 should be grasped as close to the body as possible to prevent self-taloning and iatrogenic tibiotarsal fracture. Once the legs are secured, the wings should be secured against the body to prevent feather damage. A towel or body wrap can be used to maintain the wings against the body. Always restrain these animals in an open area in a room, as the animal’s feathers may become damaged if the wings strike a counter or table surface. The head of the raptor should be grasped using the same techniques described to restrain other birds, with the index finger and thumb positioned under the mandible. In our experience, owls, eagles, and accipiters are the most prone to bite. Wearing leather gloves will protect the handler against bite injuries. Most medium to large raptors require a minimum of two people to safely restrain them.

MAMMALS Squirrels Adult squirrels can inflict deep bite wounds using their incisors. Because of this, it is important to always restrain these animals while wearing a pair of thick leather gloves. The index finger and thumb of one hand can be placed underneath the mandible to control the head, while the free hand can be used to support the body. We strongly recommend wearing gloves when handling these animals, because if they bite a member of the staff they need to be, by law, euthanized. Orphaned squirrels can generally be handled using a single hand.

Bats Injured bats pose a special risk to veterinarians and their staff. In most states, these animals remain an important vector of the rabies virus. Because these animals have special requirements regarding their care, they should only be handled by individuals with a significant amount of chiropteran experience. When working with these animals, gloves should be worn at all times. Bats are escape artists, and great care must be taken during handling to prevent accidental escape. One method of restraint is to position the bat dorsoventrally across its thorax and use the thumb and forefingers in apposition.8 Care should be taken not to restrict the animal’s ability to breathe. Scruffing appears to be stressful to many chiropteran species and should be avoided. It is important to always support the body of a bat while handling its wings to prevent iatrogenic fractures of the appendicular skeleton.

MANUAL OF EXOTIC PET PRACTICE

supported, one quick kick of the rear legs could result in a vertebral fracture. Rabbits can also be restrained by wrapping a towel around the body of the animal (“bunny burrito”), leaving the head exposed. If this technique is used, again, it is important to support the hindquarters of the animal. Another method to assist in the restraint of rabbits is that of “hypnosis,” or “trancing.”6 The technique is described as an immobility response that occurs after placing the animal on its dorsum, resulting in a cessation of spontaneous movement and a failure to respond to external stimuli for several minutes.

Mesopredators Restraining adult mesopredators requires some background knowledge of these animals and is generally best done under sedation. If these animals are presented in a cage or carrier, we prefer to anesthetize them for removal from the cage: Ketamine (5-10 mg/kg IM) (Ketaset, Ft. Dodge Animal Health, Ft. Dodge, IA) or Telazol (tiletamine-zolazepam; 3-5 mg/kg IM) (Ft. Dodge Animal Health). Some individuals recommend removing the animals using a rabies pole, but this method is very stressful to the animal. Once anesthetized, the animals can be examined and sampled for various diagnostic tests. The animals can be moved by scruffing them at the nape and supporting the body with the free hand. For larger specimens, two or more handlers are recommended. Juvenile mesopredators can be restrained manually. Leather gloves should be used to reduce the risk of bite injuries. The juveniles can be handled using the same technique described for the adults. The tail of the opossum can also be used when manipulating or moving an opossum, although we do not believe the entire weight of the animal should be supported by the tail.

Deer Adult deer should be anesthetized for a physical examination to minimize the stress and risk of injury to the patient and the handler. Anesthetic agents commonly used for deer are found in Table 19-4. Fawns can be restrained by supporting the head and body. Placing a hand under the thorax or abdomen when the animal is in a standing position will minimize the likelihood of the animal falling into a down position. In some cases, however, it is easier to examine a fawn while it is lying down.

Lagomorphs

Reptiles

Finding a balance between gentle and secure restraint is essential when handling lagomorphs. These animals may react to any perceived danger by either becoming motionless or fleeing. In those cases where they remain motionless for an extended period of time, the handler should be prepared for them to mount a sudden escape response. Rabbits can be restrained by grasping (scruffing) the skin over the nape of the neck with one hand and using the free hand to support the hindquarters. The skeleton of these animals is lighter than comparable mammals, whereas the musculature associated with the rearlimbs is massive and strong. If the hindquarters are not

Reptiles should be handled carefully to prevent injury to the patient. These animals have only a single occipital condyle supporting the skull and cervical spine, so the head and neck should always be supported to ensure that the cervical spine is not bearing the weight of the animal’s body. It should also be taken into account that during times of shedding, reptiles are more susceptible to skin damage. In the past, hypothermia was considered an appropriate manner of restraint for reptiles; however, this primitive technique is no longer considered appropriate. There are several possible disadvantages associated with this technique: (1) There

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Chapter 19

Figure 19-7 Chelonians can be securely restrained by grasping the lateral surfaces of the plastron and carapace.

is no published evidence that suggests that lowering a reptile’s body temperature until it no longer responds to external stimuli abolishes the perception of pain, (2) as the animal warms it may suddenly awaken, which can cause danger to the handlers if it is a hazardous species, (3) the animal may die if the hypothermic state is extended, (4) stress and immunodeficiency may result from hypothermic states because immunoglobulin synthesis is temperature dependent, and (5) several safe chemical restraint agents exist for those times when manual restraint is not considered sufficient.9 Many noninvasive, short procedures can be done using manual restraint alone, but it should be remembered that although reptiles may not react to painful or stressful stimuli in a manner similar to mammals, they do feel pain and experience stress. Confinement and reduction of vision can minimize the stress experienced by the animal. For example, a reptile can be placed into a dark bag for radiographs because they will generally remain motionless for the procedure. Placing a nonocclusive bandage material over the animal’s eyes may also be used to reduce motion for a diagnostic procedure such as radiographs. Chelonians can be restrained by grasping the lateral surfaces of the shell in the area of the bridges (Figure 19-7). For chelonians that are capable of extending their neck and biting (e.g., snapping turtle: Chelydra serpentina), grasping the shell more caudally is recommended. When the head of a chelonian needs to be restrained, the index finger and thumb can be placed behind the points of the ramus of the mandible. It is important that the handler does not exert excessive pressure when extracting the head and neck, as this can lead to cervical injury. For those species that can completely enclose themselves within their shell (e.g., box turtle: Terrapene carolina), chemical anesthesia is recommended. Placing the animal in a shallow volume of water may stimulate them to exit their shell, but waiting for this to occur can be time consuming.

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Small lizards may be held in one hand, with the index finger under the throat and the thumb on the head or neck. Larger lizards can be held by grasping them behind the head with an index finger and thumb, while securing the rear legs against the base of the tail. It is important to be gentle when manipulating the tail of a lizard capable of tail autonomy. Applying gentle pressure on the eyes of some species of lizards may induce a vagal response (e.g., bradycardia, lethargy). Large snakes and venomous snakes require at least two experienced handlers for restraint. Specialized equipment is recommended to manage these animals, including snake hooks, clear tubes, and pinning sticks. The snake can be placed on a foam mat, and a rod or hook may be used to pin the head. Once pinned, the head should be grasped behind the angles of the jaw with the thumb and forefinger, taking care not to apply too much pressure. A second person should assist the primary handler by supporting the body of the snake. Clear plastic tubes, either commercial or homemade and slightly larger in diameter than the snake, can be used to restrain venomous snakes. The snake should be placed on the ground with the tube directly in front of it, and the snake encouraged to enter the tube. When one third of the snake is in the tube, the snake and tube are grasped together to immobilize the snake. Preplaced holes in the tube can be used to allow for sampling and injections.

AMPHIBIANS Restraint and handling of amphibians should be kept to a minimum. Because amphibians are susceptible to skin damage and desiccation, they should be handled with moistened gloves. Larval amphibians can succumb to hypoxia if they are removed from their aquatic habitat for an extended period, and they should only be removed from the water for short periods of time. Some toad species, such as the marine toad (Bufo marinus), can present a danger to the handler by releasing bufotoxin from the parotid glands. Wearing exam gloves can minimize the likelihood of toxin exposure to the handler. Anurans (frogs and toads) can be restrained by placing an index finger and thumb into the axillae or wrapping these fingers around the forelimbs. For small specimens, cupping them in a hand is also recommended. Most salamanders and newts require minimal restraint and can be held in an open hand. The head of caecilians and amphiumas should be restrained because they can bite.

Chemical While many procedures can be carried out with manual restraint, there will be instances where the patient is too difficult to restrain, the procedure too painful, or the stress associated with the procedure too great for manual restraint to be used alone. When this is the case, it is advisable to use chemical restraint. Although the expense associated with managing the case using anesthesia is greater, the reduced stress and time saved to try to carry out the procedure will outweigh

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this disadvantage. Both injectable and inhalant anesthetics are available for use in a multitude of wildlife patients, and a list of commonly used anesthetics can be found in Table 19-4. When selecting an anesthetic(s) for a wildlife case, it is important to consider the patient’s status and the procedure being performed. Some of the points to consider include the the animal’s weight, age, reproductive status, stress level, type of injury, length of procedure, and need for analgesia.

TABLE 19-4

PHYSICAL EXAMINATION AND COMMON ABNORMALITIES The overall aim of wildlife rehabilitation is to return animals to the wild in a condition that allows them to compete with members of its own and other species and to successfully reproduce and behave in a species-appropriate way, with no physical or mental impediments that would affect their chances of performing normally in their environment. The physical

Anesthetic Agents Used to Anesthetize Wildlife Patients

Agent

Dose

Comments

3%-5% induction, 1%-3% maintenance 5%-8% induction, 2%-5% maintenance 10 mg/kg IV, to effect

Safe, mask or intubate, low cardiopulmonary risk Safe, mask or intubate, low cardiopulmonary risk Must be given IV or IO, rapid recovery Not recommended, rough recovery Not recommended, rough recovery

3%-5% induction, 1%-3% maintenance 5%-8% induction, 2%-5% maintenance 5-10 mg/kg IV, to effect 1-5 mg/kg 0.15-0.25 mg/kg 0.5-0.1 mg/kg 5-10 mg/kg 10-40 mg/kg 80-100 mg/kg 5-15 mg/kg 3-5 mg/kg 5 mg/kg

Safe, mask or intubate, low cardiopulmonary risk Safe, mask or intubate, low cardiopulmonary risk Must be given IV or IO, rapid recovery Deer (combine with ketamine)

Isoflurane Sevoflurane Propofol Medetomidine

3%-5% induction, 1%-3% maintenance 5%-8% induction, 2%-5% maintenance 10-15 mg/kg IV, to effect 0.15 mg/kg IM 0.75-0.15 mg/kg IM

Ketamine

5-15 mg/kg 10-25 mg/kg

Telazol

3-5 mg/kg 5-8 mg/kg 5-10 mg/kg

Safe, mask or intubate, low cardiopulmonary risk Safe, mask or intubate, low cardiopulmonary risk Must be given IV or IO, rapid recovery Alligators: reverse atipamezole (equal volume) Chelonians: in combination with ketamine (5-15 mg/kg IM) Snakes, lizards: induction Chelonians: induction; not recommended for surgical procedures without additional coverage (inhalant, medetomidine) Snakes: induction Lizards: induction Chelonians: induction

200-5000 mg/L 300-350 mg/L 400-450 mg/L 35-45 mg/kg ICe 5-15 mg/kg IV

Lower doses for larvae Anurans Salamanders Salamanders All

Avian Isoflurane Sevoflurane Propofol Xylazine Ketamine Mammals Isoflurane Sevoflurane Propofol Xylazine Medetomidine Ketamine

Tiletamine-Zolazepam

Deer (combine with ketamine) Mesopredators Lagomorphs Rodents Deer (combine with xylazine) Deer (combine with medetomidine) Mesopredators; not recommended for lagomorphs

Reptiles

Amphibians MS-222 Clove oil Propofol

MS-222, tricaine methane sulfonate; ICe, intracoelomic; IM, intramuscular; IV, intravenous.

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examination represents the first opportunity the veterinarian has at determining the likelihood an animal can be rehabilitated and released. In some cases, the injury will be lifethreatening or not correctable, and humane euthanasia will be warranted. In other cases, the ultimate outcome will be based on the veterinarian’s ability to provide medical and surgical assistance. For these reasons, it is important that the veterinarian fully evaluate these animals at the time of presentation to ensure that he or she identifies all of the problems an animal presents with, as well as make a determination as to the value of pursuing or not pursuing a case. The first, and most important, step in performing a physical examination is to correctly identify the presenting species and its age group. For those with limited knowledge regarding species characterization, we recommend maintaining a library of field guides (both national and state) to assist with species identification. Maintaining friendships with local ornithologists, mammalogists, and herpetologists might also be helpful when attempting to identify resources to assist with species identification. The Internet can also be a valuable resource for identifying species too. If the identification of the actual species is not possible, it would still be useful to be able to characterize the animal to family or generic level. Correctly identifying the animal will allow the veterinarian to determine the (1) natural diet (e.g., carnivore, omnivore, herbivore), (2) habitat and feeding strategy (e.g., aerial feeder, arboreal, ground dwelling), (3) social behavior (e.g., gregarious and nonterritorial, solitary and territorial), (4) movements (e.g., resident, partial migrant, summer or winter visitor), (5) approximate age (e.g., dependent/independent juvenile, or adult) (Figure 19-8), and (6) gender. This information will aid in determining the care an animal should receive and may also determine its release potential. If the animal is being presented from an outside source, a thorough history should be obtained. Unfortunately, the

Figure 19-8 It is important for veterinarians to be able to identify the general age of their patients. In the case of this greathorned owl, the downy feathers on the head confirm that it is a juvenile.

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opportunity to obtain a thorough history from these animals is often limited because the individuals presenting the animals may have only recently encountered the animals. Those historical questions that can generally be answered include: (1) Where was the animal found? (2) Under what conditions was the animal found? (3) What day and time was the animal found? (4) How long has the rescuer had the animal? (5) Has the rescuer fed, watered, or medicated the animal? (6) What has the animal been doing while in captivity? (7) Were there any obvious lesions on the animal? Once an adequate history is obtained, the animal should be observed from a distance to gauge the severity of the illness and determine whether to proceed with the physical exam or allow the animal to recuperate. Avian and reptilian species will attempt to mask any signs of illness when a perceived predator approaches. If the animal is unable to do this, and appears obviously ill, the condition is likely extremely serious. During the observation period, information can be gathered about the animal’s posture, conformation, respiratory and neurologic status, and gastrointestinal output (e.g., feces, vomitus). As with the physical examination of companion animals, it is extremely important to follow the same routine for each examination so that no body system is missed. An example of a standard wildlife physical examination can be found in Box 19-3, and a list of the most common findings during an examination in Table 19-5.

Avian General Examination Birds that present in shock should be stabilized. Overmanipulating these animals can result in acute cardiopulmonary arrest. Once the bird is stable, a thorough and systematic approach to the physical examination is essential. In addition to the previously described exam protocol, an examination of the pectoral area should be done. Evaluating the condition of the large pectoralis muscles lying over the keel will aid in the assessment of the animal’s body condition. A bird can be presumed to be in good body condition when the muscles form an “A” off of the keel. If the pectoralis muscles are concave and the keel is prominent, then the animal is in poor condition. This is generally indicative of chronic disease.10 An animal that has bulging pectoralis muscles, making the palpation of the keel difficult, can be presumed to be overconditioned. A distended abdominal wall (coelomic cavity) may be indicative of ascites, neoplasia, egg binding peritonitis, bowel distension, or hepatomegaly. In all but laying hens, the abdominal muscles associated with the coelomic cavity should be slightly concave. Both wings and legs should be extended and examined. Feather quality, muscle mass, and bone integrity should be assessed. Special care should be taken to evaluate joint function. The most common reasons that wild birds are presented to veterinary hospitals are traumatic injuries, poisonings, and infectious diseases. Traumatic injuries most commonly result from collisions with vehicles or windows, predation by domestic animals (e.g., dogs and cats), or gunshot wounds. Traumatic injuries frequently result in fractures, soft tissue injury, ruptured air sacs, nervous system damage (e.g., con-

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BOX 19-3

MANUAL OF EXOTIC PET PRACTICE

Standard Protocol for Performing a Physical Examination on a Wildlife Patient

1. Identify species to determine diet, housing needs, and restraint techniques. A geographically appropriate field guide is essential. 2. Observe the animal from a distance for alertness, aggression, mental status, body condition, and gastrointestinal and respiratory functions. 3. Restrain animal using appropriate techniques for that species. 4. Observe the eyes for pupil size and symmetry, pupillary light responses, menace reflexes, corneal disease, and the presence of hyphema or hypopyon. Perform fundic examination. 5. Examine the ears for injury and ectoparasites. 6. Examine the mouth for abscesses, mucous membrane tackiness, capillary refill time, and presence of parasitism. 7. Examine the nares for discharge. 8. Palpate the head for traumatic injuries or abnormal masses. 9. Palpate the lymph nodes (mammals only). 10. Thoroughly inspect the integument for signs of dermatitis, ectoparasitism, and abnormal masses. 11. Palpate the abdomen or coelomic cavity. 12. Palpate the extremities for obvious injury, asymmetry, muscle atrophy, and loss of feathers or fur. 13. Check anus or vent for abnormalities, diarrhea, or hemorrhage. 14. Auscult the heart and lungs. 15. Weigh the animal.

cussion, paralysis), and damaged plumage and beaks. Poisonings may be intentional or unintentional (Figure 19-9). It is important to attempt to identify the source of the poisoning, if at all possible, to prevent exposure to children, domestic pets, and other wildlife. Rodenticides, insecticides, herbicides, and heavy metals are the most common poisons reported in wildlife. Infectious disease in wild birds may be associated with an individual bird’s exposure and immune status, or it may be related to a disease outbreak. Chlamydophila psittaci, poxviruses, mycoplasmosis, and trichomoniasis represent diseases that commonly circulate through wild bird populations, while aspergillosis is more often an individual bird disease that occurs as a result of a single direct exposure or the stress associated with captivity. Endoparasites and ectoparasites are frequently found in wild birds, but they are rarely the primary cause of disease.

PASSERINES Small passerines commonly present as a direct result of a traumatic injury associated with predation or an in-flight collision (e.g., window, automobile). Soft tissue and orthopedic injuries are the most common result. If the trauma results in closed fractures of the ulna, radius, or carpometacarpus, the fracture(s)

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Figure 19-9 This great blue heron presented after falling into an industrial pond full of liquefied plastic.

may be stabilized by taping the flight feathers together and supporting the wing. Because of their high metabolism, passerines can potentially heal these fractures in 2 to 4 weeks. Unfortunately, fractures of the humerus or fractures of both the radius and ulna have a guarded to grave prognosis. If healing does occur with these fractures, normal flight is often compromised. Closed fractures of the femur with the fragments in near alignment will usually heal with cage rest; however, if the fracture occurs distal to the femur, light splints can be placed to align fracture segments. Small passerines will generally tolerate supporting their weight on one leg while the other heals. There is some debate about the long-term survival rates of one-legged small birds in the wild. Cat predation of passerines is a major concern in the United States. It has been estimated that these animals are responsible for killing millions of birds every year. In our practices, fledglings are the most commonly presented. The severity of wounds will vary greatly, from minor soft-tissue wounds to severe lacerations, punctures, and fractures. In most cases, feather loss is apparent as well. The wounds are often contaminated with mixed Gram-positive (e.g., Staphylococcus spp. and Streptococcus spp.) and Gram-negative bacteria (e.g., Pasteurella multocida and E. coli). Euthanasia is appropriate in severe cases. Superficial wounds carry a better prognosis and can be managed with appropriate antibiotic therapy and wound management. Passerines are also susceptible to the previously mentioned infectious diseases, especially poxvirus, mycoplasmosis, and ectoparasites. A higher prevalence of disease may be expected in these birds because they often congregate at high densities in feeding stations (e.g., suburban bird feeders). Starlings and members of the Corvidae family are susceptible to infection by the nematode Syngamus trachea (gapeworm), which can cause severe pulmonary disease and respiratory distress. Anesthesia may be required to facilitate an examination for the following circumstances: nervous or stressed birds, fractious birds, dyspneic birds (e.g., allows direct delivery of oxygen

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TABLE 19-5

Common Species Presentations for North American Wildlife

Species Squirrels

Bats

Rabbits and hares

Otters

Foxes, raccoons, coyotes

Opossums

Deer

Birds

Reptiles

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Common presentations Orphaned: fallen out of nest Traumatic injuries: automobile, domestic predators Viral disease: West Nile virus, papilloma virus Bacterial disease: Gram-negative opportunists Ectoparasites: mites, ticks, fleas Endoparasites: coccidians, nematodes Traumatic: fractures, wing tears, entanglement Poisoning: rodenticides, oil contamination Viral disease: rabies virus Bacterial disease Fungal disease Ectoparasites Endoparasites8 Orphaned: abandoned and kidnapped Traumatic injuries: automobile, domestic predators Ectoparasites: mites, fleas Endoparasites: coccidians, nematodes Colic: Bacterial dysbiosis Orphaned: parent trapped/killed Traumatic injuries: automobile, domestic predators Bacterial disease: Gram-negative opportunists, septic bite wounds Poisoning Ectoparasites: mites, fleas Endoparasites: protozoa, cestodes, nematodes Orphaned: parent trapped/killed Traumatic injuries: automobile, domestic predators Viral disease: canine distemper virus, parvovirus, rabies Bacterial disease: Gram-negative opportunists, leptospirosis Poisoning Ectoparasites: mites, ticks, fleas Endoparasites: coccidians, nematodes Orphaned: parent trapped/killed Poisoning Traumatic injuries: automobile, domestic predators Bacterial disease: Gram-negative opportunists, leptospirosis Ectoparasites: mites, ticks, fleas Endoparasites: coccidians, nematodes Orphaned: kidnapped Traumatic injuries: automobile, domestic predators, fire ants Viral disease: epizootic hemorrhagic disease, rotavirus, coronavirus Bacterial disease: Gram-negative opportunists (E. coli) Ectoparasites: mites, ticks, fleas Endoparasites: coccidians, nematodes Orphaned: fallen from nest, parent killed Traumatic injuries: automobile, domestic predators, electric wires, fishing hooks Poisoning: heavy metal, mercury, organophosphates Viral disease: West Nile virus, paramyxovirus, herpesvirus, poxvirus Bacterial disease: Gram-negative opportunists Ectoparasites: mites, ticks, lice Endoparasites: coccidians, nematodes Traumatic injuries: automobile, domestic predator, lawn equipment Viral disease: herpesvirus, iridovirus, paramyxovirus Bacterial disease: Gram-negative opportunists, mycoplasmosis Ectoparasites: mites, ticks Endoparasites: protozoa, nematodes

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512 and reduces stress), musculoskeletal examinations where there is a fracture, and an ophthalmic examination when a fundic exam is required. An ophthalmic examination should be done any time a bird suffers head trauma, as the disease of the posterior chamber is common in these birds.

PIGEONS AND DOVES As with passerines, pigeons and doves often present to veterinary hospitals after sustaining a traumatic injury. These birds rely heavily on flight to evade predators and reach roosts and should not be released with impaired flight. Euthanasia should be considered for birds with open or comminuted fractures or fractures with extensive joint involvement. Neurologic disorders are commonly seen with traumatic injuries, and the physical examination should include a thorough examination of the animal’s posture, demeanor, mentation, and reflexes. Traumatic wounds secondary to strike by raptors usually present as wounds on the bird’s dorsum. Ruptured crops are commonly seen in these species, likely because their crops are usually more heavily filled than those of other avian species. These cases will present as a necrotic granulating wound and associated crop fistula. Pigeons and doves are susceptible to a range of infectious diseases. Trichomoniasis commonly causes white plaques in the oral cavity. This organism is extremely contagious in these birds. Mild cases can be simple to treat, but advanced cases carry a poor prognosis. Many of the severe cases show some improvement with treatment, only to succumb before the completion of the treatment and elimination of the signs. For this reason, we generally recommend euthanasia for the severe cases. Bacterial and fungal infections of the crop (ingluvitis) are common, especially in young birds. A foul smell in the oral cavity can be an indication of a crop infection. The crop itself may appear thickened, enlarged, and have a decreased emptying time. Septic arthritis, often the result of infection with Salmonella sp. or E. coli, is common in these birds and may resolve with broad-spectrum antibiotic therapy; however, some authors advise against treatment because of the chronic joint problems that may arise after treatment and the zoonotic potential for the handlers.1 Pigeons and doves can serve as reservoirs for pathogens that are highly infectious to other birds, including paramyxovirus (Newcastle disease—a reportable disease) and poxvirus. Pigeons or doves presenting with clinical signs consistent with these diseases should be euthanized to minimize the likelihood of disease transmission to other birds. A bird with a paramyxovirus infection may display a range of clinical signs, including head tremors, diarrhea, deformed eggs, torticollis, polyuria, polydipsia, and anorexia. Birds with pox lesions generally have multiple masses in their featherless areas (e.g., legs and surrounding the oral cavity and eyes). When these lesions are draining, they are contagious to other birds. Mycoplasma spp. infections are commonly associated with upper respiratory tract disease, resulting in swollen sinuses and ocular or nasal discharge. Other than ticks or lice, external parasites are rarely seen. Various species of coccidians can cause wasting and diarrhea. Pigeons and doves are commonly infected with Capillaria spp. and ascarids.

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GAMEBIRDS Common reasons for presentation include trauma (e.g., hit by automobile) and infectious disease. Gamebirds who have been traumatized may present with orthopedic injuries, ocular hemorrhage, or neurologic disease. Infectious diseases seen in gamebirds include viral disease, bacterial infection, fungal disease, and parasitic disease. Paramyxovirus (Newcastle disease) and pheasant coronavirus, which causes multiple masses around the head and limbs, are the most common viral diseases. These birds are susceptible to a variety of opportunistic bacterial pathogens. Some of the more common pathogens are E. coli, Salmonella sp., and Mycobacterium avium. Birds with E. coli infections generally present in critical condition and often appear hypothermic (fluffed) and depressed. Salmonella sp. is usually associated with hemorrhagic enteritis, and animals often succumb to this disease acutely. Mycobacteriosis is generally associated with general body wasting, sinusitis, and tenosynovitis. This pathogen is zoonotic, so appropriate precautions should be taken when handling and disposing an animal. Fungal infections with Aspergillus spp. can result from spore inhalation, and affected animals develop clinical signs associated with the respiratory tract, including coughing and dyspnea. The parasitic diseases reported in gamebirds are similar to those seen in pigeons and doves. Capillaria sp. is a common finding in both wild and captive gamebirds. This endoparasite can cause severe loss of body condition, and it is not uncommon for animals to succumb to infection. Commonly, gamebirds are semi-intensively farmed before release and may be prone to parasitic infections. A soiled vent and diarrhea are common findings in gamebirds with endoparasitic infections. Ectoparasites such as feather lice, hippoboscid flies, and mites may be present in large numbers on debilitated birds.

WATERFOWL Waterfowl are commonly presented to the veterinary hospital because of traumatic injury (e.g., vehicle collision, fishing line injuries), poisoning (e.g., lead ingestion), and infectious disease. Traumatic injury often results in both soft tissue and orthopedic injuries. Loss of the use of one or both legs or feet, an inability to fly, and open fractures or fractures involving joints are all indications for euthanasia. Injury due to fishing line and hooks may be obvious at presentation, or it may be unknown until further diagnostics are pursued (e.g., radiographs). The ingestion of lead shot or lead sinkers has been a major problem for wild waterfowl. Lead toxicity can result in a range of clinical signs, including acute, asymptomatic death or bright green diarrhea, progressive weight loss, anorexia, anemia, kinking of the neck, neurologic symptoms, muscle weakness, dehydration, and poor feathering. Birds with a more chronic presentation may shows signs consistent with starvation and severe anemia. Lead becomes toxic to the waterfowl when the gastric acid ionizes the lead, allowing it to become absorbed and affect various organ systems (e.g., liver). Botulism (Clostridium botulinum) is a common seasonal problem in waterfowl. Affected animals present with a flaccid paralysis of the muscles, commonly known as limberneck. Three grades of paralysis have been described: (1) cannot fly but can walk and

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swim, (2) cannot fly or walk but can swim and move by a flopping motion when assisted by the wings and unable to lift head out of water or off the ground, and (3) almost completely paralyzed.11 The prognosis for grade 1 is generally fair to poor, whereas the prognoses for grades 2 and 3 are poor to guarded and grave, respectively. Oil contamination and blue green algae toxicity can also affect wild waterfowl. Infectious diseases of waterfowl are generally divided into two categories: individual and flock. Any individual suffering from stress (e.g., low food source, stress of predators, habitat loss) can develop opportunistic infections. Aspergillosis is an example of this. On the other hand, there are also those diseases that primarily affect entire flocks. Fortunately, many of these are rare. Duck viral enteritis, which is caused by a herpesvirus, can result in hemorrhagic gastroenteritis, neurologic abnormalities, polydipsia, and signs associated with all other body systems. Duck plague, Pasteurella multocida infections, can occur in flocks at certain times of the year. Avian tuberculosis can also be disseminated through flocks that are in close association.

WADERS The most common reasons that waders present to the veterinary hospital are trauma due to fishing lines and hooks, entanglement in telephone cables, poisoning by ingestion of mollusks that have accumulated the phytoplanktons that cause red tides, poor condition after hard seasons, and oil contamination. When working with wading birds, it is important to closely examine the commissures of the mouth and frenulum of the tongue for fishing line. Some waders, especially herons, have prominent keels even when they are considered to be in good health. It is important that these animals are not misdiagnosed as being in poor body condition and oversupplemented with supportive care.

SEABIRDS As with waders, it is important to carefully examine the oral cavity of seabirds for fishing lines or hooks. The body condition of these animals can be assessed by palpating pectoral muscle mass.5 The plumage should be closely inspected for signs of oil contamination. Common reasons that seabirds present to veterinary hospitals include fishing line and hook entanglement, wing and leg fractures, aspergillosis, and oil contamination. Euthanasia should be considered with complicated or open wing or leg fractures, or those with joint involvement. Animals that have heavy ectoparasite burdens or present in extremely poor condition often carry a guarded prognosis. When considering the long-term management of these animals in captivity, it is important to minimize stress and maximize nutritional support, as these animals generally do poorly in captivity.

RAPTORS The most common reasons that raptors present to a veterinary hospital are traumatic injuries (e.g., automobile, gunshot, entanglement in high wires), pesticide toxicity, infectious diseases, and maladapted starvation. Trauma may result in ocular,

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aural, orthopedic, and soft tissue injuries. Pesticide toxicity may be a result of intentional or accidental poisoning. Organophosphate toxicity is commonly seen. Viral, fungal, and bacterial infections can cause severe disease. Many raptors are extremely susceptible to West Nile virus infections. Falcon and owl herpesvirus infections are occasionally observed in animals with neurologic disease. Raptors are also susceptible to paramyxovirus. Many raptors become infected with poxvirus when they prey on Columbiformes (e.g., pigeons and doves). Because raptors ingest prey species, the potential for them to become exposed to a variety of opportunistic bacterial pathogens exists. E. coli, Salmonella spp., Klebsiella spp., Pseudomonas spp., and Staphylococcus aureus infections are commonly reported in raptors. Hippoboscid flies and lice are commonly found on raptors at the time of presentation. In addition to being exposed to poxvirus with columbiformes, many raptors are also exposed to trichomonads. Affected animals often have large white plaques in the oral cavity. Euthanasia should be considered for any of the following presentations: loss of an eye or reduced vision (owls may be an exception), an inability to feed naturally, loss of the function of one or both wings or legs, damage or amputation of the toes or talons that prevents normal hunting and perching, any bird that has become habituated or imprinted on humans and would be dangerous if released, and inability to relate naturally to conspecifics.

Mammalian General Examination The physical examination of all mammalian patients should be thorough and consistent; this will minimize the likelihood of missing species-based problems. Most orphaned mammals can be examined using standard restraint, whereas adult animals need to be anesthetized. We always recommend wearing gloves, a surgical mask, and goggles when working with rabies vector species. The general body condition of most mammals can be determined by evaluating the epaxial muscles along the spine and the large muscles along the long bones. The amount of subcutaneous adipose tissue, as determined by pinching the skin, can also be used to assess body condition.

SQUIRRELS The gender of an orphaned squirrel can be determined by identifying the prepuce of the male on the ventral abdomen or the elongate vulva of a female. Sexually mature squirrels are easy to confirm, as the males have large external testicles. Squirrels commonly present to the veterinary hospital for traumatic injuries (e.g., hit by automobile, gunshot, domestic species predation), poisonings, and infectious diseases. Animals that present for traumatic injuries may have unilateral or bilateral epistaxis, musculoskeletal injuries (e.g., fractures), deep puncture wounds, or degloving injuries, or they may be in shock. A decision as to the long-term potential for releasing the animal should be made at the time of presentation. Although some rehabilitators do not advocate this, at times a decision to not pursue a case in which the species is more plentiful (e.g., squirrel vs. bald eagle) must be made. This is the basis for triage, especially when financial and personnel resources are

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limited. Most squirrel poisonings are associated with rodenticides or pesticides and are, unfortunately, intentional. It is important to identify and pursue potential poison cases to minimize the risk for poisoning of children, domestic animals, or other wildlife. A poxvirus is occasionally observed in wild squirrel populations. Squirrels are susceptible to West Nile virus infections. Many of the infectious agents recovered from squirrels at the time of presentation are presumed to be related to a bite injury sustained from a predator. Because the predator (e.g., dog or cat) has a mixed microflora, a broadspectrum antibiotic should be used to manage the squirrel’s injury.

ceptible to a range of bacterial diseases, including pasteurellosis, which can colonize wounds or cause respiratory disease; infection with Treponema cuniculi, a spirochete that is sexually transmitted and causes lesions on the face, external genitalia, and perineum; infection with Yersinia pseudotuberculosis, which can cause internal abscesses and septicemia; and infection with Francisella tularemia, which causes an acute septicemic disease. Because some of these diseases are zoonotic, it is important that the veterinarian and staff wear gloves and take appropriate precautions. As with most mammals, lagomorphs are susceptible to ectoparasites and endoparasites.

BATS

The most common reasons that mesopredators (e.g., opossums, raccoons, foxes, coyotes) present to the veterinary hospital are traumatic injuries (e.g., hit by automobile, gunshot, trapping), intentional poisonings, and infectious diseases. Raccoons, opossums, foxes and coyotes are frequently the victims of gunshots or trapping attempts, which may result in severe orthopedic and soft-tissue injuries. They may also present with infected bite wounds from other animals. Poisoning is generally from organophosphates or vitamin K antagonist rodenticides (e.g., warfarin), although antifreeze poisonings are also occasionally reported. To characterize the type of poison, it is important to examine the animal closely. Animals with organophosphate toxicity often present with miotic pupils, excessive salivation, and ataxia. Animals poisoned with warfarin or another vitamin K antagonist, generally present with coagulopathy, hemorrhage from different orifices, weakness, and acute death. Animals with antifreeze toxicity generally succumb to renal failure. Treatment for organophosphates and vitamin K antagonists are supportive care and atropine/2-pam and vitamin K, respectively. Mesopredators are susceptible to a variety of viral, bacterial, fungal and parasitic diseases. Viral infections are common. Animals that are febrile (reference temperatures: opossum 92°95° F; raccoon, fox, coyote, 100°-104° F) should be quarantined and handled with care. Certain viral diseases may cause a lymphopenia or lymphocytosis. Raccoons, coyotes, and foxes are all susceptible to canine distemper, parvovirus, adenovirus, and rabies. Opossums should also be considered to be susceptible to rabies, although infection is rare. Sarcoptic mange and ringworm are also common in these animals and may cause extreme pruritus, dermatologic disease, and eventually emaciation and death. Some may not exhibit signs but instead serve as reservoirs for the mites. Because both sarcoptes and ringworm are zoonotic, the veterinary staff should wear gloves and change their laboratory coats (and clothes) in between handling these animals and other animals.

Species determination is critical, and field guides are highly recommended to assist with identification. Determining the animal’s gender and age is also important. Male bats have an obvious prepuce on the ventral abdomen and can easily be differentiated from a female. Juvenile bats are smaller than their adult counterparts, are dependent, and have underdeveloped wings. Once a bat becomes independent, aging becomes more difficult. Transillumination of the phalanges should reveal still developing bones, and a magnifying lens can be used to visualize small digits. Gloves should always be worn when handling bats. Body condition score can be used as a general health assessment; extremely underweight bats will have a concave abdomen. A very thorough examination of the wings should be performed. Even minor injuries will impair flight. The body should be supported while examining each wing. The bones, joints, and soft tissue elements of each wing should be examined for fractures, tears, and joint mobility. Common reasons that chiropterans present to veterinary hospitals are traumatic injuries (e.g., hit by automobile, gunshot), fishing line entanglement, oil contamination, poisoning, and neurologic symptoms secondary to lyssavirus infection. Again, because these animals are important rabies virus vectors throughout much of the United States, all animals should be considered positive until proven otherwise and should be handled accordingly.

LAGOMORPHS Gender determination of lagomorphs can be difficult in juvenile animals. In adult rabbits, an examination of the external genitalia will often reveal the gender. It is important to note that male rabbits do have open external rings, which allows them to draw their testicles into their body cavity. If the testicles have ascended into the abdomen, it should still be possible to extract the penis. Careful examination of the head and limbs, abdomen, and perineum should be performed on every lagomorph case. The most common reasons that rabbits present to a veterinary hospital are traumatic injury, fly strike, dental disorders, viral disease, bacterial disease, and parasitism. Traumatic injuries for presenting cases can range from minor soft tissue wounds to severe fractures or spinal injury. Fly strike may result from open wounds or viral disease. Myxomatosis is a common viral disease causing skin and respiratory disease. Rabbits are sus-

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DEER Adult deer presented to a veterinary hospital pose a special risk because injured deer will make every attempt to evade capture. In most cases, these animals must be sedated the entire time they are hospitalized, or they will worsen their injuries. For some facilities, euthanasia is considered the most humane option for these cases. Common reasons deer are presented to

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Wildlife

veterinary hospitals include traumatic injury (e.g., automobile, entanglement, dog attack, gunshot), myopathy, healthy fawns mistaken for orphans, and fawns suffering from starvation after hard winters. Traumatic injury commonly results in limb, pelvic, or facial fractures, or spinal injury. Deer can survive with a missing limb, and some authors believe amputation is a viable alternative to euthanasia. Myopathy may occur following attempts to evade predators (including humans) and may be evidenced by recumbent, hyperthermic, hyperventilating deer with hindlimb paralysis. Irreversible metabolic changes usually result.4 Johne’s disease, epizootic hemorrhagic disease, coronavirus diarrhea, rotavirus diarrhea, colibacillosis, salmonellosis, and mycobacteriosis represent common infectious disease presentations for deer. Because of chronic wasting disease, many state wildlife and fisheries departments have discontinued deer rehabilitation. Veterinarians can obtain state regulations from their state wildlife agents.

Reptiles Reptiles in the process of shedding, especially snakes, should be handled gently. Overmanipulation of the animal can result in damage to the new underlying skin. Lizards, snakes, and crocodilians can be examined using manual restraint. We recommend taping a crocodilian’s mouth closed to minimize the likelihood of injury to the handler. As a group, chelonians are difficult animals to evaluate clinically. Many chelonians are capable of withdrawing into the margins of their shells when threatened, becoming “bony boxes.” Clinicians experienced in evaluating chelonians have devised methods for coaxing them out of their shells. For medium to small sized chelonians, gently touching the hindlimbs will often lead to the animal extending its head. Slow, deliberate movements can also help reduce fearful responses by the animal and retraction into its shell. Many tortoises extend their forelimbs and head if tilted slightly downward, perhaps in an effort to avoid falling. Above all, an examination of a chelonian requires patience. In the end, many chelonian patients require sedation or anesthesia to perform an exam (see Table 19-4). In those cases where the animal can be examined without anesthesia, it is important to minimize the amount of stress placed on the head and neck when extracting the animal from its shell. Excessive force can lead to cervical injury. A thorough exam should be done on every patient, and the outline found in Box 19-3 can be used for reptiles too. With the exception of obvious traumatic injuries, reptiles show few or subtle signs of disease. A magnification lens will aid in detection of lesions in smaller reptiles. From a distance, lizards should ambulate normally and resist handling. An undisturbed, healthy snake should be coiled or partially coiled, and when approached, it should explore the air with its head and flick its tongue to detect any new odors. The hydration status of a reptile can be assessed by examining skin turgor (e.g., increased skin turgor will result in “wrinkles” on the animal’s body), the oral cavity (e.g., excessive ropy saliva indicates dehydration), and the eyes (in snakes, an opaque, wrinkled spectacle is an indication of dehydration; in

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515 lizards, snakes, and chelonians, sunken eyes indicate dehydration). Hematologic data can be used to confirm that a patient is dehydrated. A standard ophthalmic exam should be done. Retrobulbar abscesses, tumors, or injury can cause unilateral exophthalmos. Unilateral enophthalmos may indicate injury, whereas bilateral enophthalmos can result from microphthalmia, inanition, or dehydration. Edema or vascular obstruction can cause bilateral exophthalmos. The cornea and lens should appear clear. One exception is a snake during ecdysis, when its spectacle may appear a gray-blue color. Corneal ulcers may be obvious or covered with plaques that are protecting healing tissue. The oral cavity and glottis should be examined for discharge, signs of injury, broken teeth, and signs of necrotic stomatitis (caseous plaques, ulcers, inflammation). A rubber or plastic spatula can be used to open the mouth of snakes and lizards to examine the oral cavity. Although these can also be used for chelonians too, a metal paperclip or dental scaler is often more appropriate. The paperclip or dental scaler can be inserted at the most rostral point of the mandible and used to draw the mandible down. In lizards that are acrodonts, care should be taken to avoid injurying the nonreplaceable teeth. The beak of a chelonian should be examined for any flaking, malocclusion, or fractures. Nares should be examined for ulceration or discharge. Tympanic membranes should be examined for the presence of ectoparasites (e.g., lizards) or aural abscesses (e.g., chelonians). The entire body should be palpated to determine the animal’s body condition and identify any abnormal masses or fractures. A snake should appear rounded, with prominent epaxial muscles running the length of the spine. If the animal is emaciated and/or dehydrated, the animal will assume a more triangular shape on cross-section as the epaxial muscle mass atrophies. In lizards, the pelvic bones and spine are more prominent in emaciated animals. Chelonian body condition can be assessed by examining the temporal muscles and the muscles covering the long bones. A chelonian’s carapace should be convex and even, and the scutes should be without lesions. Measurement of the carapace at midline should be taken. General guidelines for respiratory rate in reptiles are 10 to 30 breaths per minute, with variations according to age, temperature, and species. Thoracic auscultation has limited use but can be facilitated with the placement of a damp cloth between the stethoscope and the animal. In this way, breath sounds may be heard; however, cardiac evaluation is still difficult. Thoracic percussion can be used in chelonians to check for lung field abnormalities. The animal should be resting on the palm of one hand or platform, and then the carapace tapped with the index finger of the other hand. The sound should be symmetric. If the tone is dull, this denotes fluid or increased soft tissue mass. A “tinny” sound is generally produced if there are superficial carapacial scute or dermal bone plate lesions. Aquatic species with pulmonary disease can be placed in water and checked for inconsistencies in buoyancy (e.g., one side floating higher than the other). An ultrasonic Doppler can be used to assess the heart rate.

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516 Wounds, abnormal patterns of keratinization, and swelling of the integument should be noted. In terrestrial species, asymmetric patterns of toenail wear can be associated with lameness. All limbs should be examined for swellings, muscle atrophy, pain, and abnormal movement. The contralateral limb can be used to help discern the extent of abnormalities if the condition is not bilateral. Urates and feces should be examined. The urate portion should be chalky and should vary from white to yellow in color. Terrestrial reptiles may pass colorless liquid urine with the stool. Urates and urine may appear green if colored by biliverdin, the major bile pigment in reptiles. The appearance of green urates and urine can be an indication of chronic anorexia or liver disease. The most common reason reptiles present to a veterinary hospital are traumatic injuries (e.g., hit by automobile or landscaping equipment, domestic pet predation), poisonings, and infectious diseases. Animals suffering from a traumatic injury may present with both soft tissue and orthopedic injuries. Many of these injuries are contaminated with opportunistic Gram-negative bacteria and should be managed accordingly.10 Chemical spills or chemical run-off can cause a large number of reptile deaths, especially in aquatic species. Mycoplasmosis, iridovirus infections, and herpesvirus infections have all been associated with mortality events in wild chelonians. Other reasons for presentation include necrotic stomatitis, pneumonia, epidermal disorders, septicemia, cellulitis, parasitism, and organochlorine toxicity (a potential cause of aural abscesses in chelonian species). A thorough diagnostic work-up is required to identify a specific etiology for a reptile case.

REASONS FOR EUTHANASIA The practice of wildlife medicine should follow standard triage protocols. Animals that cannot be rehabilitated and released into the wild should be considered for either euthanasia or permanent captive placement. There are restrictions associated with captive placement, and questions regarding placement should be directed to state and federal wildlife agencies. A final decision on a patient’s outcome should be made only after careful assessment of the patient’s condition, likelihood for rehabilitation and release, and potential for placement. In assessing the significance of a physical disability or abnormal behavior pattern, the possibility must be considered that, although the condition may not be immediately lifethreatening, it would by its nature permanently prevent the animal from surviving in the wild. In such cases the alternative to euthanasia would be permanent placement in captivity. This should only be pursued if, for the foreseeable future, it could be determined that the animal would maintain a reasonable quality of life in captivity and there is a sound justification for this action. Justifications for retaining permanently disabled wildlife cases in captivity include captive breeding programs, education programs, and imprint models.12 Permanently disabled adult animals have long been used to produce offspring for both endangered species and falconry programs. Much of the conservation success associated with the bald eagle and

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peregrine falcon in the United States was based on this type of model. Many of the education programs provided by wildlife hospitals and rehabilitators use permanently injured wildlife. These types of education presentations allow children and adults to observe these animals up close and, in many cases, display the negative impact humans have on these animals. Rehabilitators that work with orphaned animals can benefit from using imprint models. Imprint models are used to teach an orphaned animal what it should do in the wild to survive. When a decision is made to euthanize a wildlife patient, a standard domestic species euthanasia protocol can be followed. We prefer to sedate or anesthetize a wildlife patient before delivering the final euthanasia solution. Birds can be masked down with isoflurane (5%, 1 L oxygen). The euthanasia solution (barbiturate overdose) can then be delivered into the basiocciptal sinus, heart, available blood vessel, intraosseous catheter, or coelomic cavity. Euthanasia dosing may vary among different solutions and manufacturers. Delivery of the drug into the heart, a blood vessel, or an intraosseous catheter will result in an almost immediate cessation of cardiac function. If the euthanasia solution is delivered into the coelomic cavity, the animal may require 5 to 20 minutes to expire. Additional doses of euthanasia solution can be given if needed. Mammals can also be masked down, although for larger specimens, an injectable anesthetic is generally used (e.g., ketamine). Again, the euthanasia solution can be given intravenously, intracardiacally, intraosseously, or intrathoracically/intraabdominally. It is also recommended that reptiles be preanesthetized before being euthanized. Again, dissociative agents are preferred. Freezing reptiles is not considered an appropriate euthanasia method.

ORPHAN CARE Orphaned animals are one of the most common wildlife presentations to the veterinary hospital. The fact that concerned citizens take the time out of their schedules to collect and present these animals is a sign that humans have great compassion for animals. In many of these cases, the citizen presenting the animal becomes emotionally attached and has a desire to monitor its progress. It is important for veterinarians to show compassion to these individuals while also explaining to them how nature works. For some of the presentations, the animals will surely be orphaned. A bird that was blown from its nest after a thunderstorm and cannot be safely replaced or juvenile opossums found on a dead female that was hit by a car represent are orphaned truly. However, a fawn or rabbit kidnapped from its nest is not. We believe that the encounters with these citizens represent a great opportunity to discuss wildlife conservation and educate the public on the importance of protecting wildlife. All orphaned wildlife presentations should be considered emergencies. Juvenile animals have limited hydration and energy stores, and after a short period of time (hours), they may develop negative energy and fluid levels. Correcting for these deficits is essential to patient survival. As with other pre-

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sentations, it is imperative to have proper species identification and a complete history when accepting orphaned animals. Many orphaned wildlife cases may have been cared for by the public before being presented to the veterinary hospital, and in almost all cases the care was inappropriate or insufficient. The natural history of the species will ultimately determine the care it should receive in captivity. As reptiles are born precocial, they do not require any special attention as juveniles and will not be discussed in this section.

Avian If a baby bird is found on the ground, every attempt should be made to locate the nest and replace the bird; however, if replacement of the animal into the nest places a human at risk, it should not be done. In most cases, the orphaned bird will be accepted by the parents, giving it the best chance for survival. It is important to realize that some orphaned animals are intentionally orphaned by the parents because of limited food resources. Although this may appear harsh to the public, it is natural selection. Most baby birds presented to veterinarians will have a subnormal body temperature, and warming the bird to an appropriate temperature to maximize the animal’s metabolic rate should be done first. A heating pad under the “nest” (nest substitutes will be discussed later) is the easiest method of providing heat, but bottles or bags of warmed fluids can be used as well. If access to an incubator is available, the nest should be placed in a temperature between 85° F and 95° F (29° C and 35° C). Ultimately, the temperature selected should be based on the age and plumage of the animal. In general, the highest temperatures are reserved for the hatchlings, moderate temperatures for the nestlings, and lowest for the fledglings. Placing baby birds in direct sunlight is not recommended, as heat prostration is likely. Being able to recognize hyperthermia or hypothermia in an orphaned bird is essential to correcting the problem. In general, hyperthermic birds will pant, extend their necks, and hold their feathers tightly against their bodies. Birds with a subnormal body temperature will be depressed, feel cool to the touch, and elevate their feathers in an attempt to trap heat. The provision of a nest is important to maintain a bird in a specific area for heating purposes and to encourage normal development. Nest substitutes can be made in a variety of ways. Used natural nests should not be provided, as they are a source of various parasites and bacterial and fungal pathogens. The nest must be constructed to assure that the nestling rests in a natural position. In the wild, the nest provides an environment that supports the bird with it legs under its body, feet under the chest, and body weight centered in the back, knees, and lower coelomic cavity. The curved sides of the nest give the body this support and allow the nestling to push against the side of the nest for balance while feeding. Inappropriate nest construction can result in a bird’s developing skeletal anomalies (e.g., splay leg) that limit the potential for release.13 Suitable nest substitutes can be made from small plastic tubs (e.g., margarine containers or flower pots) that are

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517 perforated for air flow, or plastic berry boxes. The nest should be filled with crumpled paper, paper towels, or cloth (e.g., towel, old T-shirt). Occasionally a bird’s toenails may become caught in the cloth or paper substrate, so the animal should be monitored closely. Trimming the nails will prevent this from occurring. While the bird should be secure and supported in its nest, it should never be wrapped so that it cannot move. Orphaned birds can be categorized into one of two groups, altricial or precocial, and each has its distinct needs. Altricial species hatch in a helpless state (e.g., blind and featherless). They are unable to feed or care for themselves in any way. Precocial species are down-covered and ambulatory and can feed themselves soon after hatching. Examples of precocial species include ducks, chickens, geese, and quail. Most other species are altricial and require intensive care in captivity. Feeding regimes will vary greatly with species, but general principles do apply. A bird should be warmed before it is fed. This will maximize the metabolic rate and gastrointestinal tract motility. If the bird is dehydrated, then fluids should be given next. Animals that are dehydrated will have decreased gastrointestinal times, which can lead to obstipation or constipation. Avoid physical contact during the feedings whenever possible. Hatchling birds are prone to imprint. Feeding the birds using a surrogate (e.g., puppet look-a-like) or by hiding behind a blanket or towel will ensure the animal does not imprint onto humans. Placing a mirror in front of the animal so that its own reflection is the only animal it can see may also be done. Feeding intervals for birds should be consistent to promote regular gastrointestinal transit times. If the bird does not gape to be fed, its beak can be gently opened by pressure on the base at each side. The beaks are really delicate, and care must be taken to avoid damaging them. Most baby birds will gape when the nest is slightly disturbed, as this mimics the parent landing on the side of the nest. The bird will usually stop gaping when it is satiated. It is generally difficult to overfeed these animals, although there may be some instances when birds will continue to gape with a full crop. Feeding should be discontinued in these birds until their crop is emptied; otherwise, they may develop crop stasis. Sharp metal objects (e.g., tweezers) should not be used to feed these birds, as these objects can damage delicate mouth and crop tissues. The type of food being offered will vary with species, but many will present having been fed on bread and cow’s milk, which is not a natural diet in any species and can result in secondary complications. After each meal, the bird will pass dropping in a sac, which should be removed immediately. As the nestlings mature, they will be able to void over the side of their nest. Many species of juvenile birds benefit from placement in nests containing conspecifics of the same age. This will encourage natural behaviors and help minimize the chance of imprinting. It is ideal to place orphaned birds with a rehabilitator who may have more of the same species (as long as no injuries have been sustained). For orphaned nestlings to develop normal social behavior, they must be exposed to a conspecific during the critical period (within 36 hours after hatching for precocial species, variable for altricial species).14 In the absence of an

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appropriate adult, a surrogate should be provided. The order of preference for surrogates is juvenile or fledgling conspecifics, sibling conspecifics with a bird-skin puppet, and finally a puppet alone. Hand-rearing orphaned birds is very time consuming and has a high mortality rate. It is important to consider all of these factors when deciding to accept this challenge. When working with orphaned birds, it is important to estimate the age of the animal in order to develop a management plan for the bird. Hatchlings are naked, have closed eyes, and need to be hand-reared. Nestlings are partially feathered, will gape and call when disturbed, and will also need to be hand-reared. Fledglings are fully feathered, but they retain down feathers on the head and back and have less intensive rearing needs. Fledglings can leave the nest, but they still depend on their parents for feeding and protection. They may present to a practice as orphaned when they are displaced from their parents and should be returned near the area found under the cover of vegetation if no predators are present. Emaciated orphans should be fed every 15 minutes for 1 or 2 hours until they have regained their strength. Feedings should then be extended to half-hour intervals and eventually hourly intervals. Diurnal orphans can be fed from 7 am until 7 pm. Nocturnal species are generally fed on a similar basis, but with the room darkened. In general, small bird nestlings will consume 10% to 20% of their body weight in food daily. Body weight should be measured daily to ensure that the nestling is gaining an appropriate amount of weight. The frequency of feeding will vary with the age and species of the bird (Table 19-6). In the wild, diurnal species are fed only during the day and are brooded overnight. Crepuscular and nocturnal species are likewise primarily fed during their peak activity periods. Once a bird fledges, it should be encouraged to feed independently on appropriate foods. The foods can initially be presented in a shallow bowl but then ideally presented in more natural ways that encourage foraging. For nestlings less than 2 weeks old, a substitute nest, as described previously, should be provided. Branches, natural vegetation, and shallow water containers should also be provided. Once the bird has fledged, it should be placed in an outdoor aviary. By 3 or 4 weeks, most small birds will be independent and can be released using soft-release methods. A soft-release program is one in which the birds are housed in an outdoor aviary that allows for natural foraging for about 2 weeks before release.

TABLE 19-6

The choice of food used for hand-rearing orphans will vary with species, but it should be as varied as possible to ensure the proper balance of carbohydrates, fats, proteins, vitamins, and minerals. The food should also be readily available, affordable, and easily prepared. The nestling feeding experience will affect the feeding success later in the bird’s life; thus, it is important to provide as natural a diet as possible. Other important considerations to include when developing an orphaned animal rehabilitation program are the need to expose the animal to conspecifics to develop its normal song, lessons on predator avoidance, and migratory function (if appropriate). For some birds, exposure to their own species’ song must occur within 50 days after hatching. This can be provided with tape recordings if no adults are available. It is imperative that orphaned animals not be exposed to mammalian or avian predators; otherwise they may become habituated to their presence and lose their natural protective response to avoid these animals. For nocturnal migratory species, a full view of the setting sun and night sky must be provided (especially in the second month posthatch) for development of migratory function.14

PIGEONS AND DOVES In the wild, squabs (hatchling pigeons and doves) differ from other altricial chicks, requiring less than six feedings per day.3 The primary reason for this is associated with the type of nutrition (crop milk) these animals receive from their parents. For these birds, crop milk provides the sole source of nutrients during the first few days of life and remains the primary food for the first week to 10 days after hatching. Gradually, crop milk is mixed with the adult diet and fed to hatchlings by both adult parents. Crop milk is crucial for the survival of squabs, as they hatch in a relatively undeveloped state. Squabs are unable to use food found in their environment, cannot digest an adult bird’s diet, hatch with their eyes unopened, and do not possess feathering for thermoregulation after hatching. The composition of crop milk is similar to that found in mammals; it contains water, fat, protein, and ash. The primary difference between the avian milk and mammalian milk is in the carbohydrate and protein fractions. Pigeon milk is devoid of lactose. It does, however, contain high levels of crude protein and fat. When working with these animals, attempts to mimic this formula should be made. The following formula (for a day’s feeding) has been recommended: Mix one hard-boiled egg yolk (mashed), 3 tablespoons mixed baby cereal, 3 tablespoons oatmeal, and 3 tablespoons cornmeal.13 We have used both a

Recommended Feeding Schedule for Orphaned Birds Based on Age

Age

Appearance

Feeding regime

1-4 days 5-7 days 8-14 days 15-21 days 22-28 days 29-42 days

Egg tooth evident, naked, eyelids fused Naked, early development of pin feathers Feathers covering the body, complete feathers on wings and tail Wing and tail feathers fully grown Fledged Capable of flight

q15 min for 12 h/day q30-60 min for 12 h/day q60-90 min for 12 h/day q2h for 12 h/day, encourage self-feeding q2-3h for 12 h/day, encourage self-feeding Should be self-feeding

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commercial critical care psittacine formula and a psittacine neonatal formula (Lafeber Company, Cornell, IL) with good success in these animals too. Pigeons and doves also feed differently than other birds. A squab gets its food by thrusting its beak far down either side of its parents’ esophagus, where it can have access to the crop milk. Squeakers, or nestling pigeons and doves, will feed the same way but instead take seeds from the parents’ crops. To compensate for this feeding technique, we tube the squab or squeaker or cut the tip off of a 1-ml or 3-ml syringe, fill the syringe with gruel, and place the beak of the bird in the openended syringe. This technique can be used to simulate the normal strategy of having the beak inserted into the esophagus of the parent bird. Once the beak is inserted into the syringe, the bird will thrust its head and start imbibing the gruel. Squeakers that are free-feeding can be offered water-soaked gamebird chow.

PRECOCIAL YOUNG There are two different feeding strategies among the precocial birds: free-feeding birds (e.g., gamebirds, waterfowl, and plovers) and those fed by their parents (e.g., grebes, divers, rails, terns, and gulls). A variety of homemade diets may be made that are similar to the diet described for doves and pigeons, but fortunately many of these animals readily accept chicken or turkey starter diets. To further diversify these diets, live insect pupae (e.g., mealworms) can be added. All precocial birds should have a shallow dish of water near their food.

GAMEBIRDS Orphaned gamebirds are not commonly presented to veterinary hospitals because the chicks are precocial, and social imprinting is uncommon. Soon after birth, these birds leave their nest and maintain a close relationship with their nestmates and parents. When the parent birds denote danger, they set a decoy for a predator to protect their juveniles. Even truly orphaned chicks are more capable of caring for themselves than other avian species. If a chick is in immediate danger, handrearing can be attempted; however, certain species, such as quail, are especially difficult to hand-raise because they require a diverse diet comprised of various insects and weed seeds.13 Orphaned gamebirds should be offered a quiet, secure cage. Most of these birds will accept chick crumb or poultry pellet and fresh water in a shallow dish. Artificial heat, as discussed in the general avian section, should be provided. A suspended clean feather duster can be used as a synthetic parent.

WATERFOWL Whereas certain species of orphaned waterfowl are easy to care for, others are not. Mallard ducks and Canada geese will readily adapt to captive care and accept a gamebird chow or chicken starter soaked in water. Supplementing the starter diet with invertebrates (e.g., mealworms) is also highly recommended. On the other hand, orphaned wood ducks do not adapt well to captivity and often experience high mortalities. To limit these losses, every attempt should be made to transfer these cases to a specialized facility for care.

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WADERS Wader chicks are rarely seen in captivity but may present if the parents are killed or they have been displaced from their nest. Once stabilized with supportive care, healthy orphans can be offered an adult appropriate diet (in smaller amounts). Once fully fledged (about 3 weeks of age for smaller species, 6 weeks for larger species), the chicks are independent. Immediately after fledging is the optimal time for release.2

SEABIRDS Other than gulls, seabird orphans are rarely presented to veterinary hospitals, as they usually migrate out to sea soon after fledging. Orphaned gulls, on the other hand, commonly present as orphans after being displaced from nests. They must be reared in groups of conspecifics to avoid imprinting. Chopped fish, day-old chicks, or fish-flavored cat foods can be offered. An avian-specific multivitamin and mineral supplement should also be added to the diet.5

RAPTORS Although it is important to limit human contact with all orphaned avian wildlife, this is considered most important with raptors. Imprinted raptors pose a risk not only to themselves but also to humans. Raptors that have lost their fear of humans have been known to attack humans that enter their territory. Veterinarians and wildlife rehabilitators have a duty to the animals in their care, and to the community in which these animals are released, to minimize the potential for imprinting. Orphaned raptors should be raised with similar aged conspecifics if it is at all possible, as this will minimize the likelihood of imprinting. If no similar aged orphans are available in the immediate vicinity, every effort should be made to contact local rehabilitators to find a companion, or at a minimum, a surrogate adult from the same species. Surrogate raptors can be an invaluable resource to those who are rehabilitating orphaned raptors. A surrogate puppet can also be used as a last resort. When preparing a diet and feeding regime for a particular raptor, it is important to correctly identify the species and have a realistic age estimate. Food choices should mimic the natural diet of the species as closely as possible. Rodents (e.g., mice, rats, gerbils, guinea pigs, rabbits), chickens (chicks), and invertebrates (e.g., crickets and mealworms) are the most commonly available commercially raised prey items for raptors. Some facilities recommend feeding wild squirrels and songbirds that die of noninfectious causes (e.g., traumatic injury), as this is economical and a natural process for many of the opportunistic feeding raptors. Unfortunately, these types of prey items may still serve as a source of parasites or other latent infectious disease for the raptors. Raptors should never be offered a wild prey item that was euthanized using a barbiturate overdose. To elicit a feeding response in a raptor, the food item should be placed in front of the beak or touched to the tactile bristles at the base of the beak. Newly hatched raptors should be fed every 2 to 3 hours for 2 to 3 days. It is important not to overfeed these animals, as constipation can occur. For the

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520 first 2 to 3 days of life, no bone, fur, or feathers should be offered. At 3 days to 1 week of age, mice, pinky rats, or smaller songbirds can be offered every 3 to 5 hours, with no large bones or bone splinters and no fur or feathers. After the first week, the amount of fur and feathers offered, as well as the size of morsel, should be gradually increased until the raptor chick is eating the entire contents of the prey item. Raptor chicks will generally eat more food than an adult of the same species, reaching a peak at 2 to 3 weeks when they will eat more than double an adult.3 Downy raptors should be fed every 4 to 6 hours and fed as much as they will take at one feeding. Do not offer artificial casting materials (e.g., dog fur, cotton), as they have been implicated in causing intestinal blockage and crop impaction.15 Once the chick’s eyes are open and they are investigating with their beaks, fresh carcasses should be left in the environment. The body weight of these animals should be monitored closely and the diet adjusted accordingly. At 2 to 3 weeks of age, various sized perches and branches should be offered. Once the chick is fully feathered and attempting to use its wings, it can be moved to an outdoor aviary to begin learning to fly and build strength.

Mammals GENERAL All orphaned mammals should be provided an enclosure that is dry and warm and has an appropriate nesting material (e.g., shredded paper, towels). Excessive humidity often leads to the development of severe dermatitis. The patient’s body temperature and hydration status should be monitored closely. Orphaned mammals that are dehydrated or hypothermic will reduce their caloric intake. Because these animals do not have a ready source of energy reserves, anorexia or cachexia can be fatal. The fecal output of each orphan should be monitored closely. A reduction in fecal output may be suggestive of a decreased gastrointestinal transit time and should be corrected immediately. Orphaned mammals do not need to be bathed or washed. All feeding utensils should be cleaned thoroughly before being used to limit the transfer of infectious diseases between patients. Orphans should be weighed daily and their diet altered if necessary.

SQUIRRELS With squirrels, it is important to, first, determine whether the animal is actually an orphan. Juvenile squirrels are frequently displaced from their nest after a major weather event (e.g., periods of high winds), but the parent(s) will later retrieve them and return to the nest. If this appears to be the case with a recent presentation, then the animal should be returned to where it was found and placed in a protected area (e.g., shrubs) until the parents return. In general, orphaned squirrels are easy to care for. These animals should be housed in a secure, warmed enclosure. An electric heating pad or warmed water bottle can provide supplemental heat. A nest substitute can be placed in the enclosure to mimic an animal’s natural setting. A small open cardboard

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box with tissue paper, paper towels, or a cotton blanket will serve as an appropriate nest substitute. A squirrel’s eyes open between 19 and 21 days of age. Males and females may need to be separated; sibling penis suckling is common as the penis can be mistaken for a teat. Omnivore (e.g., canine) or carnivore (e.g., feline) milk replacers most closely represent the natural composition of a squirrel’s milk. The formula should be made daily, refrigerated between feedings, and warmed to body temperature before each feeding. A small syringe (e.g., 1-ml or 3ml) can be used to nurse the squirrel. For feeding, the animal can either be placed in lateral recumbency or sternal recumbency. We prefer positioning the animals in sternal recumbency. The formula should be delivered at a pace found acceptable to the squirrel, as feeding the animal too quickly can lead to the aspiration of the milk. Neonates should be fed every 2 hours, with a total volume of 3 ml/day. Anogenital stimulation should be performed after each feeding until the squirrel has urinated and defecated. Dehydrated orphans should be fed a balanced electrolyte solution for one or two feedings and gradually introduced to the milk formula over the following three to four feedings. Diarrhea and bloat commonly occur in animals that have reduced or hypermotile gastrointestinal transit times. When these problems occur, the volume of formula should be decreased.16 Neonates are messy feeders, and excess formula should be cleaned from their face after each feeding with a warm washcloth. After being fed, neonatal squirrels will sleep until their next feeding. Once a squirrel’s eyes have been open for about a week (25-28 days old), begin offering formula from a shallow dish (