The Laboratory Nonhuman Primate

Author: Jeffery D. Fortman | Jeffrey D. Fortman | Terry A. Hewett | B. Taylor Bennett | Lisa Halliday

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Library of Congress Cataloging-in-Publication Data Fortman, Jeffrey D. The laboratory nonhuman primate / Jeffrey D. Fortman, Terry A. Hewett, and B. Taylor Bennett. p. cm. — (Laboratory animal pocket reference series) Includes bibliographical references (p. ). ISBN 0-8493-2562-5 (alk. paper) 1. Primates as laboratory animals—Handbooks, manuals, etc. I. Hewett, Terry A. II. Bennett, B.T. (B. Taylor) III. IV. Series SF407.P7 F67 2001 636.9′8—dc21

2001037497

This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage or retrieval system, without prior permission in writing from the publisher. The consent of CRC Press LLC does not extend to copying for general distribution, for promotion, for creating new works, or for resale. Specific permission must be obtained in writing from CRC Press LLC for such copying. Direct all inquiries to CRC Press LLC, 2000 N.W. Corporate Blvd., Boca Raton, Florida 33431. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation, without intent to infringe.

Visit the CRC Press Web site at www.crcpress.com © 2002 by CRC Press LLC No claim to original U.S. Government works International Standard Book Number 0-8493-2562-5 Library of Congress Card Number 2001037497 Printed in the United States of America 1 2 3 4 5 6 7 8 9 0 Printed on acid-free paper

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dedication

To my wife, Michele, and daughter, Claire, for their unwavering love, support, and patience. — Jeffrey D. Fortman

To laboratory caregivers of nonhuman primates great and small. — Terry A. Hewett

To all the staff of the Biologic Resources Laboratory with whom I have had the pleasure to work over the years, and to my wife and children who have supported me in that work. — B. Taylor Bennett

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acknowledgment The authors wish to acknowledge the generous and significant contributions of Joi Holcomb (illustrations), Jay McElroy (illustrations), and Maria Lang (photography).

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preface The use of laboratory animals, including nonhuman primates, continues to be an important part of biomedical research. With many species of laboratory animals, the person responsible for animal facility management, animal husbandry, and regulatory compliance is also responsible for the performance of technical procedures directly related to the research project. Due to the special requirements for housing and management of nonhuman primates, it would be unusual for one individual to have all of these responsibilities; but even in institutions where these responsibilities are shared, there is a need for a quick reference source for investigators, technicians, and animal caretakers who provide care for nonhuman primates used for research, teaching, and testing. This handbook is intended to be such a reference source and should be particularly valuable for those individuals who may not have extensive training and experience with these unique animal species. The handbook is organized into six chapters: “Important Biological Features” (Chapter 1), “Husbandry” (Chapter 2), “Management” (Chapter 3), “Veterinary Care” (Chapter 4), “Experimental Methodology” (Chapter 5), and ”Resources” (Chapter 6). Because much of the information in the literature on nonhuman primates originates from a small number of institutions that care for large numbers of nonhuman primates on a routine basis, the number of articles in a given area or on a specific subject is often very limited, making it difficult to do a comparative review of the literature. This fact makes it difficult for authors of a text such as this to provide a critical review of the literature in putting together the necessary information to which the reader needs access. Thus, the information contained in this book is a combination of the authors’ knowledge of the literature, the practices of their colleagues at other institutions, and their own combined experience of more than 50 years caring for nonhuman primates. © 2002 CRC Press LLC

The final chapter, “Resources,” provides the user with lists of possible sources and suppliers of additional information, animals, feed, sanitation equipment, cages, and veterinary and research supplies. The lists are not exhaustive and do not imply endorsement of listed suppliers over unlisted suppliers. These lists are meant to be a starting point for the readers to develop their own lists of preferred suppliers. The literature resources in this book are listed in two categories: References when the information contained in the text can be traced to a specific peer-reviewed publication; and Selected Readings when the information is considered to be of general knowledge to those who have experience working with nonhuman primates. Readers who find themselves in the position of providing care for nonhuman primates and without the necessary formal training or experience or ready access to individuals with that experience are encouraged to seek out such individuals and rely heavily upon them for advice and direction.

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the authors Jeffrey D. Fortman, D.V.M., received his doctorate degree in Veterinary Medicine from the University of Illinois at Urbana, Champaign in 1985, and completed a postdoctoral training program in laboratory animal medicine at the University of Illinois at Chicago in 1991. He is a Diplomate of the American College of Laboratory Animal Medicine. He works at the University of Illinois at Chicago as the Associate Director of the Biologic Resources Laboratory and has 13 years of experience in the clinical veterinary care and management of nonhuman primates, and supporting research utilizing Old and New World species. Terry A. Hewett, D.V.M., received her doctorate degree in Veterinary Medicine from Colorado State University in 1986, and completed a residency in laboratory animal medicine at the University of California, Davis in 1991. She is a Diplomate of the American College of Laboratory Animal Medicine. She works at the University of Illinois at Chicago as a clinical veterinarian and has 12 years of experience in the clinical veterinary care of nonhuman primates and supporting research utilizing Old and New World species. B. Taylor Bennett, D.V.M., Ph.D., received his doctorate degree in Veterinary Medicine from Auburn University and his Ph.D. from the University of Illinois Medical School. Dr. Bennett is a Diplomate of the American College of Laboratory Animal Medicine. He is currently the Associate Vice Chancellor for Research Resources and the Director of the Biologic Resources Laboratory of the University of Illinois at Chicago. Dr. Bennett has served as the President of the Association of Primate Veterinarians, the President of the American Association for Laboratory Animal Science, and a member of the Board of Directors of the National Association for Biomedical Research, the American © 2002 CRC Press LLC

College of Laboratory Animal Medicine, and the Association of Laboratory Animal Practitioners. Dr. Bennett’s professional interests are centered upon improving the quality of care provided to laboratory animals. As part of this interest he has been heavily involved in many educational programs and projects at all levels of animal care and use. In this capacity, he has developed a training course for animal technicians seeking AALAS certification from which the AALAS Instructional Guide for Technician Training was developed. He served on the editorial review board for The Biomedical Investigator’s Handbook for Researchers Using Animal Models, which is published by the Foundation for Biomedical Research. He has served as the senior author of the Essentials for Animal Research: A Primer for Research Personnel, which was published by the National Agricultural Library, and he was the senior editor for the two-volume ACLAM text, Nonhuman Primates in Biomedical Research. He has given more than 100 presentations and published more than 50 papers.

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contents 1 IMPORTANT BIOLOGICAL FEATURES Introduction Taxonomy New World Monkeys (NWM): General Characteristics New World Monkeys (NWM): Commonly Used Species in Research Old World Monkeys (OWM): General Characteristics Old World Monkeys (OWM): Commonly Used Species in Research Functional Morphology Limbs and Vertebral Column Muzzle, Nose, and Olfactory Senses Visual and Auditory Senses Digestive System The Skull and Brain Reproduction/Placentation/Growth and Development Behavior Solitary Existence Multi-Male/Multi-Female Groups Single-Male/Multi-Female Groups Family Groups Communication Visual Signals Tactile Signals Body Language Signals Anatomic/Physiological Features Normative Values Clinical Chemistry Parameters Hematology © 2002 CRC Press LLC

Blood Coagulation Values Blood Gases Blood Types Tooth Eruption Times Reproductive Biology Sex Determination Reproductive Cycle Sex Skin Breeding Systems Pregnancy Diagnosis Parturition 2 HUSBANDRY Introduction Housing General Considerations for Primate Housing Facilities Room Design Features Equipment Maintained in Room Primate Enclosures Materials Cage Design Cnsiderations and Features Environmental Conditions Environmental/Psychological Enrichment Special Considerations Nutrition Dietary Requirements Novel Foods and Foraging Treats Potable Water Sanitation Transportation Shipping Crates Certificates of Health and Acclimation Status Recordkeeping Individual Animal Records Group/Colony Records Institutional Recordkeeping Identification Permanent Methods Temporary Methods

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3 MANAGEMENT Introduction Regulatory and Accrediting Agencies, and Compliance The United States Department of Agriculture (USDA) The National Institutes of Health (NIH), Public Health Service (PHS) The United States Food and Drug Administration (FDA) and the Environmental Protection Agency (EPA) The Centers for Disease Control (CDC) The Fish and Wildlife Service (FWS) Association for the Assessment and Accreditation of Laboratory Animal Care International (AAALAC) Institutional Animal Care and Use Committee (IACUC) Occupational Health and Safety Training Safe Work Practices Personal Protective Equipment Physical Injuries B Virus Exposure Allergic Reactions Experimental Hazards Zoonoses B Virus (Cercopithecine herpesvirus 1) Tuberculosis Bacterial Agents of Gastrointestinal Origin Protozoal Agents of Gastrointestinal Origin 4 VETERINARY CARE Preventive Health Program Sources Quarantine Conditioned Colony Health Surveillance Separation of Species Clinical Management Basic Veterinary Supplies Clinical Signs of Illness in Nonhuman Primates Therapeutic Agents Common Clinical Problems Viral Diseases Bacterial Diseases Parasitic Diseases © 2002 CRC Press LLC

Reproductive Conditions Miscellaneous Conditions Anesthesia and Analgesia General Principles Peri-Anesthetic Management Anesthetic Agents Analgesic Agents Principles of Inhalation Anesthesia Endotracheal Intubation Aseptic Surgery Facilities/Features/Equipment Personnel Pre-Operative Preparation Operating Room Procedures Post-Operative Care Euthanasia 5 EXPERIMENTAL METHODOLOGY Introduction Restraint Physical Restraint Methods Chemical Restraint Operant Conditioning and Training Methods Sampling Techniques Blood Collection Urine Collection Bone Marrow Aspiration and Biopsy Cerebrospinal Fluid Collection Semen Collection Amniotic Fluid Collection Compound Administration Parenteral Administration Methods Oral Administration Methods Miscellaneous Procedures Disarming Canine Teeth Bimanual Rectal Palpation Necropsy 6 RESOURCES Organizations Publications © 2002 CRC Press LLC

Books Periodicals Electronic Resources Primate Sources Possible Commercial Sources of Nonhuman Primates Contact Information for Nonhuman Primate Sources Nonhuman Primate Transportation Resources Nonhuman Primate Transportation Services Laboratory Services Feed Equipment Sanitation Cages, and Research and Veterinary Supplies Possible Sources of Cages, and Research and Veterinary Supplies Contact Information for Cages, and Research and Veterinary Supplies Primate Research Centers NCRR-Supported Regional Primate Research Centers Other Primate Research Centers Equivalents and Conversions REFERENCES SELECTED READINGS

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1 important biological features introduction Nonhuman primates may well be the first recorded animal subjects for scientific research.1 Since that initial use, they have been utilized in many areas of biomedical and behavioral research where their similarity to humans makes them uniquely valuable animal models. Of particular note is the role that nonhuman primates have played in virological research and in the development and testing of important vaccines for diseases such as polio and hepatitis. They have also been used historically in the areas of reproductive physiology, behavior and learning, and neurophysiology. Of the commonly used laboratory animals, primates are unique in that they are not a domesticated species, and even those that have been specifically bred for use in research laboratories are not far removed from their ancestors who were captured in the wild. For this reason, it is important that those who care for and work with these animals understand the natural environment from which these species arose, how that environment has affected their evolutionary development, and how that development affects their behavior when they are housed in a laboratory environment.

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taxonomy Nonhuman primates belong to the order Primates which contains three suborders: (1) Prosimii, which are often considered to be preprimates and include a variety of Asian and African species that are small, generally nocturnal animals who rely more on their sense of smell than their vision; (2) a newly recognized suborder, Tarsioidea, which includes the Tarsius spp. that may represent the bridge between the pre-primates and the true primates; and (3) Anthropoidea, which are the true primates and include two infraorders: the Platyrrhine or New World monkeys (NWM) and the Catarrhine or Old World monkeys (OWM).

New World Monkeys (NWM): General Characteristics New World monkeys (NWM) are found in Central and South America and consist of two families of primates: the Callitrichidae, which include the marmosets and tamarins, and the Cebidae, which include howler, woolly, spider, woolly spider, owl (night), and squirrel monkeys as well as titis, sakis, capuchins, and uakaris. The marmosets and tamarins are small, fruit-eating animals that are active in the daytime and live in small groups in an arboreal environment. They are unique among the primates in that except for the big toe, all of their digits have long, sharp claws. Marmosets and tamarins are very territorial and make high-pitched, bird-like calls. The capuchin-like monkeys (Cebidae) are a much more diverse family whose members vary in size from the large (6 to 8 kg) fruit-eating howler monkey, to the smaller nocturnal owl monkey (1 kg), to the even smaller squirrel and titis monkeys (<1 kg). Their diets vary and, with the exception of the owl monkey, they are active in the daytime and live in groups that vary from 2 (the monogamous titis monkeys) to 30 (howler monkeys). • As platyrrhine monkeys, the NWM have broad flat noses with round nostrils separated by a broad fleshy septum. • They exhibit a great variety of types, body forms, and sizes and little sexual dimorphism. • They all have tails and some are prehensile. They are completely arboreal with long limbs and no ischial callosities. They have long curved nails (marmosets and tamarins have claws) and most lack digital mobility. © 2002 CRC Press LLC

• They lack cheek pouches, have the prosimian dentition, and require a source of vitamin D3. • They exhibit no swelling of the sexual skin.

New World Monkeys (NWM): Commonly Used Species in Research There are six species of NWM that one might normally encounter in a laboratory animal facility. Three of these species are members of the family Callitrichidae, which includes the marmosets and tamarins. • The common or tufted marmoset, Callithrix jacchus jacchus, weighs between 350 and 400 g, adapts well to captivity, and is the most widely used of the Callitrichidae (Figure 1.1). • The cotton-top tamarin, Sanguinus oedipus oedipus, weighs between 425 and 550 g and is frequently used in colon cancer research (Figure 1.2). • The moustached tamarin, Sanguinus mystax, weighs between 600 and 650 g and is frequently used in hepatitis A research. There are three species of the family Cebidae that are found in research laboratories.

Fig. 1.1 Callithrix jacchus jacchus, common marmoset. (Photo used with permission from the Biologic Resources Laboratory’s slide collection.) © 2002 CRC Press LLC

Fig. 1.2 Sanguinus oedipus oedipus, cotton-top tamarin. (Photo used with permission from the Biologic Resources Laboratory’s slide collection.)

Fig. 1.3 Saimiri sciureus, squirrel monkey. (Photo used with permission from the Biologic Resources Laboratory’s slide collection.) © 2002 CRC Press LLC

• The owl monkey, also known as the night monkey (Aotus trivirgatus), is probably the most unusual of the New World primates because it is the only nocturnal true primate. These animals weigh from 900 to 1200 g and, due to genetic variations within the genus, must be karyotyped prior to breeding. • The most commonly used NWM is the squirrel monkey, Saimiri spp. These animals weigh from 500 to 1000 g and should also be karyotyped before breeding (Figure 1.3). • The largest NWM commonly found in laboratories is the capuchin monkey, Cebus spp. They weigh from 3 to 5 kg and have exceptional manual dexterity and a semi-prehensile tail (Figure 1.4).

Old World Monkeys (OWM): General Characteristics The Old World monkeys (OWM) of Africa and Asia belong to the family Cercopithecidae. Within this family are the guenons, macaques, baboons, and mangabeys, which belong to the subfamily Cercopithecinae. Nonhuman primates in this subfamily tend to be large (10 to 20 kg), terrestrial primates with omnivorous dietary habits that forage for their food in open country. Some species of Asian macaques have a more arboreal lifestyle as do the African mangabeys. A second

Fig. 1.4 Cebus apella, blacked-capped capuchin monkey. © 2002 CRC Press LLC

subfamily, the Colobinae, includes the colobine monkeys of Africa and the leaf-eating monkeys of Asia. These animals all have a specialized digestive mechanism for processing a folivorous diet. • The Catarrhine or Old World monkeys (OWM) have narrow noses with comma-shaped nostrils separated by a narrow nasal septum. • They lack prehensile tails, have ischial callosities, and many have sex skin that undergoes variable degrees of swelling. • They are diurnal, have opposable thumbs, do not require dietary vitamin D3, and typically have cheek pouches. There is marked sexual dimorphism, especially among terrestrial savannah dwellers.

Old World Monkeys (OWM): Commonly Used Species in Research The OWM most commonly used in biomedical research belong to the family Cercopithecinae — the cheek-pouched monkeys — and include two major groups: the Asian macaques, and the baboons, guenons, and mangabeys of Africa. • Macaques have the greatest geographic distribution of the nonhuman primates. They are large, robust animals with large cheek pouches, prominent ischial callosities, variable sexual swelling, and marked sexual dimorphism. The most commonly used macaques in biomedical research are Macaca fascicularis, the crab-eating macaque or cynomolgus monkey, with females weighing from 2 to 6 kg and the males weighing 4 to 8 kg; and Macaca mulatta, the rhesus monkey, which is larger, with females weighing from 4 to 9 kg and the males weighing 6 to 11 kg (Figures 1.5 and 1.6). • Baboons, Papio spp., are very sexually dimorphic terrestrial primates. The males have a profuse mantle of fur and large canines. Both sexes have prominent ischial callosities, and the females show marked sexual swelling. The most commonly used species of baboon used in biomedical research are of the species Papio anubis (the olive baboon; Figure 1.7) and Papio cynocephalus (the yellow baboon and hybrids of these species, Papio cynocephalusanubis). Baboons are relatively large primates, with the females weighing from 11 to 15 kg and males weighing 22 to 30 kg. Due to their large size, baboons are only housed in a few laboratory animal facilities in the United States. © 2002 CRC Press LLC

Fig. 1.5 Macaca fascicularis, cynomolgus (crab-eating) macaque.

Fig. 1.6 Macaca mulatta, rhesus macaque. (Photo used with permission from the Biologic Resources Laboratory’s slide collection.) © 2002 CRC Press LLC

Fig. 1.7 Papio anubis, olive baboon. (Courtesy of T. Butler, Southwest Foundation for Biomedical Research.) • The guenons are arboreal primates, weigh from 2 to 6 kg, and are distinguished by their striking facial hair and the brightly colored genitalia of the males of the species. They are found throughout a wide range of Africa, have reduced ischial callosities, and the females do not have sex skin. The most commonly encountered guenon in the laboratory setting is the African green monkey, Cercopithecus aethiops. • Mangabeys are long-legged, slender primates with large ischial callosities, little sexual dimorphism, and the females show marked reddening and swelling of the sexual skin. They often have white eyelids and prominent cheek tufts. The sooty mangabeys, Cercocebus torquatus atys, are subject to natural infections with leprosy and are an asymptomatic carrier of Simian immunodeficiency virus (SIV). The largest Old World primate used in biomedical research is actually an ape and belongs to the family Pongidae, the great apes. The chimpanzees, Pan troglodytes, are large powerful animals, with the females weighing approximately 40 kg and the males 50 kg. Chimpanzees are maintained in only a few research facilities in the United States. © 2002 CRC Press LLC

functional morphology Functional morphology, or the relationship of form and function, is an important issue to understand when working with nonhuman primates. The differences between the functional morphology of nonhuman primates and the other laboratory animals are based on evolutionary relationships and differences in locomotor activity and habitat usage — the arboreal and terrestrial surroundings that are part of their natural environment.

Limbs and Verterbral Column Nonhuman primates have clavicles and their radius and ulna are separate, which allows them to pronate and supinate their forelimbs. In species with a true prehensile tail, the tail has pressure pads with flexion creases and ridges on the ventral surface. Ischial callosities are seen in OWM and are more prominent in the large species that squat or rest upright in trees or raised rock formations.

Muzzle, Nose, and Olfactory Senses Development of olfactory sensitivity is inversely proportional to the development of mobility and sensitivity of the hands and the overlapping of the visual fields. In general, primates have a poor sense of smell compared to other mammals, although scent marking is important in the prosimian primates and several species of NWM. A welldeveloped muzzle is indicative of an enlarged masticatory apparatus, as seen in the baboons, or the enlarged larynx of the howler monkeys.

Visual and Auditory Senses The development of the visual senses is a requirement for a nonhuman primate’s arboreal lifestyle. To be able to judge spatial relationships, they have overlapping visual fields, which results in the frontal position of the orbits and a reduced muzzle size. They also have a cone-type retina with a fovea centralis. Nocturnal prosimian species have mobile external ears.

Digestive System The dentition consists of incisors, canines, and two types of cheek teeth comprised of either 32 or 36 teeth with functional specialization of the incisors and canines, and modification of the cusp pattern of © 2002 CRC Press LLC

the cheek teeth. NWM have a dental formula of (2I,1C, 3PM, 3M/2I, 1C, 3PM, 3M) × 2 with the third molar missing in marmosets and tamarins [(2I,1C, 3PM, 2M/2I, 1C, 3PM, 2M) × 2]. OWM have a dental formula of (2I,1C, 2PM, 3M/2I,1C, 2PM, 3M) × 2. The canine teeth of nonhuman primates show tremendous variation in size, shape, and projection, which is influenced by both sexual dimorphism and/or dietary needs. The longest and sharpest canines are seen in ground-living Cercopithecidae and are the most massive in the Pongidae. Nonhuman primates, although largely vegetarian, are generally considered omnivorous. There is some specialization of the diet.

The Skull and Brain The skull is enlarged, and the foramen magnum is directed inferiorly to accommodate truncal uprightness. In the brain there is progressive development of the cerebral cortex and the cerebellum, which is linked to their visual and tactile acuity. There is an increase in the size of the fiber tracts that connect the cortical and sub-cortical areas and the spinal column.

Reproduction/Placentation/Growth and Development There is a tendency toward year-round breeding, although in the wild, environment will influence seasonality. Nonhuman primates have a unicornuate uterus and usually have single births, although marmosets and tamarins usually have multiple births. Placentation in the true primates is hemochorial, which permits free exchange of serum proteins and favors maternal iso-immunization against mutant proteins. As a rule, nonhuman primates have a prolonged gestation. The postnatal life is prolonged to accommodate the complex learning that must take place. Puberty is delayed, ensuring proper development of the required social skills, and adolescence is accompanied by a marked growth spurt.

behavior Primate behavior is adaptive in nature and this ability to adapt to the environment is a key element in the evolutionary process that ensures survival of the species. Their social behavior is the result of the overall group dynamic that is necessary to optimize survival. Unlike most other mammals, the behavior of primates is learned, and © 2002 CRC Press LLC

not instinctual; thus, primates are more adaptable to different environments. Because their behavior is learned, the management of newborn and juvenile primates must include arrangements for group or social housing that exposes the younger animals to the normal behavior of conspecifics. The ecological niche in which a species develops provides the basis for its behavior patterns. These patterns evolve as part of the process of adaptation and are a major factor in determining behavior patterns once an animal is removed from that niche. The environment in which an animal is maintained will strongly affect its current behavior, regardless of its rearing background and experience. The more experience that an animal has had prior to entering its current environment, the better it should be able to adapt. Nonhuman primates have a complex social structure compared to most animals maintained in the laboratory environment. There are four primary types of social units: solitary existence, multimale/multi-female groups, single-male/multi-female groups, and family groups.

Solitary Existence This type of social unit is, by its very nature, not conducive to procreation of a species and thus is not common in nonhuman primates. Orangutans are an example of a species that uses this type of social structure, as the males tend to associate with the females only for the purpose of breeding. Females, however, usually live with one or more prepubescent offspring.

Multi-Male/Multi-Female Groups This type of social unit is made up of several males and females with various numbers of offspring. There is a dominance hierarchy among males, and the dominant male tolerates the other males. The females form the permanent nucleus of the group. In this matriarchal society, social ranking is determined by the social rank of the animal’s mother, and the order of rank is inverse to the birth order, with the younger females having a higher rank than their older sisters. In males, this matriarchal social ranking lasts until puberty when they leave the group and establish their own identity in another group. This type of social structure is seen in several species commonly found in laboratory animal facilities (squirrel monkeys, capuchins, macaques, and baboons). © 2002 CRC Press LLC

Single-Male/Multi-Female Groups This type of social unit is made up of one male and several females and their offspring. The males will not tolerate the existence of other males; thus, there is a lot of tension and fighting. Of the animals found in laboratory animal facilities, the African green monkey may exhibit this social structure, and macaques can be successfully housed using this social grouping.

Family Groups Family groups are made up of a mated pair and their young. In monogamous families, young are ejected from the group as they reach adulthood. In extended families, the adults are not ejected, but there is often suppression of estrus in the younger females. This type of social group is common among the marmosets and tamarins.

communication Nonhuman primates primarily use visualization and vocalization to communicate, although scent marking is common in prosimians and some NWM. It is important to understand the scent-marking habits of the species that are housed in the laboratory in order to design and implement appropriate sanitation programs that minimize potential adverse effects on normative behavior.

Visual Signals Visual signals are an important component of primate behavior. Everything from the coat color of an animal to complex facial expressions can play an important role in determining an animal’s behavioral response. In macaques, an open mouth gesture (Figure 1.8) is a threat, whereas lip-smacking is a submissive gesture. An openmouth with the top teeth covered is a gesture of play, while the seeming casual yawn (Figure 1.9) that exposes the canine teeth is a warning, “look at my teeth.” An open-mouth grin (Figure 1.10) is a sign of anxiety or fear, whereas a stare (Figure 1.11) is a threatening gesture. Moreover, animal care staff should avoid direct eye contact (a stare) with nonhuman primates because this can be perceived by the animals to be a threatening gesture.

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Fig. 1.8 Open-mouth gesture indicates threat/aggression.

Fig. 1.9 Yawn display indicates threat/aggression.

Fig. 1.10 Open-mouth grin indicates fear or anxiety.

Fig. 1.11 Stare indicates threat/aggression.

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Tactile Signals Grooming is probably the most important tactile behavioral display in primates and the use of group or pair housing in the laboratory allows animals to participate in this important behavior.

Body Language Signals While the end result of what can best be called “body language signals” is the visual image that it portrays, it is important to understand that body language plays an important survival role in the nonhuman primate’s natural environment. Nonhuman primates will rarely show signs of weakness or pain and this fact must be recognized when assessing the behavior and well-being of an animal. It may therefore be important to observe the animals without them knowing that they are being watched in order to accurately assess their well-being. The classic posture of submission, in which an animal presents its rear end or back to an animal (Figure 1.12), thus minimizing any perceived threat, must be recognized by the staff when handling nonhuman primates. The larger-than-life pose in which nonhuman primates, especially males, bristle out their hair and appear much bigger than they really are, is an important threatening display and should be recognized.

Fig. 1.12 Submissive posture. © 2002 CRC Press LLC

anatomic/physiological features As mentioned previously, the Primate Order is distinguished by its lack of specialization and this is especially true when it comes to the anatomical features. While nonhuman primates have many of the anatomical features of quadrupedal animals, human anatomy books are often much more useful resources than the standard veterinary anatomy books. The same statement holds true for basic physiology, and applied physiology that is so important when dealing with disease conditions. While the lack of specialization is a key feature of nonhuman primates and human reference books are a valuable resource when questions on anatomy and physiology arise, there are several anatomic/physiologic features that should be mentioned. • Sexual dimorphism is quite noticeable in some species of nonhuman primates, and this is especially noticeable in terms of the actual body size and the impressive canines that can often be found in males of some species. The most striking examples of sexual dimorphism can be seen in those species whose lifestyle is largely terrestrial in nature (e.g., the baboons). • The tails of two subfamilies of NWM, Atelinae and Alouattinae, are truly prehensile with a tactile pad on the ventral surface while the Cebinae have a functionally prehensile tail without a tactile pad. • The subcutaneous fat of nonhuman primates is yellow, as is the fat in the abdominal cavity. • The uterus is unicornuate and the ovaries are closely approximated to the uterine body by relatively short fallopian tubes. • Ovulation and luteinization are spontaneous in nonhuman primates, and the OWM have a menstrual cycle that in some species of Cercopithecinae is notable for the external signs of their cyclicity. The changes that occur during the menstrual cycle are most noticeable in terms of the swelling that occurs in the perineal area, although females in some species also show signs of color change and swelling on the hindlimbs and on the face and head.

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normative values Nonhuman primates exhibit a wide range of individual variation, both between species and within species in their basic biological parameters. Normative data (Tables 1.1 and 1.2) have been summarized from the information available in the literature and from institutions that house large numbers of the species of interest. Diet, age, social ranking, method of housing, and duration of housing in a laboratory setting will influence these parameters. It is necessary for each individual laboratory to establish normative values for their facility. Interpretation of physiological parameters should be made in consultation with veterinary staff knowledgeable about the animal and methodology used. Normative values can be a useful adjunct to other means of clinically assessing individual animals when serial data is not available; however, care should be exercised when interpreting values given the many sources of variability. A few examples follow that are not intended to be an all-inclusive list, but rather illustrate some of the frequently encountered sources of variability. Bodyweights of normal adult male squirrel monkeys may vary with season, while that of adult females of all species will vary with gravidity. Caloric intake may be affected by an animal’s access to food. Small New World monkeys normally pick through food and eat choice morsels first, more food wastage is common with these species, and personnel feeding these animals must account for these behaviors when determining food needs. Low socially ranking animals will often exercise more and spend less time eating than higher ranking cohabitants in group-housed confinement systems. Heart rate and respiratory rate vary tremendously, depending on methods and circumstances under which they are measured. Anesthestics commonly used to chemically restrain animals for safe handling may have cardiovascular effects as well as cause a loss of body heat. Marmosets normally have a circadian variation in their body temperature con4 cordant with their diurnal activity. Hence, daytime body temperatures are significantly elevated from those taken in the evening when animals are resting. Normative values determined from population means and standard deviations may not reflect a given population if the individual data points have a nonnormal distribution, such as might occur in facilities housing animals of differing ages.

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TABLE 1.1: NORMATIVE PHYSIOLOGICAL WORLD SPECIES Parameter Weight: adult male Weight: adult female Respiratory rate Heart rate Rectal temperature Daily food intake

AND

Units Saimiri sciureus2 grams 892–1094 grams 700 /min 55–58 beats/min 215–263 °F 101.7–103.6 °C 38.7–39.8 g 45–60

BIOLOGICAL DATA

OF

NEW

Callithrix jacchus jacchus3 318–493 309–387 NFa 4 194–242 (telemetry) 98.2–101.5 (telemetry)4 36.8–38.6 (telemetry)4 20/adult + fruit

Note: Superscript numbers refer to references at end of book. a

NF, not found.

TABLE 1.2: NORMATIVE PHYSIOLOGICAL WORLD SPECIES Parameter Weight: adult male Weight: adult female Respiratory rate Heart rate (telemetry) Rectal temperature Daily water intake Daily food intake Urine excreted/day Blood volume

Units M. kg kg /min beats/min °F °C ml g ml ml/kg

AND

BIOLOGICAL DATA

mulatta5 M. fascicularis 6–11 4–85 5 4–9 2–6 35–50 30–546 98–1227 115–2436 5 98.6–103.1 98.6–103.1 5 37–39 37–39 6 400–600 350–950 400–600 350–5506 NFa 150–5506 5 50–96 55–75

OF

OLD

Papio spp. 22–305 5 11–15 22–356 85–905 5 98.6–103.1 5 37–39 6 400–600 1000–15006 150–4006 5 50–70

Note: Superscript numbers refer to references at end of book. a

NF, not found.

Clinical Chemistry Parameters Tables 1.3 and 1.4 provide normative serum biochemical values for New and Old World species, respectively. As with normative and hematological values, much variability exists within species and between species, and individual laboratories should determine normative serum biochemical values for their facility utilizing management and handling techniques compatible with the experience of the staff, conditioning of the animals, and objectives of the researchers. The values provided in Tables 1.3 and 1.4 can be used in the health assessment of individual animals in consultation with veterinary staff familiar with the animals and techniques used to obtain the samples as well as to perform the laboratory tests. A few common sources of © 2002 CRC Press LLC

variability include sample handling errors such as low blood glucose resulting from serum not being separated promptly from the clot and hemolyzed serum samples producing elevations in serum potassium concentration.16 Chemistry values can also vary due to the age of an animal,17–21 as young healthy animals will exhibit an elevation of serum alkaline phosphatase. With the exception of propofol, most commonly used anesthetics do not affect serum biochemistry values.22 Propofol is formulated in a lipid vehicle that can artificially elevate serum triglyceride levels. Lipemia caused from obtaining samples from nonfasted animals will increase triglyceride, cholesterol, and serum calcium values above those samples obtained from fasted animals.

TABLE 1.3: NORMATIVE SERUM CHEMISTRY PARAMETERS (RANGE = MEAN ± 1 SD) OF NEW WORLD MONKEYS Parameter Units Total protein g/dl Albumin g/dl Alkaline phosphatase µ/l LDH µ/l AST µ/l ALT µ/l GGT µ/l CPK µ/l BUN mg/dl Creatinine mg/dl Glucose mg/dl Sodium mEq/l Potassium mEq/l Chloride mEq/l Calcium mg/dl Total bilirubin mg/dl Direct bilirubin mg/dl Amylase µ/l Lipase µ/l Triglycerides mg/dl Cholesterol mg/dl Phosphorus mg/dl

Saimiri sciureus2 Callithrix jacchus jacchus8 5.9–7.9 6.4–8.0 3.6–4.8 4.4–5.8 183–533 34–88 11–235 108–328 90–280 106–196 74–294 38–72 0–207 5.8–15.39 0–1942 920–24109 29–49 15–29 0.7–1.1 0.4–0.69 72–133 124–220 143–157 153–169 4.7–6.7 3.5–4.7 103–11810 93–121 8.7–10.5 9.1–11.7 0.2–1.4 0–1.02 0–0.4 0.08–0.229 327–704 337–1523 NFa 1–65 42–108 63–209 87–216 136–234 3.2–6.610 4.2–7.0

Note: Superscript numbers refer to references at end of book. 9Reference range converted to ±1 SD. a NF, Not found.

© 2002 CRC Press LLC

TABLE 1.4: NORMATIVE SERUM CHEMISTRY PARAMETERS (RANGE = MEAN ± 1 SD) OF OLD WORLD SPECIES ANESTHETIZED KETAMINE HYDROCHLORIDE Parameter Total protein Globulin Albumin Alkaline phosphatase LDH AST ALT GGT CPK BUN Creatinine Glucose Sodium Potassium Chloride Calcium Magnesium Total bilirubin Direct bilirubin Amylase Lipase Triglycerides Cholesterol Iron Phosphorus HDL Plasma osmolality

M. fascicularis11

Units

M. mulatta7

g/dl g/dl g/dl µ/l

6.4–7.0 2.2–2.6 4.1–4.5 504–821

7.2–8.0 3.1–4.013 3.8–4.2 89–166

6.7–7.6 3.0–4.2 2.8–4.2 97–400

µ/l µ/l µ/l µ/l µ/l mg/dl mg/dl mg/dl mEq/l mEq/l mEq/l mg/dl mEq/l mg/dl mg/dl µ/l µ/l mg/dl mg/dl µg/dl mg/dl mg/dl mOsm/kg

450–788 19–38 31–50 66–97 425–1173 15–21 0.45–0.71 46–60 142–149 3.3–3.7 104–110 8.6–9.2 1.38–1.60 0.15–0.23 NFa 213–331 a NF 24–42 122–154 a NF 5.7–7.2 15 63–69 282–295

232–43513 20–57 23–6114 69–16314 121–893 15–21 0.7–1.114 48–69 142–150 3.3–3.9 14 109–119 8.5–9.3 NFa 0–0.12 NFa 287–43114 a NF 44–76 106–148 NFa 6.7–9.714 19–103 NFa

203–350 29–49 34–56 28–50 217–566 12–17 0.75–1.25 58–108 147–152 3.3–4.4 95–103 8.4–9.6 NFa 0.1–0.3 0.05–0.15 166–321 1.5–8.5 51–88 71–128 42–95 2.1–3.8 33–79 NFa

Note: Superscript numbers refer to references at end of book. a

WITH

15

Papio spp.12

Range = Mean ± S.E.

NF, not found.

Hematology Tables 1.5 and 1.6 provide normative hematological parameters for New and Old World species, respectively. As with normative and serum biochemical values, significant variability exists within species and between species, and individual laboratories should determine normative hematological parameters for their facility utilizing management and handling techniques compatible with the experience of the staff, conditioning of the animals, and objectives of the researchers.17–20,23–25 New World species can often be safely hand © 2002 CRC Press LLC

restrained and Table 1.5 lists values from unsedated animals. Old World species are frequently sedated with ketamine hydrochloride for chemical restraint; hence, Table 1.6 reflects data from anesthetized animals. Elevation of white blood cell count with a corresponding lymphopenia is often seen in animals that are frightened or stressed TABLE 1.5: NORMATIVE HEMATOLOGICAL PARAMETERS (RANGE = MEAN ± 1 SD) OF UNSEDATED NEW WORLD SPECIES Parameter Hematocrit RBC WBC Hemoglobin Neutrophils Lymphocytes Eosinophils Basophils Monocytes Platelets MCV MCH MCHC

Units % × 106/ml × 106/ml g/d1 % % % % % × 103 fl pg g/dl

Saimiri sciureus2 38.6–43.6 6.3–7.1 6.0–9.1 12.2–13.6 36–6610 27–5510 0–1210 <110 0–610 NFa 58–63 19–20 31–33

Callithrix jacchus jacchus9 32–54 4.6–6.6 4.9–11.3 12.6–19.6 27–59 35–67 0–1.4 0–2.1 0.4–6.2 180–382 66–78 14–32 34–45

Note: Superscript numbers refer to references at end of book. a NF, not found.

TABLE 1.6: NORMATIVE HEMATOLOGICAL VALUES (RANGE = MEAN ± 1 SD) OF OLD WORLD SPECIES Parameter Hematocrit RBC WBC Hemoglobin Neutrophils Lymphocytes Eosinophils Basophils Monocytes Platelets MCV MCH MCHC

Units M. mulatta7 % 37–40 × 106/ml 5.1–5.6 6 × 10 /ml 4.2–8.1 g/dl 12.0–13.1 % 26–52 % 39–72 % 0–4 % 0–0.4 % 1–4 × 103 260–361 fl 71–75 pg 22.8–24.5 g/dl 31.0–33.4

M. fascicularis11 33.1–37.5 5.3–6.3 6.1–12.5 11.0–12.4 35–61 34–56 1.3–9.1 0–0.214 0.4–3.0 300–51214 59–66 19–21 32–35

Note: Superscript numbers refer to references at end of book.

© 2002 CRC Press LLC

Papio spp.12 36–41 4.6–5.3 6.7–12.5 11.7–13.5 48–76 22–50 0–2 0 0.5–3.5 233–399 74–80 24–26 32–34

at the time of sampling. Aged animals or those raised in indoor captive situations may show lower white blood cell counts than animals originating from outdoor or wild caught populations. As stated in previous sections, care must be employed when interpreting hematological parameters. Veterinary staff can help interpret incidental from pathologic changes with the aid of physical examination observations and, if necessary, serial blood sampling from selected individuals.

Blood Coagulation Values Table 1.7 provides reference values for coagulation in Old World species. These values are particularly subject to laboratory variation, and hence it is recommended that reference ranges be determined by individual laboratories that may have need to perform these tests.

Blood Gases Arterial blood gases and blood pH for rhesus monkeys, baboons, and squirrel monkeys are provided in Table 1.8. These parameters are usually obtained as part of an anesthetic monitoring protocol to ensure adequate ventilation for animals undergoing prolonged or complex surgical procedures. In general, trends in these values are used to assess an animal’s response to anesthetic manipulation by personnel trained in anesthesia. Care should be used when interpreting values from a single sample. Inadvertent venous sampling and partial arterial occlusion (such as occurs during aortic crossclamping procedures) are examples in which aberrant values might be obtained, and personnel interpreting results should be aware of the methods used to obtain the sample. Always include an overall patient assessment when evaluating blood gas results and consult with veterinary personnel familiar with their usage.

TABLE 1.7: COAGULATION REFERENCE RANGES (MEAN ± 1 SD) WORLD SPECIES Parameter Prothrombin time Activated partial prothrombin time Fibrinogen

Units seconds seconds mg/dl

M. mulatta13 M. fascicularis14 10.3–11.7 9.9–11.1 NFa NFa

18.2–22.6 NFa

Note: Superscript numbers refer to references at end of book. a NF, not found.

© 2002 CRC Press LLC

IN

OLD

Papio spp.12 12.3–13.7 29.5–34.9 142–190

TABLE 1.8: ARTERIAL BLOOD GAS VALUES (MEAN ± 1 SD) IN RHESUS MONKEYS (MACACA MULATTA) AND BABOONS (PAPIO SPP.) ANESTHETIZED WITH KETAMINE HYDROCHLORIDE AND SQUIRREL MONKEYS (SAIMIRI SCIUREUS) ANESTHETIZED WITH PHENCYCLIDINE HYDROCHLORIDE Parameter pH pCO2 pO2

Units mmHg mmHg

M. mulatta7 7.36–7.42 34–42 81–96

Papio spp.12 7.33–7.39 32–40 61.5–86.5

Saimiri sciureus10 7.32–7.40 29–33 72–81

Note: Superscript numbers refer to references at end of book.

Blood Types Macaques and baboons are often used in research that requires transfusion of whole blood. Blood groups of these species are discussed in the literature.26–28 Table 1.9 summarizes the human A, B, O blood groups of Old World species. Personnel familiar with blood typing human blood can perform A, B, O typing on baboon and macaque serum or saliva using an agglutination inhibition technique. Direct hemagglutination is not used because these animals do not have ABO group antigens on their red blood cells.26 In addition to typing by the human A, B, O antigens, it is possible to identify simian-type blood groups; however, reagents must be prepared against primate red cells and consequently this is not readily done by facilities housing small numbers of these animals. Transfusion without regard to blood type can be done for life-threatening, single-time treatments in Old World species. Large volumes of antigen may prevent transfusion reactions from developing in sensitized incompatible recipients too. For research that might involve the administration of multiple transfusions, the compatibility of the donor and recipient will determine the red blood cell viability in the recipient. Antibody-related accelerated elimination of incompatible blood cells may render a transfusion 27 worthless if pretransfusion compatibility testing is not performed. Transfusion for the purpose of supplying platelets may still be justified between partially incompatible donor/recipient pairs. TABLE 1.9: ABO BLOOD GROUPS Species M. mulatta M. fascicularis Papio spp.

A Rare Present Present

Source: From Reference 26.

© 2002 CRC Press LLC

OF

AB Rare Present Present

OLD WORLD SPECIES B Common Present Present

O Likely present Present Rare

Tooth Eruption Times Tooth eruption times for the permanent teeth of New and Old World species are provided in Tables 1.10 and 1.11, respectively. Tooth eruption times are helpful in estimating the age of subadult nonhuman primates when their date of birth is not available. The number of premolar and molar teeth differs between squirrel monkeys, marmosets and Old World species. The first molar is the first permanent tooth to erupt in all species shown in Tables 1.10 and 1.11, followed by the incisor teeth. The canine teeth in male squirrel monkeys, macaques, and baboons take longer to erupt than in females and it is one of the last teeth to erupt. The rear molar erupts last in Old World species. TABLE 1.10: MEAN ESTIMATED TIME IN MONTHS PERMANENT TEETH IN NEW WORLD SPECIES

I1 2 I C P2 3 P P4 M1 2 M M3

Saimiri sciureus Maxillary Mandibular 10 9 10 11 19a–22b 19a–22b 13 13 12 13 12 12 5 4 8 7 19 15

UNTIL

ERUPTION

OF

Callithrix jacchus jacchus Maxillary Mandibular 7 7 9 9 11 11 10 10 9 9 8 8 4 4 8 7

a

Female tooth eruption time. Male tooth eruption time. Source: From Reference 29. b

TABLE 1.11: MEAN ESTIMATED TIME IN MONTHS PERMANENT TEETH IN OLD WORLD SPECIES

I1 2 I C P3 4 P M1 M2 M3

M. mulatta Maxillary Mandibular 30 30 33 31 46 42 42 43 44 44 18 17 40 38 72 68

a

UNTIL

M. fascicularis Maxillary Mandibular 30 30 32 31 41 40 42 43 42 43 20 17 41 40 69 66

Female tooth eruption time. Male tooth eruption time. Note: Superscript numbers refer to references at end of book. Source: From Reference 29. b

© 2002 CRC Press LLC

ERUPTION

OF

P. anubis Maxillary Mandibular 36 30 41 40 46a–73b 43a–67b 62 67 66 67 21 21 50 54 85 85

reproductive biology The reproductive biology of nonhuman primates, in particular Old World species, is very similar to that of humans. As a result, nonhuman primates have been used as a model to study the mechanisms and processes associated with fertility, infertility, pregnancy, and parturition. Moreover, many species of nonhuman primates have been bred extensively in captivity to support production programs for research and species preservation; thus, a wealth of information is available on the reproductive biology of nonhuman primates. Although it is not within the scope nor the intent of this section to cover in detail all aspects of nonhuman primate reproduction, basic information on the reproductive biology of the more commonly used species in research is presented in Tables 1.12 and 1.13. In addition, pertinent information on sex determination, the reproductive cycle, sex skin, breeding systems, pregnancy diagnosis, and parturition is also presented.

Sex Determination In addition to the sexual dimorphism exhibited by some species, adult male and female nonhuman primates can be differentiated by TABLE 1.12: NORMATIVE REPRODUCTIVE PHYSIOLOGICAL AND BIOLOGICAL DATA FOR THE SQUIRREL MONKEY (SM), COMMON MARMOSET, (CM) AND COTTON-TOP TAMARIN (CT) Parameter Sexual maturity Female Male Estrous cycle length Gestation period Birth weight Average number of young per parturition Post-partum estrus Newborn solid food consumption Weaning age Interbirth interval

Units

SM29

CM29

CT29 1.5–2 1.5–2 15a 166a,33 44a,35 235

Years Years Days Days Grams

2.5–3.5 3.5–531 8–10 148–16032 95–11034 135

1.5–2 1.5–2 28a 144a,33 27a,35 235

Days postpartum Weeks

None

7–11

4–6

4

Months Years

6 35 1

2–6 35 0.5

a

33

Mean. Note: Superscript numbers refer to references at end of book.

© 2002 CRC Press LLC

33

33

7–11 433 2–6 35 0.6

TABLE 1.13: NORMATIVE REPRODUCTIVE PHYSIOLOGICAL AND BIOLOGICAL DATA FOR THE RHESUS MONKEY (Mm),a CYNOMOLGUS MONKEY (Mf),b AND OLIVE BABOON (Pa)c Parameter Sexual maturity Female Maled Seasonal breeders Menstrual cycle length Menstruation Gestational period Birthweighte Newborn solid food consumption Weaning age Interbirth interval a b c

Mm30

Mf

Days Days Days Grams Months

2.6–3.5 3–4 Yes 26–30 e 4.6 165–178 47535 1.5–3.0

3.0–3.4 3–4 No 28–32 1–536 155–165 34535 1.0

3–4 3.5–4.5 No 30–37 1–637 173–193 86538 3–6

Months Years

7–14 1.035

12–18f 1.135

6–15 18–24

Units Years

30

Pa30

Mm = Macaca mulatta (rhesus monkey). Mf = Macaca fascicularis (cynomolgus monkey). Pa = Papio anubis (olive baboon).

d

Although males produce viable sperm at this age, they are not considered socially mature, nor do they contribute significantly as a breeder until 2 to 3 years later. Mean. f Some B virus free production colonies wean between 3 to 4 and 7 to 8 months.39 Note: Superscript numbers refer to references at end of book. e

observing the external genitalia. Males have a prominent pendulous penis and scrotal sac, which in most species is located posterior to the penis. Females have a vulva and a shorter anogenital distance than is seen with males. There are some noteworthy species- and age-associated variations that can make it difficult for a novice observer to correctly determine the sex of certain species of nonhuman primates. • The size of the labial folds and clitoris can vary significantly among species, especially New World species. The most extreme example is the spider monkey, in which the clitoris of the adult female is more prominent and pendulous than the penis of the 40 adult male. Adult female New World species with elongated genitalia can be differentiated from males by examining the perineum for the presence of a scrotum. • Newborn and young callitrichids (marmosets and tamarins), and squirrel and cebus monkeys, can be difficult to sex because the external genitalia are of similar size, and in the case of female callitrichids and squirrel monkeys, a false scrotum or pudendal pad © 2002 CRC Press LLC

is present. To correctly identify the sex of these young animals, the external genitalia should be manipulated to demonstrate a round preputial opening (male) or a slit-like vaginal opening (female).33, 34, 41, 42 • In all nonhuman primates, the testes descend into the scrotum before or shortly after birth. Following this initial descent of the testes, there is considerable variation between species as to whether they will remain in the scrotum prior to puberty. In many species, the testes will ascend into the inguinal canal or abdomen only to redescend into the scrotum near puberty. This can further complicate the sex determination of young New World species. The age at which final testicular descent occurs in some of the more common species used in research is (a) callitrichids 8 to 11 months; (b) rhesus monkeys approximately 3 years; (c) cynomolgus macaques and baboons approximately 4 years.40

Reproductive Cycle The reproductive cycle of nonhuman primates and the seasonality with which it occurs vary among species. New World species do not menstruate. Their reproductive cycle is therefore referred to as an estrous cycle, which refers to the period of time from the beginning of one estrus period to the beginning of the next. Estrus is defined as the period of time the female is sexually receptive and nearing ovulation. In general, female callitrichids do not exhibit any obvious external changes that can be used to predict ovulation. This includes changes in outward physical appearance, behavior and vaginal cytology.30, 43 Callitrichids are not considered seasonal breeders because they will cycle, breed, and produce young throughout the year. Because they have a postpartum estrus and a relatively short gestation period, callitrichids will generally give birth twice a year.33, 43 Female squirrel monkeys do not typically exhibit the traditional behavioral changes associated with estrus; however, changes in vagi30 nal cytology can be used to determine ovulation. Squirrel monkeys are seasonal breeders. They have a distinct 3-month breeding season 31 followed approximately 6 months later by a birthing season. The majority of births in the Northern Hemisphere occur between May and August.44, 45 Both male and female squirrel monkeys demonstrate weight increases in association with the onset of their annual breeding season. The torso of males becomes bulkier and there is an increase in testicular size and spermatogenesis. These changes are © 2002 CRC Press LLC

referred to as the “fatted male” condition and are associated with an increase in testosterone.46, 47 The Old World species and great apes exhibit true menstrual cycles because the lining of their uterine wall undergoes periodic sloughing. The length of the menstrual cycle is determined by the number of days between the onset of menses from one cycle to the next. The presence of menses is determined by visual examination of the external genitalia or vaginal swabs for the presence of blood. Menstrual cycles are generally of 4 to 5 weeks duration, with ovulation and an associated increase in the number of matings tending to occur midcycle.48 Baboons and cynomolgus macaques breed and produce young throughout the year, whereas rhesus monkeys maintained under natural lighting conditions are seasonally polyestrous with a hiatus in ovulatory cycles during the late spring and summer months. The peak birthing season for rhesus monkeys maintained under natural lighting conditions occurs during the late spring and early summer 30, 36, 48 months.

Sex Skin Many Old World species of nonhuman primates have sex skin. Sex skin is usually associated with an animal’s perineal region and changes in the skin occur in response to hormonal fluctuations. The location, coloration, and response to the cyclic hormonal changes associated with the menstrual cycle can vary significantly among species. Some noteworthy species variations are described below. • African green monkeys lack sex skin.30 • The sex skin of the female cynomolgus macaque is located at the base of the tail. This tissue will undergo mild swelling midcycle (near ovulation). There are no skin color changes associated with the menstrual cycle in this species.36 • The sex skin of the female rhesus monkey undergoes a series of changes at puberty. When an animal reaches puberty, the skin over the perineum and the back of the legs will become edematous. When the swelling subsides, the affected skin will take on a red coloration. The coloration of this skin in the adult animal can change in conjunction with the hormonal fluctuation of the animal’s menstrual cycle, being palest at the time of menstruation. In some animals, the sex skin will become slightly swollen as the animal nears ovulation. The sex skin usually involves the © 2002 CRC Press LLC

Fig. 1.13 Hind limb in a rhesus macaque with extensive furrowing of sex skin. animal’s perineum and the caudal aspect of the hind legs; however, it can also be found along the caudal aspect of the animal’s abdomen and over the animal’s brow. Sex skin along the caudal lateral aspect of an animal’s legs may have a furrowed appearance (Figure 1.13).36 • Changes in the sex skin of the baboon, chimpanzee, and sooty mangabey can be quite dramatic. In these species, the perineum (sex skin) will undergo marked cyclic changes in size that correlate with the various stages of the menstrual cycle. The perineum in animals near menstruation is flat (Figure 1.14); whereas, in animals near ovulation, the skin will become very swollen (Figure 1.15). Moreover, near ovulation (maximal swelling), the skin of the perineum will appear smooth, shiny, and have a deep red coloration. The cyclic changes in the sex skin of these species can 30,35,36 be used to estimate the time of ovulation. © 2002 CRC Press LLC

Fig. 1.14 Perineal sex skin of a baboon near menstruation.

Fig. 1.15 Perineal sex skin of the same baboon in Fig. 1.14 near ovulation.

© 2002 CRC Press LLC

Breeding Systems A variety of breeding systems are used to produce nonhuman primates for biomedical research. The system employed is dependent in part on the animal species, available facilities, space constraints, labor constraints, production needs, and the needs of the research project. • The mating system typically used for breeding captive callitrichids is monogamous pairing.33 • The mating systems typically used for breeding squirrel monkeys are a single-male, multiple-female or a multiple-male, multiplefemale social configurations.31,44,45 • Old World species such as cynomolgus macaques, rhesus monkeys, and baboons breed well in free ranging environments (corrals and islands) and harem groups maintained in large cages or 35,39,49-52 pens. To support specific research projects, some Old World species are time-mated, which involves maintaining males and females separately, except for the periovulatory period during which the female is housed with the male for a short period of time. • A common method for determining the optimal time for mating rhesus and cynomolgus monkeys is to determine the average menstrual cycle length of the previous three cycles, divide the average cycle length by two, and subtract three. The female is then placed with the male for 2 to 3 days, beginning with this calculated breeding date. The first day of menses should be consid35,46 ered the first day of the menstrual cycle. • In the baboon, the optimal time for mating is determined by calculating from the previous three menstrual cycles when the average day deturgescence (a decrease in sex skin swelling) first occurs and subtract three. The female is then placed with the male for 3 to 4 days beginning with this calculated breeding date. The first day of menses should be considered the first day of the menstrual cycle.35,37

Pregnancy Diagnosis In the laboratory environment, the most practical methods of determining pregnancy are palpation and ultrasonography. Palpation techniques have been used to determine pregnancy in both New and Old World species. Using an abdominal palpation technique, © 2002 CRC Press LLC

experienced personnel can determine pregnancy in callitrichids and squirrel monkeys by palpating a 4- to 5-mm structure (uterus) in the caudal abdomen as early as week 4 or 5 of gestation. Moreover, in these species, manual palpation techniques are often used to estimate gestational age and day of parturition.32–34 In Old World species, the abdominal palpation technique can be used to determine pregnancy by mid to late first trimester; however, a bi-manual rectal palpation technique is recommended for the early detection of pregnancy. Using such a technique, experienced personnel can determine pregnancy in macaques and baboons as early as day 16 of gestation.35 For a description of the bimanual rectal palpation technique, see Chapter 5. In recent years, diagnostic ultrasound has become an important tool in the management of nonhuman primate breeding colonies. Ultrasonography can be used to determine pregnancy in rhesus and cynomolgus monkeys and baboons as early as days 14 and 18, respectively; however, its utility extends well beyond the mere diagnosis of pregnancy. Diagnostic ultrasound can also be used to determine fetal viability, estimate fetal age, assess fetal development, identify the location of the placenta, diagnose reproductive disease conditions, and support a variety of research projects. Tables 1.14, 1.15, and 1.16 provide predictive values for the embryonic and fetal development of the more commonly used Old World species. These values can be used to determine normal growth and estimate gestational age of a developing fetus.

Parturition Parturition, the act of giving birth, is very similar among nonhuman primates. Behavioral changes reported to precede parturition in Old World species include restlessness, changes in eating and sleeping habits, frequent urination, and genital manipulation. Typically, nonhuman primates give birth to a single newborn (twinning is common in the callitrichids) at night or during the early morning while in a squat or upright position. Uncomplicated labor usually lasts 1 to 4 hours, and the vast majority of babies are delivered head first. Licking or cleaning the newborn is a common process for many species, as is the consumption of the placenta. The mothers of Old World species will assist the newborn in finding the nipple; whereas newborn callitrichids and squirrel monkeys receive little direct assistance from their mothers.33–35

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TABLE 1.14: PREDICTED VALUES FOR GESTATIONAL SAC (GS) AND GREATEST LENGTH (GL) FOR THE RHESUS, CYNOMOLGUS MACAQUES (Mmf),a,35 AND OLIVE BABOON (Pa)b,53 c

GD 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 a

GS(Mmf) (cm) 0.14 0.27 0.39 0.51 0.63 0.75 0.87 1.00 1.12 1.23 1.35 1.47 1.59 1.71 1.83 1.94 2.06 2.17 2.29 2.40 2.51 2.62 2.74 2.87 2.96 3.07 3.17 3.28 3.39 3.49 3.60 3.70 3.80 3.90 4.01 d 4.11 — — — — — — — — — —

GS(Pa) (cm) — — — 0.1 0.3 0.4 0.6 0.7 0.9 1.0 1.2 1.3 1.4 1.6 1.7 1.8 2.0 2.1 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9 3.0 3.1 3.2 3.2 3.3 3.4 3.4 3.5 3.6 3.6 3.7d — — — — — — — — —

GL(Mmf) (cm) — — — — — — 0.29 0.33 0.38 0.43 0.49 0.55 0.62 0.69 0.77 0.85 0.94 1.03 1.13 1.23 1.34 1.45 1.57 2.87 2.96 3.07 3.17 3.28 3.39 3.49 3.60 3.70 3.80 3.90 4.01 4.11d — — — — — — — — — —

GL(Pa) (cm) — — — — — — — — — — — — 0.3 0.4 0.4 0.5 0.6 0.7 0.8 0.9 1.1 1.2 1.3 1.4 1.5 1.6 1.8 1.9 2.0 2.2 2.3 2.5 2.6 2.8 2.9 3.1 3.3 3.4 3.6 3.8 4.0 4.1 4.3 4.5 4.7 4.9

Mmf = Macaca mulatta (rhesus macaque) and Macaca fascicularis (cynomolgus macaque). b Pa = Papio anubis (olive baboon). c GD = gestational day. d No data reported beyond this day of gestation. © 2002 CRC Press LLC

TABLE 1.15: PREDICTED VALUES FOR BIPARIETAL DIAMETER FOR THE RHESUS MACAQUE (Mm),a,35 CYNOMOLGUS MACAQUE (Mf),b,35 AND OLIVE BABOON (Pa)c,53 GDd 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88

Mm (cm) 1.3 1.4 1.4 1.5 1.5 1.6 1.6 1.7 1.7 1.8 1.8 1.9 1.9 2.0 2.0 2.1 2.1 2.2 2.2 2.3 2.3 2.4 2.4 2.4 2.5 2.5 2.6 2.6 2.7 2.7 2.8 2.8 2.8 2.9 2.9 3.0 3.0 3.1

Mf Pa (cm) (cm) 1.3 1.2 1.4 1.3 1.4 1.3 1.5 1.4 1.5 1.5 1.6 1.5 1.6 1.6 1.7 1.6 1.7 1.7 1.8 1.7 1.8 1.8 1.8 1.9 1.9 1.9 1.9 2.0 2.0 2.0 2.0 2.1 2.1 2.1 2.1 2.2 2.2 2.2 2.2 2.3 2.3 2.3 2.3 2.4 2.4 2.4 2.4 2.5 2.4 2.6 2.5 2.6 2.5 2.7 2.6 2.7 2.6 2.8 2.7 2.8 2.7 2.9 2.7 2.9 2.8 3.0 2.8 3.0 2.9 3.1 2.9 3.1 2.9 3.2 3.0 3.2

a

GDd 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126

Mm (cm) 3.1 3.1 3.2 3.2 3.3 3.3 3.3 3.4 3.4 3.4 3.5 3.5 3.6 3.6 3.6 3.7 3.7 3.7 3.8 3.8 3.8 3.9 3.9 3.9 4.0 4.0 4.0 4.0 4.1 4.1 4.1 4.2 4.2 4.2 4.2 4.3 4.3 4.3

Mf Pa (cm) (cm) 3.0 3.2 3.1 3.3 3.1 3.3 3.1 3.4 3.2 3.4 3.2 3.5 3.2 3.5 3.3 3.6 3.3 3.6 3.3 3.7 3.4 3.7 3.4 3.8 3.4 3.8 3.5 3.8 3.5 3.9 3.5 3.9 3.6 4.0 3.6 4.0 3.6 4.1 3.7 4.1 3.7 4.1 3.7 4.2 3.8 4.2 3.8 4.3 3.8 4.3 3.8 4.4 3.9 4.4 3.9 4.4 3.9 4.5 3.9 4.5 4.0 4.6 4.0 4.6 4.0 4.6 4.0 4.7 4.1 4.7 4.1 4.7 4.1 4.8 4.1 4.8

GDd 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165

Mm (cm) 4.3 4.4 4.4 4.4 4.4 4.4 4.5 4.5 4.5 4.5 4.5 4.6 4.6 4.6 4.6 4.6 4.6 4.6 4.7 4.7 4.7 4.7 4.7 4.7 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 4.8 f 4.8

Mm = Macaca mulatta (rhesus macaque). Mf = Macaca fascicularis (cynomolgus macaque). c Pa = Papio anubis (olive baboon). d GD = gestational day. e No data reported between gestational day 135 and parturition (day 184). f Average day of parturition. Note: Superscript numbers refer to references at end of book. b

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Mf Pa (cm) (cm) 4.1 4.9 4.2 4.9 4.2 4.9 4.2 5.0 4.2 5.0 4.2 5.0 4.2 5.1 4.3 5.1 4.3 5.1e 4.3 — 4.3 — 4.3 — 4.3 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.4 — 4.5 — 4.5 — 4.5 — 4.5 — f — 4.5 — — — — — — — — — — — — — — — — — — — —

TABLE 1.16: PREDICTED VALUES FOR FEMUR LENGTH FOR THE RHESUS MACAQUE (Mm),a,35 CYNOMOLGUS MACAQUE (Mf),b,35 AND OLIVE BABOON (Pa)c,53 GDd 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88

Mm (cm) 0.3 0.3 0.4 0.4 0.5 0.5 0.5 0.6 0.6 0.7 0.7 0.7 0.8 0.8 0.9 0.9 1.0 1.0 1.0 1.1 1.1 1.2 1.2 1.3 1.3 1.3 1.4 1.4 1.5 1.5 1.6 1.6 1.6 1.7 1.7 1.8 1.8 1.9

Mf (cm) 0.3 0.4 0.4 0.5 0.5 0.5 0.6 0.6 0.7 0.7 0.8 0.8 0.8 0.9 0.9 1.0 1.0 1.0 1.1 1.1 1.2 1.2 1.3 1.3 1.3 1.4 1.4 1.5 1.5 1.5 1.6 1.6 1.7 1.7 1.7 1.8 1.8 1.8

Pa (cm) 0.2 0.3 0.3 0.4 0.4 0.5 0.5 0.5 0.6 0.6 0.7 0.7 0.8 0.8 0.9 0.9 1.0 1.0 1.1 1.1 1.2 1.2 1.2 1.3 1.3 1.4 1.4 1.5 1.5 1.6 1.6 1.7 1.7 1.8 1.8 1.9 1.9 2.0

a

GDd 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113 114 115 116 117 118 119 120 121 122 123 124 125 126

Mm (cm) 1.9 1.9 2.0 2.0 2.1 2.1 2.2 2.2 2.2 2.3 2.3 2.4 2.4 2.5 2.5 2.5 2.6 2.6 2.7 2.7 2.7 2.8 2.8 2.9 2.9 2.9 3.0 3.0 3.1 3.1 3.1 3.2 3.2 3.2 3.3 3.3 3.3 3.4

Mf (cm) 1.9 1.9 2.0 2.0 2.0 2.1 2.1 2.1 2.2 2.2 2.3 2.3 2.3 2.4 2.4 2.4 2.5 2.5 2.5 2.6 2.6 2.6 2.7 2.7 2.7 2.8 2.8 2.8 2.9 2.9 2.9 3.0 3.0 3.0 3.1 3.1 3.1 3.1

Pa (cm) 2.0 2.1 2.1 2.2 2.2 2.3 2.3 2.4 2.4 2.4 2.5 2.5 2.6 2.6 2.7 2.7 2.8 2.8 2.9 2.9 3.0 3.0 3.1 3.1 3.2 3.2 3.3 3.3 3.4 3.4 3.5 3.5 3.6 3.6 3.7 3.7 3.8 3.8

GDd 127 128 129 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145 146 147 148 149 150 151 152 153 154 155 156 157 158 159 160 161 162 163 164 165

Mm (cm) 3.4 3.4 3.5 3.5 3.5 3.6 3.6 3.6 3.7 3.7 3.7 3.8 3.8 3.8 3.9 3.9 3.9 3.9 4.0 4.0 4.0 4.0 4.1 4.1 4.1 4.1 4.2 4.2 4.2 4.2 4.2 4.3 4.3 4.3 4.3 4.3 4.3 4.4 4.4f

Mm = Macaca mulatta (rhesus macaque). Mf = Macaca fascicularis (cynomolgus macaque). Pa = Papio anubis (olive baboon). d GD = gestational day. e No data reported between gestational day 135 and parturition (day 184). f Average day of parturition. Note: Superscript numbers refer to references at end of book. b c

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Mf (cm) 3.2 3.2 3.2 3.3 3.3 3.3 3.3 3.4 3.4 3.4 3.5 3.5 3.5 3.5 3.6 3.6 3.6 3.6 3.7 3.7 3.7 3.7 3.8 3.8 3.8 3.8 3.8 3.9 3.9f — — — — — — — — — —

Pa (cm) 3.9 3.9 4.0 4.0 4.1 4.2 4.2 4.3 4.3e — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

2 husbandry introduction The care and management of nonhuman primates requires knowledgeable staff, well-designed and maintained facilities, and caging and equipment appropriate for the animals in order to maintain their health, both physically and psychologically, and to minimize the risks to personnel. All components of husbandry, including housing, environmental conditions, enrichment methodology, nutrition, sanitation, transportation, recordkeeping, and identification, should be designed and implemented in consideration of the species of nonhuman primates that a facility maintains. Adherence to general concepts of good husbandry as outlined in the Guide for the Care and Use of Laboratory Animals54 (Guide) and a commitment to providing measures to promote psychological well-being utilizing The Psychological Well-Being of Non-human Primates55 will provide direction while still allowing customized solutions to achieve the objectives of good husbandry.

housing Nonhuman primates may be housed in a variety of enclosures ranging from islands and outdoor corrals, where environmental conditions are not closely regulated, to completely indoor housing in individually ventilated rooms or enclosures. Outdoor housing facilities should provide animals with shelter for moving out of wind, rain, or sun (Figure 2.1). Indoor housing facilities rely heavily on heating, cooling, and ventilation systems to provide animals with environments adequate

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Fig. 2.1 Outdoor field cage with shelter. (Courtesy of S. Pearson, Covance Research Products, Alice, TX.) for their needs. The ideal design of indoor facilities should include provision for redundant or backup ventilation to allow for ventilation equipment service and repair without interruption in ventilation to primate housing areas.

General Considerations for Primate Housing Facilities • Functional areas within a facility should be considered, depending on the operational needs of the facility. Commonly utilized functional areas include: food storage, including refrigeration, diet kitchen; nursery; cage/equipment storage; treatment room; post-operative recovery room; radiology/special procedure rooms such as ultrasonography/endoscopy/laparoscopy/fluoroscopy; surgery with associated preparatory and support areas; isolation for immunosuppressed animals or animals with infectious disease; janitorial supply area; necropsy, preferably with downdraft ventilated table(s) and carcass storage; record storage; locker and shower facilities for animal carestaff; infectious waste handling/storage; clinical pathology laboratory; and administrative support areas. • Proximity to cage washer if primate cages and equipment will be sanitized out of the room. • Separation of quarantined animals from conditioned animals. All husbandry of conditioned animals should occur prior to that of quarantined animals unless separate staff can be used for the two different areas. © 2002 CRC Press LLC

• Utilization of animals may determine that animals be located close to procedural areas, if special equipment or caging is necessary. • Prior to any equipment purchases, clearance in all areas where the equipment may be located and travel through should be carefully determined including ceiling height, door height and width, cage wash dimensions, available clearance to access and exit cage wash, and elevator dimensions. Consideration must be made for additional space necessary for transport carts and lift equipment (Figures 2.2 and 2.3), as most items are heavy and awkward to manipulate in confined spaces.

Fig. 2.2 Transport cart loaded with Harford-style macaque cages.

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Fig. 2.3 Lift used to move macaque cages off wall-mounted racks.

Room Design Features • In general, animal rooms are designed following the recommendations found in the Guide.54 They can be multipurpose rooms that hold individual cages or cages suspended from rolling racks or brackets mounted onto walls. Waste falls to the floor or into troughs or trays beneath the cages. All rooms should have floors that slope toward drains that have a minimum diameter of 4 to 6 in., and the drains should be located where they can be easily accessed for maintenance. A one-way viewing window to allow room checks without entering the room can be beneficial should an animal escape from its cage, and when showing visitors through primate housing areas. • Floors should be smooth to facilitate cleaning, but not slippery when wet. The surface should be impervious to moisture and © 2002 CRC Press LLC

resistant to damage from impact. Walls and doors in rooms, corridors, and elevators throughout the facility, where primate caging might travel, should have bumper guards of appropriate height to protect walls from being damaged by moving cages and equipment. Light fixtures in animal rooms should be set into the ceiling or walls and have moisture- and tamper-resistant light diffusers. Rooms housing animals should not have suspended ceilings or nonlocking ceiling access ports that escaped nonhuman primates might open and enter. Any electrical outlets should be water resistant and grounded. • Room temperature must be monitored and wall-mounted maximum-minimum thermometers are an inexpensive method frequently used to track daily temperature fluctuations inside housing areas.

Equipment Maintained in Room • Floor-cleaning implements generally include a hose on a wallmounted bracket and a squeegee. • Experimental equipment should be kept to a minimum in rooms housing more than a few nonhuman primates. Often, remote monitoring stations can be set up in adjacent rooms through the use of tether systems or telemetric monitoring devices to separate electrical equipment from the moisture and animal waste found in animal rooms. • Animals should not have access to any equipment kept in the room, particularly if it is electrical.

Primate Enclosures Caging standards include minimum space requirements based on the species and individual bodyweight (Table 2.1). Enclosures should provide for the animal’s comfort, promote normal growth and development, and ensure health by the use of materials and design features that facilitate thorough sanitization (Figure 2.4). Housing systems must also accommodate study requirements and allow for 56, 57 interaction between animals when possible.

Materials Cages should be constructed of nontoxic construction materials. Stainless steel (Type 304) is commonly used for smaller cages. It is © 2002 CRC Press LLC

TABLE 2.1: NONHUMAN PRIMATE PRIMARY ENCLOSURE SPACE REQUIREMENTS 54, 66 Groupa

a

Weight

Floor Area/Animal

Height

lb

kg

ft2

m2

in.

cm

1

<2

<1

1.6

0.15

20

50.8

2

2.2–6.6

1–3

3.0

0.28

30

76.2

3

6.6–22.0

3–10

4.3

0.40

30

76.2

4

22.0–33.0

10–15

6.0

0.56

32

81.28

5

33.0–55.0

15–25

8.0

0.74

35

91.44

6

>55.0

>25

25.1

2.33

84

213.36

Group 1—marmosets, tamarins, and infants (<6 mo. old) of various species. Group 2—capuchins, squirrel monkeys, and similiar-sized species, and juveniles (6 mo.–3 yr. old) of various species. Group 3—macaques and African species. Group 4—male macaques and large African species. Group 5—baboons and nonbrachiating species > 33.0 lb. (over 15 kg). Group 6—great apes >55.0 lb. (25 kg) and brachiating species.

Fig. 2.4 disposal.

Macaque cages with shared trough system for waste

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very durable and resists oxidation and corrosion from exposure to the animal excreta and the acids and disinfectants used for sanitization. Extruded aluminum is sometimes used, particularly for larger cages, to decrease the weight, yet provides a unit that is strong enough to contain larger animals safely. Aluminum may oxidize when alkaline cleaning agents are used, but this process can be reversed by an acid wash cycle. Recently, some of the newer plastics have been employed and may serve well for housing smaller species although they may not be as durable as metal caging. Numerous references detailed elsewhere provide information on housing designs that have been successfully used to maintain a variety of species of nonhuman primates used in research.58

Cage design considerations and features Lifestyle of species New and Old World species have different lifestyles and behaviors that influence some aspects of cage design. For example, most New World species are primarily arboreal with long tails. These animals require perches and devices to climb, and these are best provided in a cage that has additional height as compared to those designed for more terrestrial species such as the macaques (Figure 2.5). Perches for Old World species should allow the animals to sit on their ischial callosities, yet allow waste to fall to the floor of the cage or preferably beneath the cage out of reach of the animals. Marmosets, tamarins, and owl monkeys require nesting boxes. Handling and restraint In general, individual cages for species such as macaques and baboons are fitted with a false back called a “squeeze-back” that allows animals to be restrained within the cage without having to be physically handled (see Chapter 5). The cages used for macaques are usually made from stainless steel rods for top, front, and floor. The sides and back are made of solid stainless steel. The false back of stainless steel rods fits in front of the true back and can be drawn forward by an operator at the front of the cage by means of handles that attach to the false back (Figure 2.6). Latches secure the false back in the desired position. Cages holding groups of animals often utilize a series of chutes in graduated sizes that animals can be conditioned to run into. They can be separated individually via guillotine doors or sliding horizontal dividers, or the chutes may terminate at a squeeze-

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Fig. 2.5 New World monkey gang cage with perches and climbing devices. (Courtesy of A. Brady, University of South Alabama.) back cage. For small nonhuman primates such as marmosets and tamarins, the nesting box can be used as a transfer unit when fitted with a sliding door (Figure 2.7). Often, chemical restraint methods are used on nonhuman primates (see Chapter 4). For additional information on handling techniques and restraint devices, see Chapter 5. Floors Primary enclosures for nonhuman primates usually have slatted or mesh floors with sufficient openings to allow feces to fall through the enclosure to facilitate sanitation, animal health, and safety of personnel. However, facilities that house groups of animals in indooroutdoor units or outdoor corrals or corncribs have solid floors. These can be successfully managed with appropriate waste removal procedures and vermin control programs.

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Fig. 2.6 Schematic drawing of a squeeze-back macaque cage with sidebar latches.

Fig. 2.7 New World monkey nest box. (Courtesy of K. Boehm, University of Wisconsin.)

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Provision of food and water J-feeders are commonly used for Old World species because the feed will drop into the lower portion of the feeder and animals can be readily fed without opening the cages (Figure 2.8). Also, the animals cannot reach outside the cage through these feeders. Biscuits may not always drop all the way down into J-feeders, and animals with poor dexterity or those who are debilitated may not be able to reach the food unless care is taken when filling the feeders. New World species require elevated feeding stations because they normally feed high up in trees in the wild. Water can be provided by automatic watering systems or bottles. Young, aged, or debilitated animals may not be able to reach automatic watering devices. New animals must learn to use automatic watering devices and carestaff need to check the systems daily to ensure that they are functioning properly.

Fig. 2.8 J-feeder.

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Environmental Conditions Temperature, relative humidity, illumination, ventilation, and noise are factors that can impact animal health and require regular monitoring. Contingency plans should be in place should problems with these parameters arise that could compromise the health and safety of the animals. Those housed in indoor facilities where they cannot use behavioral methods to avoid adverse conditions and are not adapted to environmental extremes are especially vulnerable to extremes in temperature. • Most species of nonhuman primates evolved in warmer climates, hence, the recommended temperatures for indoor housing areas are 64 to 84°F.54 Outdoor housing facilities must provide animals with shelter to move out of wind, sun, and rain, thereby allowing animals to seek comfort or refuge away from environmentally stressful conditions. • Indoor facilities should maintain relative humidity in rooms 54 between 30 and 70%. • Lighting is often regulated by timers set to give 12 to 14 hours of light; 10 to 12 hours of darkness in indoor housing areas. It may be either natural or artificial, but should be sufficient to allow adequate inspection of animals and good housekeeping practices. Providing full-spectrum light indoors will promote synthesis and absorption of vitamin D3, which New World monkeys cannot synthesize from vitamin D2.55 Light timers should be regularly checked to ensure correct function. Breeding activity of species with seasonal breeding cycles (i.e., Macaca mulatta and Saimiri sciureus) may be affected by changes in the light-dark cycle. • For animals housed inside, improper room ventilation can influence the spread of aerosol-borne diseases such as tuberculosis. The relative air pressure between rooms housing nonhuman primates and ancillary areas such as corridors should be such that the animal room is negative to adjacent areas. All air entering a room that houses nonhuman primates should be completely 54 exhausted with no air recycled through other areas of a facility. The ventilation rate should be sufficient to prevent buildup of odors, and remove excess heat and humidity without creating drafts and excess noise. Because animals produce heat, ventilation needs for rooms with large numbers of animals will be increased as compared to rooms housing few animals.

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• Nonhuman primate housing areas often have increased noise levels as compared to areas housing rodent species. Rodents, rabbits, and quieter animals such as cats should be located in rooms distant from nonhuman primate areas.54

Environmental/Psychological Enrichment Methods used to improve the environment of captive nonhuman primates and enhance their psychological well-being are termed environmental enrichment.55 Nonhuman primates have well-developed social behaviors due to their natural lifestyles that include living in pairs or groups. Programs to address the psychological well-being of captive nonhuman primates must address both social and environmental means of enriching these animals’ lives. Each plan should include:

1. Methods to address social interaction Social contact can be achieved through housing systems that allow groups to live together. Groups of animals are often housed together in outdoor corrals, corncribs, and large indoor/outdoor runs or rooms equipped for gang housing (Figure 2.9). Methods of group housing include: Gang housing Social groups are best formed with animals that are not familiar with each other. This limits the formation of coalitions among the animals, which can result in severe aggression. Care must be taken when removing animals from an established group in order to preserve social stability between the remaining animals. Removal of the highest socially ranked female from a stable group of macaques often results in attempts by lower ranked females to elevate their social status. Coalitions of lower ranked animals may fight with the remaining higher ranked animals, and individuals may become severely injured. Newly assembled social groups of animals should be closely observed to ascertain that all animals are able to maintain themselves in the group. Early identification of injured animals can reduce the occurrence of serious complications resulting from wounds and trauma. Pair housing Pairing nonhuman primates provides them with social interaction which, is beneficial for psychological well-being when gang housing © 2002 CRC Press LLC

Fig. 2.9 Group housed macaques. (Courtesy of S. Pearson, Covance Research Products, Alice, TX.) is not feasible. Manufacturers of cages for nonhuman primates have developed cage designs that allow pair housing or attempts to pair. The key to the success of pairing nonhuman primates is identification of compatible pairs. Frequently this is achieved through the use of caging that has been modified to allow prospective pairs to have limited access to each other by means of visual contact or min56 imal contact through a mesh screen (Figure 2.10). After a short conditioning interval, personnel familiar to the animals should do the observations, so that animals will interact with each other rather than the observer. These people should have knowledge of the animals’ individual behavior in order to observe each pair’s interactions over a period of time (e.g., 30 minutes for 3 to 5 days), paying special attention to the amount of affiliative vs. aggressive behavior exhibited between the animals. Affiliative gestures include lip smacking, grooming, and nonthreatening body language such as the presentation of the hindquarters to the other animal. Aggression is usually conveyed via the open mouth display of teeth; erection of hair, especially around the head and neck; cage shaking; biting attempts; and direct eye contact between the individuals during displays of threatening body language such as bouncing. Initially,

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Fig. 2.10 Duplex macaque cage modified to allow pair housing or limited contact through a mesh screen panel.

animals are observed when only visual contact is allowed. If observations suggest potential compatibility, limited direct contact through mesh screens or bars is allowed. Finally, animals that appear to have established a compatible relationship are allowed full contact. Observations for negative interactions must be done until observers are confident that animals are compatible. Only pairs that can be fed together without fighting and who show friendly gestures towards each other should be allowed to remain together without close scrutiny. It is important to assess each pair on a regular basis to be sure that the nutrition of each animal is not compromised. Disarming canine teeth on male nonhuman primate species with large canine teeth (e.g., macaques and baboons) can help reduce the severity of lacerating bite wounds should animals fight. However,

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even female macaques are capable of inflicting severe crush wounds. Bite wounds are often not readily visible, and observers must be familiar with the animal’s individual behavior and report those that appear depressed or avoid interactions with their cagemate. Additional considerations for social interaction Even animals that cannot be offered direct access to other animals can benefit from contact via visual, auditory, and mesh screen routes. Every consideration should be made to avoid housing nonhuman primates in isolation from others of their own or compatible species. When receiving new animals into quarantine, care should be taken to arrange shipments of more than one animal in order to avoid social isolation, although the animals are usually individually housed during quarantine for preventative health considerations. Creative use of mirrors (Figure 2.11) and clear plexiglass barriers may allow individually housed animals visual contact when further contact with other animals is not possible.

Fig. 2.11 Mirror on rolling stand. © 2002 CRC Press LLC

2. Opportunities to engage in species-typical behavior Many factors are thought to influence activities in nature, including the type of social organization, mating system, group size, group composition, spacing patterns, patterns of emigration and immigration, habitat characteristics, locomotor patterns, sleeping places, daily activity patterns, food availability, feeding patterns, reproductive cycles, parental care, communication, age of puberty, and normal postures for resting and sleeping.55 Environmental enrichment often focuses on providing animals with novel ways to interact with their surroundings through the use of toys or other objects placed inside or attached to their cages, and ways to increase their activity by providing food foraging opportunities. Increasing animals’ time spent foraging provides stimulation to all sensory organs, and carestaff should be encouraged to use their ingenuity when developing foraging strategies. Prior to implementing extensive new enrichment programs, limited trials of new enrichment devices and taste tests of foraging foods should be performed to ensure the safety and suitability of the items. The cost of items for foraging should be factored into the husbandry management budget for nonhuman primates. Foraging for food is a normal behavior of nonhuman primates that accounts for a significant amount of their daily activity in the wild. Captive animals may be induced to forage for natural foodstuffs such as seed mixtures, fruit, or vegetables, as well as various candies, dog 59 biscuits, and other processed foods such as cereals (Figure 2.12).

Fig. 2.12 Foraging mix components. © 2002 CRC Press LLC

The use of specialty foods rather than food biscuits has been shown to increase the amount of time animals spend doing foraging activities.60 Foraging devices provide captive animals the opportunity to express this behavior. Simple devices that require the animals to pick food out of a mixture of nonnutritive materials (such as wood chips or straw) or manipulate regular food biscuits through a maze or puzzle feeder (Figures 2.13 and 2.14), can prolong the amount of time animals work at feeding themselves.61, 62 Turf and fleece rugs sprinkled

Fig. 2.13 Macaque foraging tube.

Fig. 2.14 Puzzle feeder. © 2002 CRC Press LLC

Fig. 2.15 Baboon foraging tray. with tidbits, foraging trays (Figure 2.15), probe feeders, puzzles baited with food treats, and hanging manipulanda containing treats are other examples of foraging devices.59 All devices should be nontoxic, durable, safe, and ideally easy to clean, sanitize, and reuse. Facility managers must consider the extra labor required to provide nonhuman primates with opportunities to forage based on the number of animals and available labor. Units with small numbers of nonhuman primates have additional flexibility in devising and selecting devices. New designs of devices should be carefully assessed on small groups of animals before investing in large quantities of a particular unit. Novelty plays a role in the utilization of some devices. Stimulating new sensations such as taste, touch, and smell can increase the usefulness of a device. For species such as marmosets and tamarins, that scent mark as a method of communication, sanitation programs should be designed to avoid removing all scent-marked territorial objects such as the perches and nest boxes, because to do so creates unnecessary distress in these animals.63 Providing auditory stimulation is another way to enrich an animal’s environment. Radios or compact disc players can be used to provide a variety of natural or soothing sounds that may help provide a medium to reduce the impact on animals of other noise such as from ventilation systems, plumbing, and metal equipment.

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3. Housing that permits suitable postural and locomotor expression There are numerous citations in the literature describing a variety of nonforaging enrichment methods that permit animals to express normal postural and movement activity.64, 65 Perches, swings, chew toys, and tree branches are examples of items that enrich the animals’ environment and provide psychological enrichment. Caging should include perches and other enrichment devices, affixed to or placed inside, that address the animals’ need to express normal postures as well as play with and manipulate objects. Devices should be designed to be removable from outside the cage or oriented so that they do not interfere with the operation of the squeeze-back mechanism within standard cages. Perches are often built into the cage of the same materials as the cage (Figure 2.16). However, they can be made of polyvinyl chloride (PVC) pipe for small nonhuman primates (Figure 2.17) or branches from nontoxic trees may be used. Toys can be made of various materials, such as PVC pipe, various nontoxic plastics, or rubber used in chew toys and balls (Figure 2.18). Mirrors of coated plastic or stainless steel are often well received, as are nontoxic tree branches. These various manipulanda can all be satisfactory, depending on the management of the animals and equipment involved. Items should be easily sanitizable. Toys and equipment should be regularly inspected for sharp edges and excessive wear. A program for replacing worn items can allow incorporation and assessment of novel items into the enrichment program.

Special Considerations Developing enrichment programs should be done in consultation with persons experienced in primate behavior and familiar with the facility and its available resources to support animals with individual needs. According to the Animal Welfare Regulations,66 the enrichment program needs to address the following considerations: • Programs need to address the special social needs of infants and juvenile nonhuman primates. Young nonhuman primates must learn appropriate social responses after birth initially from their parents; and as they grow more independent, from conspecifics and peers. Addressing the needs of these animals requires special knowledge and careful planning because their social skills develop over an extended period of time (i.e., years). © 2002 CRC Press LLC

Fig. 2.16 Collapsible perch for macaques.

Fig. 2.17 Polyvinylchloride pipe perch.

Fig. 2.18 Enrichment devices. © 2002 CRC Press LLC

• Programs need to address nonhuman primates with observable behavioral abnormalities and those that appear to be in psychological distress. Animals with abnormal behaviors may be responding to perceived stress in the only way they can, vs. how animals with better-developed social skills might react to the same event. Care and experience are required to maintain animals with abnormal behaviors because they may have abnormal responses to what may appear to be nonthreatening situations or novel stimuli. • Programs must address the needs of animals used in Institutional Animal Care and Use Committee-approved research that requires restricted activity. • Programs must address the needs of individually housed animals that are unable to see and hear nonhuman primates of their own or compatible species. • Programs must allow great apes weighing over 110 lb (50 kg) to express species-typical behavior.66

nutrition Dietary Requirements 67

Nonhuman primate dietary requirements have been published. Within the research setting, nutritional needs are usually met by feeding commercially available, nutritionally balanced, nonhuman primate diets (see Chapter 6). Milk replacers are available for infants, and biscuits for other life stages. Balanced nutrient liquid formulations are also available for adult animals. Semi-purified and chemically defined diets can be purchased; however, their palatability should be assessed before feeding them for long periods. New diets to reduce the formation of dental calculus and provide high fiber have also recently been developed for nonhuman primates. Key concepts with respect to nonhuman primate nutrition include: • Old World species are usually fed a diet containing approximately 15% (dry matter) protein, while New World species receive a diet with approximately 25% protein.

• New World species require vitamin D3 (cholecalciferol), while Old World species can metabolize vitamin D2 (ergocalciferol) into vitamin D3. Commercial diets are supplemented with vitamin D3. © 2002 CRC Press LLC

• All nonhuman primates require vitamin C, a nutrient that is quite perishable. Ascorbic acid is oxidized in aqueous solutions by dissolved oxygen; hence, feeding biscuits soaked in milk, fruit juice, or water may require that animals be supplemented with additional vitamin C. It is also degraded by exposure to light, elevated temperatures, and metals that act as catalysts, (e.g., copper). Consequently, it is recommended that properly stored biscuits containing vitamin C should be used within 90 days of milling. • Although primates may actually consume 3 to 5% (dry weight) of their bodyweight on a daily basis, there is considerable wastage. Aged animals with arthritis or those with missing digits may have trouble obtaining biscuits out of J-type feeders. Those animals with few teeth or dental disease may be reluctant or unable to chew hard biscuits. Carestaff must be alert to these potential problems. Marmosets and tamarins should be fed from elevated feeding stations. Feeding smaller amounts of food several times a day reduces wastage and more closely mimics the animals’ feeding pattern in the wild. Small New World species are especially prone to hypoglycemia resulting from prolonged fasting periods and require access to food throughout the day. Creating foraging opportunities that utilize the biscuits through the use of overhead grid feeders can increase the amount of time it takes animals to feed as well as minimize wastage; however, this may not 55 be appropriate for all ages of animals. • Animals that are group housed require multiple, well-spaced feeders to allow all animals daily access to enough food for adequate nutrition. New pairs of animals should be observed to ensure that each animal has access to the food. Dominance, either physical or social, can prevent low-ranking nonhuman primates from accessing food. Providing sight barriers around feeding stations may improve situations in which dominant animals monopolize access to food through visual threats.

Novel Foods and Foraging Treats Novel foods are often incorporated into nonhuman primate feeding regimens. Food preferences vary among species. Several key principles when using novel food and foraging treats should be kept in mind:

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• Fresh fruits and vegetables need to be thoroughly washed prior to feeding, particularly those items that may be contaminated with soil that can harbor pathogens from fecal origin. However, it is preferable to allow the animals to peel the items or open nuts and seeds at will. Suppliers of fresh foods to grocery stores are a good source of clean, fresh fruits, vegetables, and nuts. • Pet and farm supply stores carry bird food mixes and grain items that are attractive to most commonly used nonhuman primate species. • Mealworms, cooked eggs, crickets, and neonatal mice have been used as supplemental feed for New World species. Because mice can carry the lymphocytic choriomeningitis virus, which is transmissible to the marmosets and tamarins, many institutions housing these species prefer to use other supplemental foodstuffs. • The selection of foodstuffs should take into consideration the type of waste management system a facility uses for its nonhuman primates. Care should be taken to protect drains from becoming clogged with waste from fresh produce. Promptly removing waste minimizes the potential pest control problems associated with moist garbage remaining in the animal rooms. • Insects and other vermin can be brought into a facility with fresh foods. Any foodstuffs, and the containers or bags they arrive in, should be carefully inspected upon receipt prior to entering a facility. Containers or bags that are open, soiled, or wet should not be accepted. • Open bags of food must be stored inside covered containers with tight-fitting lids. These containers should be sanitized on a regular basis.

Potable Water In general, water is supplied to animals continuously via automatic watering systems or water bottles. If water cannot be provided continuously, animals must have access to water for a minimum of 1 hour twice daily, unless otherwise required by the attending veterinarian or an Institutional Animal Care and Use Committee-approved research proposal.66 If water is supplied by automatic systems, animal care personnel must be trained to check the function of these devices on a regular basis to ensure that water is being delivered to

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the animals. Newly received, aged, infirm, and recently weaned animals must be carefully assessed for their ability to recognize, access, and operate watering devices.

sanitation The standards contained in the Code of Federal Regulations must be followed.66 The following points are relevant to sanitation: • Daily removal of excreta and food waste from inside each indoor primary enclosure is required. Cage designs that allow waste to fall through the floor of the cage are less labor intensive to keep sanitary. Removal of excreta underneath primary enclosures is required as often as necessary to prevent animals from becoming soiled and to reduce disease hazards, insects, pests, and odors. • Complete sanitation of indoor cages (or primary enclosures), including food and water receptacles, must be performed at a minimum of every 2 weeks. Animals are not to become wet, harmed, or distressed during this activity. Acceptable methods of sanitation include live steam under pressure, washing with hot water at 180°F and soap or detergent such as in a mechanical cage washer, or washing in the room with detergent or disinfectant solutions (or a combination product) to remove organic material followed by a clean water rinse. An acid wash may be necessary to remove mineral deposits. To achieve the best sanitation, animals should first be removed from the primary enclosure and the cages then transported to a mechanical cagewasher. • Animals housed in primary enclosures containing material that cannot be readily sanitized by the above methods (e.g., outdoor enclosures) should be maintained so that contaminated material is regularly removed to prevent odors, diseases, and infestation by insects and vermin.

transportation In most instances in the United States, nonhuman primates are transported in motor vehicles with self-contained climate control units separate from the driver’s compartment, or by commercial airliners. Prior to shipping animals, one should carefully review all

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applicable laws and regulations. The following points are relevant to the transportation of nonhuman primates: • Transportation of nonhuman primates requires adherence to the standards published in the Code of Federal Regulations66 as well as those pertaining to interstate/international movement of animals if applicable. In addition, see Chapter 3 for information to comply with the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) and the United States Endangered Species Act (ESA).68, 69 • The International Air Transport Association (IATA) publishes requirements for international shipping.70 These include detailed specifications for shipping crates, isolation of animal shipments from other cargo, handling of loaded crates for personnel safety, and many additional requirements covering a multitude of issues. Airlines are not required to carry animal shipments, and many airlines have opted not to handle these shipments. Consequently, there are fewer options for shipping these animals.

Shipping Crates While it is possible to construct a shipping crate to meet the requirements listed in the Code of Federal Regulations66 or the IATA’s specifi70 cations, it is common practice to purchase disposable crates for single-use purposes from commercial producers (Figures 2.19 and 2.20). Reusable crates made of extruded aluminum can also be constructed by manufacturers of caging equipment. These more durable crates are recommended for larger species like baboons, especially those traveling by commercial airlines where the crates will by handled several times and animals will be sitting at intermediate points en route to their final destination. All crates must be large enough to allow animals to sit and lie down in normal positions. Food and water receptacles should be affixed so that they can be accessed from outside the crate. There should be no sharp edges inside the crates and openings should not permit animals to put any bodypart outside the crate. Absorbent, nontoxic bedding should be provided if animals are to be housed on solid floors. Crates with slotted or mesh floors must have a solid tray beneath to prevent waste from flowing outside the crate. Ventilation openings must meet the size and location stipulations of 66 70 the Animal Welfare regulations or IATA requirements if applicable. Crates must be constructed so that these openings cannot be occluded by placing the crates in contact with each other. Signage required on the crates includes feeding and watering instructions specifying when © 2002 CRC Press LLC

Fig. 2.19 Disposable shipping crate for a baboon or large macaque.

Fig. 2.20 Disposable double-unit shipping crate for small nonhuman primates.

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animals were last offered food and water (no less than 4 hours prior to shipment) and that water must be given every 12 hours and food every 24 hours during transport. Lettering at least 2 inches high specifying “Wild Animals” with a directional indication for which side is “Up” must be present. Crates should be labeled with the shipping address of the consignee, including the consignee’s contact person. This signage should be posted on two sides of each crate. The health certificate and shipping forms are to be affixed to the crate in such a way that the documents can be easily examined en route if necessary.

Certificates of Health and Acclimation Status Health certificates A health certificate is required for interstate/international movement of animals. Generally, tuberculin skin testing should have been performed within a reasonable interval (30 days) prior to shipment, unless contraindicated. The United States Department of Agriculture (USDA) provides Health Certificates for interstate and international shipping of small animals, including nonhuman primates. These are obtained from the state veterinarian. When state health certificate forms are used, they must be completed by a USDA-accredited veterinarian. In addition, the Animal Care branch of the Animal Plant and Health Inspection Service within the USDA requires that APHIS Form 7020, entitled “Record of Acquisition, Disposition or Transport of Animals (other than dogs and cats)”, must be completed and accompany shipments of nonhuman primates from one institution to another for the purpose of sale, transfer, or exchange. This form can be downloaded from the United States Department of Agriculture’s Animal Care Web site by Internet users (http://www.aphis.usda.gov/ac/forms.html).

Certificates of acclimation Certificates of acclimation to temperatures less than 50°F must be made by a veterinarian within 10 days of shipping if shipping is to be handled by carriers that cannot provide conveyances that maintain 66 temperatures between 45 and 85°F at all times. These certificates must state a minimum acceptable lower temperature that is within the professionally accepted range for the nonhuman primate, considering its age, condition, and species. The consignor’s name and address, the number of nonhuman primates contained in each primary enclosure, and the species of nonhuman primate should be included on the certificate. © 2002 CRC Press LLC

recordkeeping Recordkeeping generally involves those records pertaining to individual animals, groups or colonies of animals, and records to ensure institutional compliance.

Individual Animal Records These records should contain the following information: • The animal’s unique identification, date of birth or date of receipt, physical examination findings, laboratory results (e.g., blood tests, urinalysis, fecal flotation, fecal smears, fecal cultures), dates and results of tuberculin screening tests, dates when weighed, and a summary of the animal’s health history as it pertains to the animal’s usage and health or behavioral problems. • Recently, additional recommendations for contents of health records have been circulated by the USDA.71 Diagnosis, prognosis, and treatment plans are requested when appropriate, along with the corresponding dates, details, and results of all medically related observations, tests, and treatments. Name, dosage, route of administration, frequency, and duration of all treatments should be documented. Dates of resolution, and criteria or schedule for reevaluation should be indicated. If animals have activity restrictions, veterinarians should document these in the health record. • All animal health records must be held for at least 1 year after the animal’s disposition or death. Animals transferred to another party should be accompanied by a copy of that animal’s health record, which contains the individual’s medical history, information on any chronic or ongoing health problems, and information on the most current preventative medical procedures.71

Group/Colony Records Records of breeding animals may have breeding histories and genetic analysis to facilitate the selection of genetically unrelated breeding pairs. Documentation of social rank, if animals are group housed, can be an important component of managing groups of animals to maintain a stable social hierarchy.

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Recordkeeping can be designed around the needs of the individual institution housing nonhuman primates. Often, periodic reports need to be generated that reflect the animals’ usage. Computerized database management programs can be used to format the desired information to provide periodic reports. Census data, project histories, experimental schedules, clinical and surgical case histories, reproductive indices, and financial statements are examples of the types of reports that are often made on a regular basis.

Institutional Recordkeeping Institutional recordkeeping is usually designed to follow the format established in the table of contents of the Guide. The Association for the Assessment and Accreditation of Laboratory Animal Care, International, provides a document entitled “Evaluation of Institutional Animal Care and Use Program”; this document outlines major areas where recordkeeping is utilized and duplicates the format of the Guide. It details information that the records should contain. For those facilities that purchase animals through a vendor that has imported the animals for the purposes of education, research, or exhibition, records of disposition of animals must be maintained in 72 accordance with Centers for Disease Control specifications. Noneducational, nonscientific, or nonexhibition use of these animals is not permitted.

identification Methods of individual identification of nonhuman primates can be classified as permanent or temporary.

Permanent Methods Permanent methods include tattoos, microchip implants, and freezebranding. The major disadvantage of all these methods is that they are difficult or impossible to read at a distance. Tattoos can fade, especially in growing animals, and require some practice to produce a legible number. Microchips can be removed or migrate from the site of implantation, are expensive if large numbers of animals must be identified, and require a scanner capable of reading the chip. Freezebranding is limited by the variable depth of tissue affected, is difficult to see on long-haired species, and is therefore not commonly used.

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Temporary Methods Temporary identification methods include hair dyes, hair-clipping, or neck collars with identification tags attached. All three methods may allow the identification of animals at a distance. Disadvantages of these methods are that hair dyes must be replaced seasonally; hairclipping is for short-term identification and may not work well if animals have sparse haircoats; and neck collars with identification tags attached may be lost or defaced and can present a hazard to the animals wearing them. Neck collars and tags are most frequently used on squirrel monkeys and outdoor-housed baboons. Often, several methods of identification are used, such as tattooing along with either hair dyeing, microchip implants, or neck collars and tags.

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3 MANAGEMENT introduction The management of a facility housing nonhuman primates is affected by numerous governmental regulatory agencies, including those at the federal, state, and local levels. It is not uncommon for state and/or local agencies to have their own regulations and requirements. It is therefore important that a facility maintaining nonhuman primates for the purposes of research, testing, and teaching not only be managed in accordance with the federal regulatory agencies and accrediting organizations listed below, but also in accordance with state and local laws.

regulatory and accrediting agencies, and compliance The United States Department of Agriculture (USDA) The Animal Care division of the Animal and Plant Health Inspection Service (APHIS) within the USDA has the responsibility for enforcing the Animal Welfare Act (AWA). The AWA was originally passed in 1966 as the Laboratory Animal Welfare Act (P.L. 89-544) and has since been amended on several occasions (P.L. 91-579, P.L. 94-279, P.L. 99-198, P.L. 101-624).73 The regulations and standards promulgated to implement the AWA establish standards for the humane care of laboratory animals, including nonhuman primates. As part of the 1985 amendment to the AWA, regulations were written requiring

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institutions using “covered species” in research, testing, and teaching to have an Institutional Animal Care and Use Committee (IACUC). In addition, standards were written as part of the 1985 amendment to the AWA requiring dealers, exhibitors, and research facilities to develop, document, and follow an appropriate plan for environmental enhancement of primates. Components of the environmental enhancement program should include social housing of compatible animals, enrichment of the environment, limitations on the use of restraint devices, and special considerations that address the needs of various age groups of animals or animals displaying abnormal behavior. • The specific requirements of the AWA are found within the Regulations and Standards of the Animal Welfare document.66 • Registration with the USDA and adherence to the USDA Regulations is required of all institutions using nonhuman primates in research, testing, or teaching in the United States. • Compliance with the USDA Regulations and Standards is monitored by an active inspection program that involves the unannounced periodic inspection (at least annually) of registered facilities by a representative of the Animal Care division of APHIS. • The USDA requires registered facilities to submit an annual report on animal usage covering the federal fiscal year, October 1 to September 31. The report is due by December 1.

The National Institutes of Health (NIH), Public Health Service (PHS) The Office for Laboratory Animal Welfare (OLAW) NIH, is responsible for the implementation and administration of the Public Health Service Policy on the Humane Care and Use of Laboratory Animals (PHS Policy). The PHS Policy implements the Health Research Extension Act of 1985 (P.L. 99-158).74 The PHS Policy is based on the United States Government Principles for the Utilization and Care of Vertebrate Animals Used in Testing, Research and Training (IRAC Princi75 ples). Standards for institutional programs and facilities are 54 described in the Guide. It is required that each institution receiving PHS funds to support or conduct research involving live vertebrate animals submit to and have approved by OLAW a written Animal Welfare Assurance setting forth compliance with the PHS Policy. Assurances are evaluated by © 2002 CRC Press LLC

OLAW to determine the adequacy of the institution’s proposed program for the care and use of animals. Approval of an Assurance can be for a period of time up to 5 years. Adherence to the PHS Policy is primarily achieved through voluntary compliance.

The United States Food and Drug Administration (FDA) and the Environmental Protection Agency (EPA) Pertinent regulations are described in the Good Laboratory Practices (GLP) for Nonclinical Laboratory Studies CFR (Code of Federal Regulations) Title 21 (Food and Drugs), Part 58, Subparts A-K; CFR Title 40 (Protection of Environment), Part 160, Subparts A-J; CFR Title 40 (Protection of Environment), Part 792, Subparts A-L.76 GLP regulations establish standards for the proper conduct of nonclinical studies that use animals for safety studies funded or reviewed by the FDA or EPA. For the FDA, this includes studies involving food and color additives, animal food additives, human and animal drugs, medical devices for human use, biological products, and electronic products; whereas for the EPA, this includes studies involving pesticides, herbicides, and fungicides. In general, GLP regulations require institutions to have in place written standard operating procedures, that are rigorously followed and supported with detailed records. Compliance is assessed through an active program of periodic inspections performed by representatives from the respective agencies.

The Centers for Disease Control (CDC) The CDC is the organizational unit within the PHS that is responsible for preventing the introduction of communicable disease into the United States. According to the Foreign Quarantine Regulations (CFR, Title 42, Part 71), only facilities registered as importers with the CDC can import nonhuman primates into the United States.72 Facilities not registered with the CDC cannot receive imported nonhuman primates until after they have completed a 31-day quarantine period at a registered importation facility. These regulations also stipulate that the importation of nonhuman primates within the United States is restricted to scientific, educational, or exhibition purposes and specifically prohibits importation and distribution for use as pets. Importers must document the intended use of nonhuman primates prior to distribution, maintain detailed records, and report certain diseases (monkey pox, yellow fever, and hemorrhagic 77 fevers) within 24 hours of their recognition. © 2002 CRC Press LLC

The Fish and Wildlife Service (FWS) The FWS is the unit within the Department of Interior that regulates the trade and transportation of nonhuman primates. In this role, the FWS is responsible for enforcing the Lacey Act, the United States Endangered Species Act (ESA) and the Convention on International Trade in Endangered Species of World Fauna and Flora (CITES). The Lacey Act (P.L. 97-79)78 regulates the transportation of wild mammals and birds imported into the United States. To promulgate the Lacey Act regulations, the FWS integrated the existing live animal transportation requirements of the CITES, the International Air Transport Association, and the United States Animal Welfare Act.79 The Lacey Act is a very powerful law in that failure to comply with any federal, state, or foreign law pertaining to the transportation of animals is a violation of the Act. The ESA (P.L. 93-205)68 was passed in 1973 to prevent the extinction of various plant and animal species as determined by the Office of Scientific Authority of the FWS. It regulates/prohibits the import, export, capture, and interstate or foreign commerce of endangered and threatened species unless authorized by a permit from the FWS.69 The ESA applies equally to live or dead animals or plants, their progeny, and parts or products derived from them. Many nonhuman primates are federally listed as endangered or threatened under the ESA. Interstate sale of listed species is allowed only under a permit from the FWS. Permits and information pertaining to species federally listed as endangered or threatened can be obtained through the U.S. Fish and Wildlife Service’s Office of Management Authority. The CITES treaty is the most frequently cited international law affecting nonhuman primates. The U.S. Fish and Wildlife Service’s Office of Management Authority acts as the U.S. Management Authority for CITES. The CITES categorizes animals into appendices (I, II, and III) based on the level of vulnerability to extinction a species faces by virtue of its international trade. Species listed in Appendix I are those presently threatened with extinction; whereas those listed in Appendix II are not presently threatened with extinction, but may become so unless their trade is regulated. Species listed in Appendix III are those that are regulated domestically to prevent exploitation. To import species listed in Appendix I, an export permit from the country of origin and an import permit from the FWS must be obtained; whereas species listed in Appendix II require only an export permit from the country of origin. The CITES lists all nonhuman © 2002 CRC Press LLC

primates in either Appendix I or II, with the majority of species used in biomedical research being listed in Appendix II.79

Association for the Assessment and Accreditation of Laboratory Animal Care International, Inc. (AAALAC) AAALAC is a nonprofit organization designed to provide peer reviewbased accreditation of animal research programs. Accreditation by AAALAC is based on an institution’s adherence to the principles outlined in the Guide. Accreditation is voluntary and involves an on-site assessment of an institution’s animal care and use program at least once every 3 years. Annual reports and program changes are required. Full accreditation by AAALAC is viewed favorably by the Office of Laboratory Animal Welfare of the National Institutes of Health when determining whether an institution complies with the PHS Policy.

institutional animal care and use committee (IACUC) The IACUC plays an integral role in ensuring that nonhuman primates used in research, testing, and teaching are used appropriately and receive proper care and humane treatment. In addition, the IACUC plays an important role in ensuring that the institution complies with various governmental regulatory agencies and accrediting organizations. The composition and responsibilities of the IACUC are described in detail in the Animal Welfare Act, PHS Policy, and the Guide.54, 66, 75 Important points to note about the composition of the IACUC include: • The USDA Regulations require an IACUC to be composed of a minimum of three members; whereas the PHS Policy requires a minimum of five members. Committee members are appointed by the Chief Executive Officer of the institution. • Both the USDA Regulations and the PHS Policy require the IACUC to have a chairperson. • Both the USDA Regulations and the PHS Policy require that at least one member of the IACUC be a Doctor of Veterinary Medicine with training or experience in laboratory animal medicine or science and with responsibility for activities involving animals at the facility. © 2002 CRC Press LLC

• Both the USDA Regulations and the PHS Policy require that at least one member of the IACUC be nonaffiliated with the institution in any capacity other than as a member of the IACUC. • The PHS Policy requires that at least one member of the IACUC be a practicing scientist who is experienced in research involving animals. • The PHS Policy requires that at least one member of the IACUC have primary concerns that are in a nonscientific field. This person may be an employee of the institution served by the IACUC. • According to the PHS Policy, one person may fulfill more than one of the above requirements; however, the IACUC must still consist of a minimum of five members. • According to the USDA Regulations, an IACUC cannot have more than three members from the same administrative unit of the research facility. The written regulations should be consulted for an in-depth description of IACUC responsibilities.66, 75 In general, the IACUC is charged with the following: • Review at least once every 6 months the institution’s program for the humane care and use of animals to ensure that it complies with the requirements of the PHS Policy and the Animal Welfare Regulations and Standards. • Inspect at least once every 6 months all of the institution’s animal facilities (including study areas and satellite facilities) to ensure that it complies with the requirements of the PHS Policy and the Animal Welfare Regulations and Standards. • Prepare reports of the semi-annual evaluations described above and submit them to the Institutional Official. The report must contain a description of the nature and extent to which the facility adheres to the Animal Welfare Act, the PHS Policy, and the Guide. Reports are to be reviewed and signed by a majority of the IACUC members and must include any minority views. • Review and investigate concerns involving the care and use of animals at the institution. • Make recommendations to the Institutional Official regarding any aspect of the institution’s animal program, facilities, or personnel training.

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• Review and approve, require modifications to secure approval or withhold approval of proposed protocols and/or significant changes to previously approved protocols involving the use of animals in research, teaching, and testing. • Ensure that personnel are adequately trained and qualified to perform procedures on animals. • Ensure that the animals are properly handled and cared for. • Ensure that the investigator has considered alternatives to potentially painful or stressful procedures, and that the research does not unnecessarily duplicate previous experiments. • Ensure that sedatives, analgesics, and tranquilizers are used to relieve pain and distress, whenever possible. • Ensure that activities that involve surgery include appropriate provisions for pre-operative, intra-operative, and post-operative care. • Ensure that animals are euthanatized appropriately by personnel trained in the methodology.

occupational health and safety The PHS Policy, the Guide, the CDC, and the Occupational Safety and Health Act require that an occupational health and safety program be in place for personnel working with nonhuman primates. 54, 75, 80, 81 An effective occupational health and safety program should involve the following groups working together to achieve this goal: the IACUC, the veterinary and animal carestaff, the investigative staff, the occupational health staff, the environmental health and safety staff, and the institution’s administration. In-depth descriptions of comprehensive occupational health and safety programs can be found in the literature.54, 81–83 The use of nonhuman primates in the laboratory environment creates some unique considerations when developing an occupational health and safety program. Unlike most animals used in research, many nonhuman primates are obtained from the wild or large outdoor colonies. Animals from such sources can be asymptomatic carriers of a variety of zoonotic diseases, such as Cercopithecine herpesvirus 1 (B virus), shigella, and giardia. Moreover, the intelligence, agility, strength, and aggressive nature of nonhuman primates and the nature of the caging equipment (heavy with moving components) © 2002 CRC Press LLC

used to maintain them increase the potential for physical injury to staff. Although it is not within the scope of this chapter section to cover all facets of an occupational health and safety program, some important aspects, as they relate to the use of nonhuman primates in a research environment, are presented.

Training Personnel handling and/or performing experimental manipulations on nonhuman primates must be properly trained. Training should include a review of the institution’s occupational health and safety program; a review of zoonotic concerns; a review of appropriate protective equipment; a review of how to handle biological samples; a review of the behavioral aspects of the species utilized; a review of standard operating procedures; a review of safe work practices; and a review of appropriate handling and experimental techniques and a demonstration of the technique to be employed.

Safe Work Practices There are a number of safe work practices that personnel handling and/or performing experimental manipulations on nonhuman primates should follow to minimize injury and/or exposure to zoonotic diseases. Some of these safe work practices are briefly described below; further details can be found in the CDC’s Biosafety in Microbiological and Biomedical Laboratories and the NRC’s Occupational Health and Safety in the Care and Use of Research Animals.80, 81 • Access to nonhuman primate rooms should be restricted to individuals trained in the inherent risks and hazards associated with nonhuman primates. • Extreme care should be taken when using needles and syringes and/or other sharps during procedures involving nonhuman primates. Needles should not be recapped, but should be placed directly into a sharps container when no longer in use. Whenever possible, syringes with retractable needles should be used. • Personnel handling or manipulating nonhuman primates should wear appropriate protective clothing. • Personnel should keep their hands and other objects away from their mouth, eyes, and nose. • Personnel should wash their hands following removal of protective gloves. © 2002 CRC Press LLC

• Personnel should never eat, drink, smoke, handle contact lenses or apply cosmetics, or take medicine in areas in which nonhuman primates are used or maintained. • Procedures such as cage sanitization should be performed carefully to minimize splashing. • Surfaces (such as treatment tables) that come in contact with nonhuman primates and their bodily fluids and excretions should be appropriately disinfected.

Personal Protective Equipment The purpose of personal protective equipment is to provide a physical barrier to prevent exposure of the skin, eyes, and mucous membranes to potentially infectious materials; to prevent the contamination of clothing; and to minimize physical injuries such as scratches and bites. Personnel handling or manipulating nonhuman primates should wear as a minimum a mask, protective eye wear, disposable gloves, and a long-sleeved gown or laboratory coat.81, 84, 85 The type of protective eye wear (face shield, wrap-around protective glasses, or other devices) should be selected based on the type of work to be performed, the proximity to the nonhuman primate, and the need to provide staff with a clear field of vision. This basic level of protective equipment may need to be further augmented, depending on the experimental protocol, the species of nonhuman primate, and the technique/manipulation being performed. For example, the authors recommend double-gloving when handling or manipulating macaques. For more information on additional protective clothing, see the chapter section on restraint in Chapter 5.

Physical Injuries Nonhuman primates are intelligent, agile, strong, and in many cases quite aggressive. These characteristics, in conjunction with the need to move heavy caging equipment and the experimental methods employed in a research environment, put staff at risk for a variety of physical injuries, including bites, scratches, cuts, needle sticks, and back injuries. All injuries that occur should be reported to an appropriate supervisor and the proper procedures as outlined in the institution’s occupational health and safety program should be initiated. In addition to training, following safe work practices, and wearing appropriate protective clothing, there are a number of additional steps that can be taken to minimize physical injuries when working with nonhuman primates: © 2002 CRC Press LLC

• Whenever possible, the direct handling of nonhuman primates should be minimized. • The canine teeth of adult male nonhuman primates should be disarmed. • Whenever a nonhuman primate is sedated, the nails should be checked and trimmed if warranted. • Broken cages and equipment should be identified and taken out of service. • Equipment with sharp edges or corners should be identified and taken out of service. • Manual or hydraulic lifts should be used when stacking cages.

B Virus (Cercopithecine herpesvirus 1) Exposure Institutions that maintain macaques should develop, implement, and strictly enforce standard operating procedures to minimize potential B virus exposure based on previously published guidelines.85–87 Moreover, when a potential B virus exposure does occur, an institution should be prepared to manage the case promptly. To facilitate the prompt management of a B virus exposure, an institution should have a readily accessible B virus exposure station, as well as written standard operating procedures for the appropriate cleansing of the exposure site, the assessment and referral of the exposed employee to a knowledgeable health care professional, and the collection and submission of biological samples for diagnostic purposes. Ideally, a copy of the written procedures should be maintained at the B virus exposure station as well as by the responsible health care professional. For an in-depth description on the prevention, treatment, and risk assessment of individuals potentially exposed to B virus, refer to the CDC’s Guidelines for the Prevention and Treatment of B-virus Infection in Exposed Persons.84 Briefly, these guidelines recommend the following: • Employees should report all bites, scratches, or other injuries (no matter how trivial) that involve macaques or items potentially contaminated with macaque bodily fluids. In addition, employees should report mucous membrane exposure to macaque bodily fluids. • Decontamination procedures should begin immediately after exposure. For bites, scratches, and other injuries, the wound

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should be thoroughly cleansed with a povidone-iodine or chlorohexidine scrub for at least 15 minutes. Dakin’s solution, a dilute (0.25%) hypochlorite solution can also be used; however, it has a short shelf life and can cause local tissue injury, such as a chemical burn in people with fair skin. • If a mucosal surface, such as the eyes, nose, or mouth, has been exposed, the site should be immediately irrigated with sterile saline or tepid flowing water for at least 15 minutes. • Following appropriate cleansing of the exposure site, serum samples and viral cultures should be obtained for diagnostic purposes from both the exposed individual and the macaque involved in the incident. In the case of the exposed individual, the exposure site is cultured; whereas with the macaque, the conjunctiva and oral cavity are cultured. • The exposed individual should be evaluated by a health care professional and a risk assessment performed based on the route of the exposure, the severity of the wound, and the location of the exposure. Based on the risk assessment, the health care professional will determine if antiviral prophylaxis is warranted and propose an appropriate follow-up monitoring program. It should be noted that the use of prophylactic antiviral agents in an asymptomatic person awaiting laboratory tests is controversial and is not recommended in most circumstances. For assistance in case management, health care professionals should consult with the Viral Exanthems and Herpesvirus Branch, Division of Viral Diseases, Center for Disease Control and Prevention, Atlanta, GA 30333 (Telephone: 404-639-3629). Additional information on the collection, handling, and shipment of samples for B virus analysis can be obtained from the NIH B Virus Resource Laboratory, Viral Immunology Center, Georgia State University, 50 Decatur Street, Atlanta, GA 30303, (Web site: http://www.gsu.edu/bvirus). Specific requirements for the proper packaging and shipping of diagnostic specimens potentially containing B virus can be found in 42 CFR Part 72 - Interstate Shipment of Etiologic Agents.88

Allergic Reactions Allergic reactions to animals are among the most common conditions that affect the health of animal carestaff; however, unlike the case with many of the rodent species, sensitization to nonhuman primates is uncommon.81 Staff working with nonhuman primates can become © 2002 CRC Press LLC

sensitized to latex gloves and develop contact urticaria. Contact urticaria is characterized by red, itchy skin with raised circumscribed lesions (welts or hives). Individuals exhibiting such signs should seek assistance from an occupational health specialist. Latexsensitized individuals should wear rubber or vinyl gloves when handling nonhuman primates.

Experimental Hazards Some research protocols may require the purposeful infection of a nonhuman primate with a known human pathogen, the administration of a chemical hazard, or the administration of a radioisotope. Prior to initiation of such projects, the facility manager, veterinarian, and representatives from occupational health and medicine, and environmental health and safety should develop and implement standard operating procedures for the safe handling of the hazardous material and the treated animal.

zoonoses Zoonoses refers to diseases of animals that can be transmitted to man under natural conditions. Nonhuman primates used in research come from a variety of sources, including their natural environment, island colonies, and large corrals. Even with quality preventive health programs, animals from such sources often harbor endemic diseases, some of which are zoonotic. Moreover, the close phylogenetic relationship between nonhuman primates and people make them ideal models for the study of human infectious diseases. For these reasons, staff should be made aware of the zoonotic concerns associated with nonhuman primates and the preventive measures that need to be taken to minimize exposure. It is not the intent of this section to cover in-depth all zoonotic disease associated with nonhuman primates as a variety of sources are available that provide more thorough coverage of these diseases.81, 82

B Virus (Cercopithecine herpesvirus 1) B virus is the most important zoonotic disease associated with nonhuman primates because of its high seroprevalence among macaques and its near-70% fatality rate in infected humans. According to the CDC’s Guidelines for the Prevention and Treatment of B Virus Infection in Exposed Persons, all macaques should be presumed

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to be shedding B virus and should be handled accordingly. No other Old World or New World monkeys are known to naturally harbor B virus.84 There are a number of additional sources that should be consulted for further in-depth information on B virus.82, 89, 90 • It is estimated that only 2 to 3% of infected macaques are shedding B virus at any given point in time. During these periods of shedding, B virus can be found in the secretions and tissues associated with the oral, conjunctival, and genital mucous membranes and is transmitted among macaques by exposure to these secretions. In production colonies in which B virus is endemic, 97% of animals become infected by 2.5 years of age.90 • B virus is transmitted to humans primarily through exposure to contaminated saliva via bites and scratches; however, transmission has also been associated with needle stick injuries, exposure to infected tissues, scratches from contaminated cages, and ocular mucous membrane exposure. • Most macaques infected with B virus exhibit no clinical signs and are considered asymptomatic carriers. When clinical signs of B virus do occur in macaques, they typically consist of vesicular (blister-like) lesions of the oral cavity (Figures 3.1 and 3.2) and/or conjunctivitis. • The clinical signs associated with a B virus infection in humans can include a pruritic vesicle at the site of exposure, fever, headaches, flu-like symptoms, and fatigue. These early signs are followed by progressive neurologic symptoms that include numbness, ataxia, confusion, convulsions, ascending paralysis, and 81, 84 death. The CDC has very specific guidelines for the prevention of B virus infection in staff handling macaques.85 Briefly, these guidelines include the following: • Macaques should only be used for research purposes when clearly indicated. • Whenever possible, macaques seronegative for B virus should be used. • All macaques should be considered infected with, and shedding, B virus. • Handling and restraint methods that minimize the direct handling of macaques should be used (including chemical restraint). © 2002 CRC Press LLC

Fig. 3.1 B virus lesions on the tongue of a macaque. (Courtesy of G. Baskin, Tulane Primate Research Center.)

Fig. 3.2 B virus lesion on the lip of a macaque. (Courtesy of G. Baskin, Tulane Primate Research Center.)

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• Appropriate personal protective equipment, including protective eye wear, should be worn when handling macaques. • Cages and equipment used to maintain and handle macaques should be free of sharp edges. • Access to areas where macaques are maintained should be limited to individuals properly trained to avoid potential exposure. • Macaques with oral lesions suggestive of B virus infection should be isolated until the lesions have healed. • Persons who handle or come into contact with macaques should receive training in the proper methods of restraint, the use of protective equipment, the zoonotic nature of B virus, and the appropriate steps to be taken should a potential B virus exposure occur.

Tuberculosis Tuberculosis is the most important bacterial zoonotic disease associated with nonhuman primates. Most reported cases are caused by the acid-fast bacillus Mycobacterium tuberculosis. Susceptibility of nonhuman primates varies, with Old World species considered to be more susceptible to infection than New World species. Nonhuman primates can be exposed and infected with tuberculosis from humans during their capture and exportation from countries in 81, 90 which the prevalence of tuberculosis is high. • The primary route of transmission of tuberculosis is via inhaled aerosols from infected animals and tissues. Animals with tuberculosis of the gastrointestinal tract can shed the organism in their feces. • In nonhuman primates, tuberculosis classically involves the respiratory system and clinical signs may include coughing, labored breathing, and exercise intolerance. Because tuberculosis can involve a multitude of organ systems, signs may vary depending on the system affected. Some additional nonspecific signs associated with infection include chronic weight loss, diarrhea, and enlarged lymph nodes. • In people, the most common clinical signs associated with tuberculosis include a chronic cough with mucus and/or blood, weight loss, fatigue, weakness, fever, and chills. • The key to the prevention of tuberculosis is to minimize potential exposure. This is done by wearing appropriate protective

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clothing, employing safe work practices, limiting access to authorized personnel, and having a colony preventive health program that screens for the presence of tuberculosis in both the quarantine and maintenance colonies.

Bacterial Agents of Gastrointestinal Origin Diarrhea is the most common cause of morbidity and mortality in nonhuman primates, and the primary bacterial agents associated with diarrhea in nonhuman primates are considered zoonotic. Such agents include the Shigella spp., Campylobacter spp., and the Yersinia spp.81, 82

• These agents are transmitted via the fecal-oral route through direct contact with infected animals and their excretions or contaminated food and water. • In nonhuman primates, the clinical signs associated with these agents can vary from an asymptomatic carrier state to profuse diarrhea and dehydration. In the case of Shigellosis, the diarrhea may contain mucus and blood. Acute death has been seen in animals infected with Shigella spp. and Yersinia spp. • In people, these agents typically cause diarrhea, abdominal pain, and fever. In the case of Shigellosis, the diarrhea is often watery and contains mucus and blood. • The key to preventing infection with the zoonotic gastrointestinal bacterial agents is to minimize potential exposure. This is done by wearing appropriate protective clothing, employing safe work practices and good personal hygiene, and using appropriate sanitation methods and procedures. In addition, some facilities will screen nonhuman primates for the presence of Shigella spp. and Campylobacter spp. as part of their quarantine procedures.

Protozoal Agents of Gastrointestinal Origin Protozoal agents of gastrointestinal origin are a potential zoonotic concern, and nonhuman primates can serve as a natural reservoir for a number of these agents. These agents include Giardia spp., Cryp81, 82, 91 tosporidium spp., Entamoeba histolytica, and Balantidium coli. • These agents are transmitted via the fecal-oral route through direct contact with infected animals and their excretions or contaminated food and water. © 2002 CRC Press LLC

• In nonhuman primates, the clinical signs associated with these agents can vary from an asymptomatic carrier state to severe diarrhea and dehydration. Specifically, Cryptosporidium infections have been associated with intractable diarrhea and dehydration in infants and immunosuppressed animals; whereas Balantidium coli has been associated with severe ulcerative enterocolitis and death in great apes. Entamoeba histolytica has been reported to cause more significant disease, including marked diarrhea with blood and mucus in New World species, and young and immunosuppressed animals. • In people, these agents typically cause diarrhea, cramping, and abdominal pain. With an Entamoeba infection, the diarrhea may contain blood and mucus and the person may experience fever and chills. Cryptosporidium infections can cause prolonged bouts of profuse diarrhea in immunosuppressed individuals. • The key to preventing infection with the zoonotic gastrointestinal protozoal agents is to minimize potential exposure. This is done by wearing appropriate protective clothing, employing safe work practices and good personal hygiene, and using appropriate sanitation methods and procedures. In addition, some facilities will screen for the presence of Giardia spp., Entamoeba histolytica, and Balantidium coli as part of their quarantine procedures.

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4 veterinary care preventive health program A key component of veterinary care is an effective preventive health program. Each facility should have a preventive health program designed by a veterinarian with experience in caring for nonhuman primates. This section provides a basic overview of some of the common elements of a preventive health program as it pertains to nonhuman primates in both a quarantine and conditioned colony. Additional information on preventive health programs are described elsewhere.46, 92–95

Sources Nonhuman primates can be obtained from a variety of commercial and noncommercial domestic and nondomestic sources, including primary importers, domestic breeding colonies, academic institutions, and industrial corporations. Where animals are obtained will vary depending on the requirements of a facility, the needs of an investigative project, and the availability of animals. Many commercial sources of nonhuman primates can provide specific pathogenfree animals, such as macaques serologically negative for B virus and retroviruses. A list of commercial sources of nonhuman primates can be found in Chapter 6. Regardless of the source, animals should only be obtained from facilities that have in place a well-defined preventive health program, and are in good standing with the appropriate federal regulatory agencies and accrediting organizations.

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Quarantine All nonhuman primates coming into a facility should be isolated from the conditioned/existing colony for a specified period of time prior to release into the conditioned colony. The basic goals of a quarantine/conditioning program are to protect the animals in the existing colony from the introduction of infectious diseases, to protect personnel from zoonotic diseases, and to optimize the health and condition of newly acquired animals. A brief overview of some of the key components of a quarantine program for a facility obtaining nonhuman primates from a domestic source follows.

Quarantine facilities • In general, facilities that comply with the CDC recommendations for Animal Biosafety Level 2 are adequate for the quarantine of most groups of nonhuman primates.94 • Ideally, the designated quarantine area should be located in a building separate from the conditioned colony. However, due to the size and research emphasis of many institutions, this is not always feasible or practical.46 In such cases, the designated quarantine area should be located in an area of the facility that is not contiguous with the conditioned colony. • The quarantine facility/area should have a designated area for the treatment of sick animals that is separate from the treatment area of the conditioned colony. • The quarantine facility/area should have the means to restrict access to authorized personnel.

Quarantine husbandry practices • Only authorized, trained personnel who are included in an institution’s occupational health and safety program should have access to nonhuman primates in quarantine. • Personnel entering a quarantine area should wear, at a minimum, the protective clothing described in Chapter 3. • An essential component of a quarantine program is the establishment of standard operating procedures (SOPs) for all husbandry and veterinary practices in the quarantine facility/area. SOPs should be designed to minimize potential cross-contamination with the conditioned colony, minimize the potential for zoonotic disease transmission, and maximize the potential to detect disease processes. © 2002 CRC Press LLC

• Ideally, cages, equipment, supplies, and personnel should not be shared between the quarantine facility/area and the conditioned colony. The small size of some facilities may preclude such practices. In such cases where cages and equipment are shared, the cages and equipment must be thoroughly sanitized before being used outside the quarantine area. • Individuals who must work in both the conditioned colony and the quarantine area should perform their conditioned colony work assignments prior to entering the quarantine area. • Incoming groups of animals should be maintained in separate rooms during the quarantine period based upon source, arrival date, and species. • In general, nonhuman primates are maintained individually during the quarantine period except for special circumstances such as mother/infant pairs. Such maintenance practices allow the animal care staff to evaluate food and water consumption, urine and feces output, and identify individuals with signs of illness. • Although most animals are maintained individually during the quarantine period, quarantine should not preclude the animals from participating in other aspects of an institution’s behavioral enrichment program.

Duration of quarantine • The only legally mandated quarantine period applies to primary importers, which are those facilities registered with the CDC to bring nonhuman primates into the United States from foreign sources. The CDC requires primary importers to quarantine nonhuman primates from foreign sources for a minimum of 31 days prior to distribution to secondary importers.96 • Most research facilities are not registered with the CDC and are considered secondary importers. Recommendations on the duration of the quarantine period at secondary importers vary from 30 46, 92–94 to 90 days. • Decisions concerning the duration of the quarantine period should be based on the professional judgment of the veterinary staff, an analysis of the risk factors, the source of the animals, and the value of the conditioned colony. In general, short quarantine periods should be the exception rather than the rule.93

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Quarantine health surveillance • Animal care staff should evaluate animals daily during quarantine for food and water consumption, urine and feces output, and signs of illness. • Nonhuman primates entering a facility should be allowed to acclimate for at least 48 to 72 hours before initiation of quarantine/conditioning procedures. • Following acclimation, animals should be chemically restrained so that they can receive their first tuberculin skin test, be permanently identified, and undergo a complete examination by a veterinarian. • The examination may vary, depending on the species and requirements of the facility, but, in general, should include a physical examination, bodyweight, complete blood count, serum chemistry profile, fecal sample, and deep rectal cultures for enteric pathogens such as Campylobacter and Shigella spp. • As part of the initial examination process, some facilities require the submission of serum for diagnostic testing to verify/determine an animal’s serologic status against agents such as B virus and the retroviruses. In addition, some facilities require thoracic radiographs at the time of the initial examination and prior to release from quarantine. Thoracic radiographs not only serve as a method for the detection of tuberculosis, but also allow for the detection of pneumonia, parasitic diseases (hydatidosis), congen97, 98 ital defects, traumatic injuries, and osteoarthritis. • During the quarantine period, tuberculin skin tests should be administered in alternating eyelids every 2 weeks. In addition, it is recommended that animals in quarantine be administered tuberculin in a secondary site such as the skin over the abdomen. • Tuberculin skin test sites should be observed and findings recorded 24, 48, and 72 hours post-administration. All suspect or positive responses should be immediately reported to an appropriate supervisor for verification. Animals with unequivocal positive responses should be euthanatized, unless extenuating circumstances exist, and the quarantine group in which the positive tuberculin reactor was identified should start their quaran92 tine period over beginning at the time the reactor was identified. • During the quarantine period, animal bodyweights should be monitored closely. Animals should be weighed at least every 2 weeks in conjunction with tuberculin testing.

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• During the quarantine period, staff trained in the behavioral assessment of nonhuman primates should begin observing and documenting the behavioral repertoire and social interactions of the animals. Such assessment can facilitate the early identification of animals with aberrant behavior as well as facilitate the pair and group housing of animals upon release into the conditioned colony.

Conditioned Colony Health Surveillance There are a number of considerations in developing a preventive health program for a conditioned nonhuman primate colony.

Daily observations The daily observation of nonhuman primates is an essential component of a preventive health program. Moreover, animals must be observed daily in order to comply with various regulatory agencies.66, 75 At most facilities, the animal care staff perform this vital function as part of their daily husbandry activities. Animal care staff who work with nonhuman primates should be trained to recognize signs of illness as well as normal and aberrant behavior. Ideally, the same staff member should be responsible for the daily observation of specific rooms or groups of animals. This provides for greater continuity and allows the staff to develop experience with the animals, which can facilitate the detection of changes in behavior, body condition, food and water consumption, urine and fecal output, and fecal consistency.

Annual physical examination Nonhuman primates should undergo an annual physical examination by a veterinarian. In some circumstances, such as in the case of aged animals, biannual examination may be warranted. A thorough physical examination should be performed in a methodical and systematic manner. Results of the examination should be recorded and maintained in the animal’s health record. A routine physical examination should include an assessment of the following: • General condition. Prior to chemical restraint, the animal’s overall appearance and behavior should be assessed. This should include body condition, mental alertness, visual acuity and activity level.

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• Eyes. The conjunctiva should be examined for the presence of inflammation and discharge; and the eyes should be examined for the presence of opacities and traumatic injuries. • Ears and nose. The ears and nose should be evaluated for patency and the presence of discharge. • Mouth. The mouth of all nonhuman primates should be thoroughly examined for the presence of broken teeth, draining tracts, ulcerative lesions, dental tartar, and periodontal disease. In Old World species, the examination should include an inspection of the cheek pouches. • Skin. The skin and hair coat should be thoroughly inspected for wounds, masses, hair loss, and areas of inflammation. • Musculoskeletal system. The back and joints should be examined for swelling, joint enlargement, deformity, and contracture, and the musculature should be assessed for atrophy. The extremities should be closely examined for injuries, with special attention to the fingers, toes, and tip of the tail. • Lymph nodes. The superficial lymph nodes of the neck, axilla, and inguinal region should be examined. The presence of enlarged superficial lymph nodes could be indicative of neoplasia, tuberculosis, or other systemic bacterial, viral, or fungal diseases. • Thorax. The heart and lungs should be auscultated for the presence of abnormal sounds and also the absence of sound in lung fields. • Abdomen. The abdomen should be palpated for the presence of masses and distended/enlarged organs. • Uroanogenital region. The uroanogenital region should be inspected for the presence of ulcerative lesions, traumatic wounds, urine and fecal staining, and other abnormalities. Rectal palpation should be performed to assess the prostate, or uterus and cervix. • Vital signs. As part of an examination, the animal’s weight, temperature, and heart and respiratory rates should be determined. Some nonhuman primates may become excited during the restraint process and may as a result have an elevated body temperature. • Clinical laboratory tests. Clinical laboratory tests should be considered an important part of the annual physical examination. Tests routinely performed include complete blood counts, serum

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chemistry profiles, and fecal examination for parasites. In older animals, a urinalysis or fecal occult blood check may be warranted.

Tuberculin skin testing The tuberculin skin test is currently the most practical and reliable method of detecting tuberculosis in nonhuman primates. The frequency of tuberculin skin testing in a conditioned colony is determined in part by the research goals of the institution, the value of the colony, and the risk of exposure to tuberculosis. Where increased risk factors exist, such as the introduction of nonhuman primates from sources outside the facility, significant human exposure, or exposure to species that may carry the organism, quarterly testing is recommended. In a closed colony with limited exposure to humans, 92, 94, 95, 99 As part of semi-annual or annual testing may be sufficient. the tuberculin skin testing procedure, animal care staff should perform a cursory examination, obtain a bodyweight, and trim the animal’s nails. Procedure 1. The animal is appropriately restrained. 2. Test sites include the edge of the upper eyelid and/or the skin of the abdomen. The eyelid is the preferred testing site because it is easy to observe without the need to restrain the animal. An abdominal test is frequently performed in conjunction with the eyelid in quarantined animals and/or as part of the retesting process of an animal that had an initial suspect or positive response. The abdominal test site is used because it is easier to palpate for the presence of induration (firmness). 3. Mammalian tuberculin (0.1 ml) is injected intradermally into the test site with a 1/2 to 5/8 –inch, 25–27 gauge needle and a tuberculin syringe. A separate needle and syringe is used for each animal. See Chapter 5 for more information on how to administer an intradermal injection. 4. The injection site and any bruising that occurred as a result of the injection is recorded. Abdominal test sites are outlined in ink to facilitate later assessment. 5. Injection sites are observed at 24, 48, and 72 hours, and findings recorded.

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6. A positive tuberculin skin test (eyelid) may range from mild reddening, swelling, and drooping, to marked reddening, swelling, drooping, and in some cases ulceration and ocular discharge (Figures 4.1 and 4.2). A positive tuberculin skin test (abdomen) may appear as a red, raised wheal-like lesion around the injection site, which is very firm upon palpation.92, 95 7. All suspect or positive responses should be immediately reported to the attending veterinarian so that additional steps can be taken to differentiate between true and false positive responses. 8. Suspect or positive animals, and any animal(s) with which they are housed, should be isolated until further testing can determine the cause of the response. Access to isolated animals should be restricted. Fig. 4.1 Positive tuberculin skin test (animal’s right eye) de-monstrating mild swelling and drooping of the eyelid. (Courtesy of G. Baskin, Tulane Primate Research Center.)

Fig. 4.2 Positive tuberculin skin test (animal’s right eye) demonstrating ulceration/ necrosis and marked swelling and drooping of the eyelid. (Courtesy of G. Baskin, Tulane Primate Research Center.) © 2002 CRC Press LLC

Bodyweight assessment The bodyweight of a nonhuman primate can provide important information on the animal’s health and well-being. Weights should be obtained, recorded, and compared to previous weights on a regular basis. Animals in which a decrease in bodyweight of 10% or more has occurred should be reported to the attending veterinarian. The frequency with which bodyweights are obtained is determined by the requirements of the research project, the age and health status of the animals, the housing system in which the animals are maintained, and the professional judgment of the veterinary staff. In general, bodyweights should be assessed during physical examinations and tuberculin skin testing. Moreover, at many facilities, bodyweights are obtained in conjunction with cage sanitization.

Dental prophylaxis The maintenance of a healthy mouth can play an important role in the health and well-being of nonhuman primates. The oral cavity should be examined during annual physical examination and in conjunction with tuberculin skin testing. Animals with excessive tartar and periodontal disease should have their teeth cleaned and polished.

Immunization Recommendations and/or current practices for the immunization of nonhuman primates vary widely among facilities. Factors that should be considered in determining whether an immunization program is warranted include species susceptibility, colony risk, safety, 92 cost, and research interference. Immunization schedules typically follow human pediatric recommendations and specifics can be found in a number of sources.95, 100, 101 Some of the more common disease processes for which nonhuman primates are immunized include tetanus, poliovirus, and measles. • All nonhuman primates are susceptible to tetanus and it is recommended that all animals maintained outdoors be immunized against this agent. • Great apes are susceptible to poliovirus and it is recommended that they be immunized against this agent as part of their preventive health program. • Measles is highly transmissible between humans and nonhuman primates. Nonhuman primate colonies exposed to large numbers © 2002 CRC Press LLC

of humans are at an increased risk of exposure and may warrant prophylactic immunization. Both human measles and canine distemper/measles vaccine are effective.92, 100 These vaccines should not be administered in quarantine because they can result in a false negative tuberculin skin test.100

Separation of Species The separation of nonhuman primates is recommended to prevent disease transmission. In particular, some species such as squirrel monkeys and patas monkeys (Herpesvirus tamarinus and simian hemorrhagic fever virus, respectively) can harbor subclinical or latent disease processes that are fatal in other species of nonhuman primates. From a practical standpoint, multiple species of nonhuman primates are often maintained in the same room in a conditioned colony. To limit potential disease transmission to aberrant hosts, the following general guidelines for the separation of species should be 92, 102 followed: • New and Old World species should not be maintained in the same room. • Asian and African Old World species should not be maintained in the same room. • Squirrel, spider, and cebus monkeys should not be maintained in the same room as owl monkeys, marmosets, and tamarins.

clinical management Basic Veterinary Supplies A variety of supplies and equipment should be available to support the basic veterinary care of nonhuman primates. The sizes of catheters, needles, syringes, and feeding tubes will vary, depending on the size of the animal. Below is a minimal list of nonpharmaceutical supplies and equipment that should be available in a treatment room: • Stethoscope • Rectal thermometer and sleeves • Scale • Pen light or transilluminator © 2002 CRC Press LLC

• Laryngoscope, an assortment of endotracheal tubes, an Ambu bag, and suction bottle and tubing • Nail trimmers and silver nitrate sticks • Restraint box and/or restraint trough and limb restraint ties • Hair clippers with a number 40 blade and clipper blade lubricant • Bandage and suture removal scissors • Water-soluble lubricant • An assortment of syringes varying in size from 1 to 60 ml (luer lock and catheter tip) • An assortment of disposable needles (20–27 gauge and 5/8 –1.5 inches long) • An assortment of indwelling catheters (20–24 gauge and 1–2 inches long) • Blood collection tubes without additives (for the collection of serum) or with sodium EDTA (for the collection of whole blood) • Sterile fluids (lactated Ringers solution and 0.9% saline) and IV administration sets (standard drip set = 15 drops/ml; micro drip set = 60 drops/ml) • Sterile irrigation fluid and skin disinfectants such as povidoneiodine scrub and solution, chlorhexidine and alcohol • Wound pack containing a scalpel handle and blade, fine and heavy scissors, tissue forceps, needle driver, and hemostats • An assortment of absorbable and nonabsorbable suture material with cutting and taper point needles • An assortment of bandaging material, including nonstick pads, cotton gauze, waterproof adhesive tape, cast padding, and elastic wraps • Vaginal speculum • Sterile bacterial culture swabs and fecal sample containers • Feeding tubes (8–16 French) and mouth speculum • High caloric solutions such as Ensure Chicago, IL)

®

(Abbott Laboratories,

• Disposable underpads, and sterile surgical drapes and gloves • Circulating warm water blanket

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Clinical Signs of Illness in Nonhuman Primates The clinical signs associated with illness in nonhuman primates are very similar to those seen in many other species except that nonhuman primates are very adept at masking many of these signs in the presence of humans. For this reason, it is very important that animal care staff be trained to recognize subtle changes in an animal’s behavior and physical condition. Specific signs of illness will vary, depending on the disease condition; however, there are a number of general signs that can indicate a nonhuman primate is sick. General signs of illness include: • Increased or decreased food and water consumption • Increased or decreased urine and fecal output • Loss of bodyweight and body condition • Decreased activity, lethargy, and a disinterest in surroundings • Decreased grooming and social interaction with other nonhuman primates • Ocular and nasal discharge, dyspnea, diarrhea, and emesis • A crouched or huddled posture with hands folded over the abdomen or lying down unresponsive Note: A veterinarian with experience with nonhuman primates should be notified upon identification of a sick animal in order to determineand implement an appropriate diagnostic and treatment plan.

Therapeutic Agents It is not the intent of this section to provide an in-depth listing of all therapeutic agents used to treat nonhuman primates as there are a number of excellent existing sources.95, 103, 104 Some of the more commonly used antibiotic and antifungal drug dosages are listed in Table 4.1. Antiparasitic agents are listed in Table 4.2 and miscellaneous drugs are listed in Table 4.3.

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TABLE 4.1: ANTIBIOTIC PRIMATES

AND

Drug

Dosage

Amikacin Amoxicillin

ANTIFUNGAL DRUG DOSAGES

FOR

NONHUMAN

Route

Indication

Ref.

2.3 mg/kg sid

IM

Antibiotic

103

7–11 mg/kg sid

IM, SC

Antibiotic

103

Amphotericin B

0.25–1.0 mg/kg sid

IV

Antifungal

103

Ampicillin

5 mg/kg bid

IM

Antibiotic

103

Cefazolin

25 mg/kg bid

IM, IV

Antibiotic

103

Cefotaxime

100–200 mg/kg tid

IM

Antibiotic

103

Ceftizoxime

75–100 mg/kg bid

IM

Antibiotic

103

Cephalexin

20 mg/kg bid

PO

Antibiotic

103

Chloramphenicol

20–50 mg/kg bid

IM

Antibiotic

103

Ciprofloxacin

16–20 mg/kg bid

PO

Antibiotic

103

Enrofloxacin

5 mg/kg sid

IM, PO

Antibiotic

103

Erythromycin

40 mg/kg sid

IM

Antibiotic

103

75 mg/kg sid

PO

Fluconazole

2–3 mg/kg sid

PO

Antifungal

103 103

succinate

103

Gentamicin

2 mg/kg bid

IM, IV

Antibiotic

Griseofulvin

20 mg/kg sid

PO

Antifungal

200 mg/kg once

PO

95 95

every 10 days Imipenem

25 mg/kg bid

IV, 30-minute

Antibiotic

105

Kanamycin

7.5 mg/kg bid

IM

Antibiotic

103

Lincomycin Oxytetracycline

5–10 mg/kg bid

IM

Antibiotic

103

10 mg/kg

IM, SC

Antibiotic

103

infusion

Pencillin G, procaine

20,000 U/kg bid

IM

Antibiotic

103

Penicillin G,

40,000 U/kg every

IM

Antibiotic

103

benzathine

3 days

Pipercillin

100–150 mg/kg bid

IM, IV

Antibiotic

103

Tetracycline

20–25 mg/kg bid

PO

Antibiotic

103

Trimethoprim/

0.2 ml/kg of

SC

Antibiotic

103

IV

Antibiotic

105

sulfadiazine Vancomycin

240 mg/ml solution 40 mg/kg/day

continuous drip

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TABLE 4.2: ANTIPARASITIC DRUG DOSAGES

FOR

NONHUMAN PRIMATES

Drug

Dosage

Route

Indication

Ref.

Albendazole

25 mg/kg sid for 5 days

PO

Anthelmintic

103

Fenbendazole

50 mg/kg sid

PO

Anthelmintic

95

for 3–14 days Ivermectin

200 mcg/kg

PO, IM, SC

Anthelmintic

95

Metronidazole

35–50 mg/kg bid

PO

Antiprotozoal

103

0.1 ml/kg once

IM

Cestodes

103

40 mg/kg once

PO

Thiabendazole 100 mg/kg sid

PO

Anthelmintic

103

for 10 days Praziquantel

for 3 days

TABLE 4.3: MISCELLANEOUS DRUG DOSAGES

FOR

NONHUMAN PRIMATES

Drug

Dosage

Route

Indication

Ref.

Aminophylline

25–100 mg/

PO

Bronchodilator

103

103

animal bid Dexamethasone

0.25–1.0 mg/kg

IV, IM, PO

Anti-inflammatory

Furosemide

2.0 mg/kg

IV, IM, PO

Diuretic

Guanfacine

0.5 mg/kg

IM, PO

Self-injurious behavior

106

Kaolin/pectin

0.5–1 ml/kg

PO

Anti-diarrheal

103

IM, IV

Uterine involution,

103

95

every 2–6 hrs Oxytocin

5–20 U total dose

milk let-down Prednisolone

10 mg/kg

IV

Shock

103

0.5–1 mg/kg bid 3-5

PO

Anti-inflammatory

103

PO

Hypovitaminosis C

95

sodium succinate Prednisone

days, then sid for 3-5 days, then every 48 hrs for 10 days, then 1/2 dose every 48 hrs Vitamin C

4–10 mg/kg/day

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common clinical problems The purpose of this chapter section is to review some of the more common clinical problems seen in nonhuman primates. In general, the most common health problems encountered in a nonhuman primate colony involve bacterial enteritis, bacterial pneumonia, and traumatic injuries. The clinical conditions presented in this chapter section are categorized as either viral, bacterial, parasitic, reproductive, or miscellaneous. Where appropriate, a brief discussion of the etiology, clinical signs, diagnostic procedures, and treatment recommendations are included. A preventive health program, as well as good husbandry and sanitation practices, will limit the occurence of many of the clinical conditions described in this chapter section. For more information on these common health conditions, as well as other less-common conditions, it is recommended that one of a num107–109 ber of excellent existing sources be consulted.

Viral Diseases Measles110 • Measles is caused by a human Morbillivirus. • It has been reported in New and Old World species, and the great apes. • The primary route of transmission of measles is through inhaled aerosols from infected animals or humans. • In macaques and most Old World species, the disease is usually mild or asymptomatic unless the animal is stressed or immunosuppressed. Clinical signs most frequently seen include nasal discharge, conjunctivitis, facial edema, blepharitis, a papular skin rash, and in severe cases, pneumonia (see Figure 4.3). • In marmosets and owl monkeys, measles causes a gastroenterocolitis with a mortality rate approaching 100%. These animals present with diarrhea and lack the characteristic rash and upper respiratory signs seen in Old World species. • The diagnosis of measles is based on characteristic clinical, necropsy, and histopathological findings. • There is no specific treatment for measles other than supportive care. • Measles outbreaks can be prevented by immunizing susceptible populations of animals and limiting contact with humans. © 2002 CRC Press LLC

Fig. 4.3 Skin rash in the caudal abdomen/inguinal region of a rhesus macaque infected with measles. (Courtesy of G. Baskin, Tulane Primate Research Center.)

Herpesvirus tamarinus and Herpesvirus simplex110 • Herpesvirus tamarinus (herpes T) and Herpesvirus simplex cause very similar disease conditions in New World species. • Squirrel, spider, and capuchin monkeys serve as asymptomatic reservoir hosts of herpes T, whereas humans are the reservoir host of herpes simplex. • Transmission is through contact with asymptomatic carriers shedding the virus and/or contact with infected bodily secretions. • Herpes T is not typically associated with clinical signs in carrier species, although on occasion it has been associated with oral ulcerative lesions. • In susceptible species (owl monkeys, marmosets, and tamarins), herpes T and herpes simplex can cause a fatal disease process characterized by anorexia, dermatitis, pruritus, depression, and ulcerative lesions of the oral cavity and the gastrointestinal tract. • The diagnosis of herpes T and herpes simplex in susceptible species is based on clinical signs, necropsy, and histopathological findings. • There is no specific treatment for herpes T and herpes simplex in susceptible species other than supportive care. • Herpes T and herpes simplex outbreaks can be minimized by preventing contact of susceptible species with carrier species and humans with active herpes simplex lesions. © 2002 CRC Press LLC

Bacterial Diseases Tuberculosis111 • Tuberculosis is one of the most important bacterial diseases of nonhuman primates. Most reported cases are caused by the acidfast bacillus Mycobacterium tuberculosis or Mycobacterium bovis. • Susceptibility of nonhuman primates varies, with Old World species considered to be more susceptible to infection than New World species. • The primary route of transmission of tuberculosis is through inhaled aerosols from infected animals. Animals with tuberculosis of the gastrointestinal tract can shed the organism in their feces. • In nonhuman primates, tuberculosis classically involves the respiratory system and clinical signs may include coughing, labored breathing, and exercise intolerance. Because tuberculosis can involve a multitude of organ systems, signs vary, depending on the system affected. Some additional nonspecific signs associated with infection include chronic weight loss, diarrhea, and enlarged lymph nodes with or without draining tracts. • The tuberculin skin test is the primary method used in the antemortem diagnosis of tuberculosis. Adjunctive methods/tests that can aid in the diagnosis of tuberculosis include thoracic radiography, ELISA tests, sputum cultures, PCR screening of sputum and feces, and an in vitro blood-based assay of cell-mediated immunity that detects the presence of interferon-gamma. See Chapter 6 for laboratories capable of running such tests. • The postmortem diagnosis of tuberculosis is based on necropsy (the presence of granulomatous-like lesions) and histopathological findings (the presence of acid-fast bacilli), as well as the isolation of the organism from suspected lesions. • Due to the inherent risk a nonhuman primate infected with tuberculosis poses to the colony and staff, it is recommended that animals infected with or suspected of being infected with 94 tuberculosis be euthanatized. For valuable animals or special circumstances, tuberculosis-positive animals can be successfully treated with a regimen of isoniazid, ethambutol, and rifampin.112 • Tuberculosis in a colony can be prevented/minimized by establishing a preventive health program that includes obtaining animals from reputable sources, tuberculin skin testing animals in © 2002 CRC Press LLC

quarantine and the conditioned colony, annual (minimum frequency) tuberculin skin testing of staff that come in contact with nonhuman primates, and limiting nonessential human access to the colony.

Streptococcus pneumoniae111 • Streptococcus pneumoniae is a Gram-positive coccoid bacterium usually found in pairs or chains. • It can be carried in the respiratory tract of asymptomatic nonhuman primates as well as other species, including humans. • The primary route of transmission is through inhaled aerosols from infected animals. • Most reported clinical cases in nonhuman primates involve macaques and the great apes. Active disease is often associated with stress-related factors such as shipment, quarantine, and viral respiratory infections. • Animals infected with this agent typically present with clinical signs consistent with pneumonia, such as a cough, dyspnea, and exercise intolerance. In addition, this agent may cause bacterial meningitis. These animals will present with central nervous signs such as ataxia, head tilt/head press, paralysis, and seizures. • Diagnosis is based on clinical signs, necropsy, histopathological findings, and isolation of the organism. • Clinical pneumonia has been effectively treated using long-acting penicillins; whereas clinical disease associated with meningitis has been effectively treated with cephalosporins. • Other bacterial agents that cause pneumonia in nonhuman primates include Klebsiella pneumoniae and Bordetella bronchiseptica.

Shigellosis111 • Shigella are rod shaped, Gram-negative bacteria. The most common isolate in nonhuman primates is Shigella flexneri. • It is one of the more common enteric bacterial pathogens isolated from nonhuman primates, occurring in New and Old World species and the great apes. • Infection is spread by the fecal-oral route and asymptomatic carriers may exist within colonies.

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• Clinical signs associated with shigellosis include depression, anorexia, dehydration, weakness, and diarrhea containing variable amounts of blood and mucus. Abdominal pain is often evident and affected animals may sit in a hunched posture with hands and arms folded across their abdomen. Marked gingivitis has also been associated with nonenteric shigella infections. • Diagnosis is based on clinical signs and isolation of the organism from deep rectal swabs or fresh stool specimens. Culture and isolation of the organism is facilitated by the use of selective media such as selenite broth and salmonella/shigella agar. • Shigellosis has been successfully treated using enrofloxacin. Supportive care, including fluid and electrolyte administration, may be necessary in severe cases.

Campylobacteriosis111 • Campylobacter are curved, slender, Gram-negative bacteria. The most common isolates of nonhuman primates are Campylobacter jejuni and Campylobacter coli. • It is one of the more common enteric bacterial pathogens isolated from nonhuman primates, occurring in New and Old World species. • Infection is spread by the fecal–oral route and asymptomatic carriers may exist within colonies. • Clinical signs associated with campylobacter infections include watery diarrhea with or without blood, dehydration, and weight loss. In some cases, an animal may present with chronic or intermittent diarrhea. • Diagnosis is based on clinical signs and isolation of the organism from deep rectal swabs or fresh stool specimens. Culture and isolation of the organism is facilitated using selective media such as Skirrow’s or Brucella Agar Plates incubated at 42°C in a 5 to 10% oxygen and 5 to 10% CO2 environment. • Campylobacteriosis has been successfully treated using oral erythromycin. Many infections are self-limiting and animals may only require supportive care (fluid and electrolyte administration).

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Parasitic Diseases Gastrointestinal protozoal agents113 • Nonhuman primates are frequently infected with gastrointestinal protozoal agents. Agents associated with clinical disease include Cryptosporidium spp., Entamoeba histolytica, and Balantidium coli. • These agents are transmitted through the fecal-oral route, direct contact with infected animals and their excretions, and contaminated food and water. • The clinical signs associated with these agents can vary from an asymptomatic carrier state to severe diarrhea and dehydration. Specifically, Cryptosporidium infections have been associated with intractable diarrhea and dehydration in infants and immunosuppressed animals, whereas Balantidium coli has been associated with severe ulcerative enterocolitis and death in great apes. Entamoeba histolytica has been reported to cause more significant disease, including marked diarrhea with blood and mucus, in young and immunosuppressed animals, and New World species. • Diagnosis depends on the microscopic recognition of the organism in feces or on histopathological examination of the gastrointestinal tract. Demonstration of these organisms requires special staining methods (Figures 4.4 and 4.5). • Entamoeba histolytica and Balantidium coli have been successfully treated using metronidazole. Cryptosporidium infections are usually self-limiting and most animals can be effectively treated by providing supportive care, including fluid and electrolyte administration.

Nematodes113 • Oesophagostomum spp., or nodular worms, are the most common nematode parasite found in Old World species and the great apes. Infection is usually asymptomatic; however, heavy burdens can cause diarrhea, weight loss, and abdominal adhesions. The parasite produces firm, smooth, black or white nodules in the wall of the colon. Infection can be diagnosed by identifying eggs in the feces. Oesophagostomum eggs resemble hookworm eggs. Thiabendazole and ivermectin have been used to treat animals infected with this parasite. © 2002 CRC Press LLC

Fig. 4.4 Entamoeba histolytica trophozoite with characteristic nucleus and vacuolated cytoplasm (trichrome stain of fecal smear, magnification × 2000).

Fig. 4.5 Entamoeba histolytica cyst with chromatid body (trichrome stain of fecal smear, magnification × 2000).

Fig. 4.6 Strongyloides spp. egg is thin walled, transparent, and embryonated (wet mount, magnification × 600).

• Strongyloides cebus and Strongyloides fulleborni are commonly found to infect New and Old World species, respectively. Infection is usually asymptomatic; however, severe infections can cause diarrhea. Other clinical signs associated with larval migration include cough, dyspnea, and dermatitis. Infection can be diagnosed by identifying eggs in the feces (Figure 4.6). Thiabendazole and ivermectin have been used to treat animals infected with this parasite. © 2002 CRC Press LLC

Arthropods113 Lung mites, Pneumonyssus spp., are the most important arthropod found in Old World monkeys. The most commonly encountered member of this species is Pneumonyssus simicola, which is found in rhesus monkeys. Animals with lung mites are usually asymptomatic; however, coughing and pulmonary lesions have been associated with infestation. Pulmonary lesions typically consist of small, pale yellow to gray/tan cystic foci throughout the lungs. In addition, fibrous adhesions between the lungs and the pleural cavity may be present. The diagnosis of lung mites is based on necropsy and histopathological findings. Ivermectin has been reported to be effective in eliminating lung mite infestations in nonhuman primates.

Reproductive Conditions Endometriosis114 Endometriosis is one of the most common reproductive disorders seen in rhesus and cynomolgus macaques. In addition to infertility, suggestive clinical signs of endometriosis include cyclical abdominal pain, anorexia, weight loss, depression, and a decrease or absence of feces as well as the presence of a mass in the caudal abdomen. Infrequently, prolonged menstrual bleeding coupled with mild anemia may be observed. Pathologic findings are typically associated with the reproductive tract and include the presence of cystic structures containing a characteristic brown (“chocolate-like”) fluid as well as localized to extensive adhesions involving the bladder and colon. A definitive diagnosis of endometriosis can be established by ultrasound guided aspiration of brownish fluid from cystic structures in the caudal abdomen or by surgical visualization of the reproductive organs. Endometriosis can be managed medically with danazol, leuprolide, or medroxyprogesterone, or surgically via ovariohysterectomy. In severe cases of endometriosis, marked adhesions may preclude surgical management.

Dystocia Dystocia occurs in nonhuman primates and is considered relatively common in squirrel monkeys and marmosets. Predisposing factors include the size, presentation, and position of the fetus as well as maternal pelvic abnormalities. Squirrel monkeys are particularly prone to dystocia because of the infant’s relatively large head size

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and high birth weight, 15 to 17% of the mother’s weight for male infants and 5 to 8% for female infants.32, 35, 114 Typically, nonhuman primates give birth at night or during the early morning while in a squat or upright position. Uncomplicated labor usually lasts 1 to 4 hours, and the vast majority of babies are delivered head first. Signs indicating that a nonhuman primate is having difficulty giving birth and may require manual or surgical assistance include: • A weak, depressed animal lying down in the cage • Labor that fails to progress during a 2-hour period or that lasts for more than 4 or 5 hours • Evidence of fetal malpresentation during delivery (i.e., the presence of a limb(s) or tail in the pelvic canal or exiting the vagina) Note: Dystocia should be considered a life-threatening condition for both the mother and the unborn infant, and a veterinarian should be contacted immediately should an animal appear to be having difficulty giving birth.

Miscellaneous Conditions Traumatic injuries Traumatic injuries are commonplace in nonhuman primate facilities. Most injuries occur as a result of fighting. The most frequently encountered injuries typically involve the hands, feet, or tail. These injuries can be quite extensive and may require amputation of digits and/or portions of the tail. Fresh lacerations/wounds with minimal contamination can be managed by thoroughly cleaning/lavaging the site with a 0.05% chlorhexidine solution prior to suture repair. In addition to thorough cleaning, old contaminated or infected lacerations/wounds may require the use of wet-to-dry bandages, drains, or surgical debridement. An excellent source on wound management can be found in Small Animal Wound Management by Swaim and 115 Henderson. In group housed animals, fighting may result in crushing injuries. Crushing injuries are characterized by areas of marked soft tissue damage (bruising). The significance of these injuries is deceptive and they are often difficult to detect because of the length of an animal’s hair as well as the lack of a major laceration and external hemorrhage. Various factors released from the damaged soft tissue can

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induce acute renal failure; therefore, animals with crush-like injuries should be treated aggressively with intravenous fluids. The management of nonhuman primates with wounds can be challenging because of their intelligence, dexterity, and inclination to pick at wounds and suture lines. Numerous devices/methods have been described in the literature to protect wounds, including Robert-Jones splints, fiberglass casting materials, primate jackets, and Elizabethan collars (Figure 4.7).116 Many animals will leave bandages on if they are applied to the entire length of the affected appendage (Figure 4.8). If bandages are used, they should be checked frequently to make sure they are clean, intact, and have not slipped in such a manner as to occlude the venous return of the respective limb.

Dental conditions In addition to the accumulation of dental tartar and associated gingivitis and periodontal disease, one of the more frequently encountered dental conditions in a nonhuman primate colony is tooth root abscess. Tooth root abscesses can develop subsequent to severe periodontal disease, technical failure of canine disarming methods, and

Fig. 4.7 Primate jacket and Elizabethan collar “hoopskirt” used to protect wounds and bandages on the hind limbs.

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Fig. 4.8 Example of a full-length, soft padded bandage applied to an injured limb. traumatic injuries to teeth from chewing on cages, sticks, and other manipulanda. They typically present as a draining fistulous tract in either the oral cavity or on the face of an animal (Figures 4.9 and 4.10). A root canal procedure or extraction of the affected tooth is the treatment of choice for an abscessed tooth.

Behavioral disorders Abnormal behavior in nonhuman primates often takes the form of stereotypic behavior, that is, a repetitive action that does not appear to serve any apparent biologic purpose. Examples of stereotypic behaviors include pacing, back flipping, bouncing and rocking, as well as digit sucking, self-clasping, and excessive grooming. Nonhuman primates can also exhibit self-injurious behaviors such as hairplucking, head banging, and self-biting. A variety of factors have been proposed as being associated with the development of aberrant behavior, including rearing method, social isolation, and environmental complexity. Animals reared in total or partial isolation not only frequently develop a variety of stereotypes, but also may lack the social skills necessary to be pair or group housed as adults. There are a number of sources that should be consulted for more information on what constitutes normal and aberrant behavior as well as the 55, 117 causes of such behavior. © 2002 CRC Press LLC

Fig. 4.9 Face of a macaque with a draining fistulous tract from an abscessed tooth.

Fig. 4.10 Oral cavity of a macaque with a draining fistulous tract from an abscessed tooth.

Hypothermia Hypothermia is a frequently encountered clinical condition in nonhuman primate facilities. See Chapter 1 for species-specific temperature ranges. Animals most prone to develop hypothermia include

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the smaller New World monkeys, as well as the young, old, debilitated, and anesthetized nonhuman primates of all species. The clinical presentation of hypothermia includes lethargy, huddling, recumbency, and/or an increase in anesthetic recovery time. Body temperatures of nonhuman primates can readily be assessed using mercury rectal, digital, or tympanic membrane thermometers. There are reports that tympanic membrane thermometers in nonhuman primates read approximately 0.5 to 1.0°F less than mercury rectal thermometers; thus, baseline ranges should be established for these devices.32 Hypothermia generally occurs secondary to some other underlying process; therefore, after initiating measures to treat an animal’s hypothermia, the animal should be assessed to determine possible causes such as hypoglycemia, debilitating systemic disease, or anesthesia/chemical restraint. Below is a list of some methods to minimize further heat loss as well as provide supplemental heat. • Direct contact with metal surfaces, such as found in cages and on examination tables, should be eliminated because these surfaces can serve as heat sinks. This can be done by simply placing insulating material between the animal’s body and the metal surface. • Animals should be removed from drafty locations such as positions near supply or exhaust ducts. • Animal room temperatures should be checked and, where appropriate, increased. • Heat lamps are frequently used to provide supplemental heat to nonhuman primates; however, care must be taken to minimize thermal burns. Heat lamps should not be placed any closer than 4 feet from an animal or cage. When using a heat lamp on an animal in a cage, it should be directed only on a portion of the cage so that the animal can self-regulate his/her exposure to the heat source. Halogen lamps should never be used as a supplemental heat source. • Warm water recirculating blankets, nontoxic heat solution packs that use an exothermic reaction, and warm bags of saline can be used to provide supplemental heat to anesthetized or recumbent animals. A towel or other linen-like material should be placed between the animal’s body and the heat source. Warm air blankets can also be used to provide supplemental heat to anesthetized or recumbant animals. An electric heating pad should

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not be used as a supplemental heat source due to the potential for thermal burns and electric shock. • Hypothermic animals can also be maintained in incubators or incubator-type cages (Thermocare®, Inc., Incline Village, NV).

Hypoglycemia Hypoglycemia is another condition frequently encountered in nonhuman primate facilities. See Chapter 1 for species-specific glucose ranges. Animals at risk for developing a hypoglycemic crisis include small New World species, anorectic or fasted animals, animals receiving insulin, and the young of all species, in particular squirrel mon32, 118 The clinical presentation of hypoglycemia includes key infants. lethargy, weakness, ataxia, recumbency, and hypothermia. Hypoglycemia can readily be determined from a drop of blood using a blood glucose monitoring system available at most pharmacies. A hypoglycemic animal that is conscious can be treated by orally ® administering a 20% dextrose solution or Gatorade (Gatorade Corp., Chicago, IL). A hypoglycemic animal that is unconscious can be treated by administering intravenously 2 to 4 ml/kg of a 50% dextrose solution.119 Subsequent administration of a high caloric liquid diet such as Ensure® by stomach tube will minimize potential relapse. In addition, most unconscious/recumbent animals will be hypothermic and will require heat supplementation. Upon recovery, animals should be offered their favorite food items, monitored for a relapse, and assessed to determine the underlying cause of the hypoglycemic episode.

Acute gastric dilatation/bloat

120

Acute gastric dilatation/bloat occurs in both Old World and New World species. The cause of bloat in most cases is associated with the rapid production of gas in the gastrointestinal tract by clostridial organisms. Predisposing factors implicated in the disease include a sudden change in diet, excessive water intake, prolonged broad-spectrum antibiotic therapy, anesthesia, shipping, and fasting followed by free-choice feeding. Bloat is a rapidly progressive disease process that can result in death in a matter of hours. Clinical signs include abdominal distension, restlessness, lying down in the cage, rapid shallow respiratory pattern, increased heart rate, pale/gray mucous membranes, and a prolonged capillary refill time. At necropsy, animals have markedly distended stomachs and distended and congested intestines. In some © 2002 CRC Press LLC

cases, the stomach may rupture. In addition, subcutaneous hemorrhage, edema, and emphysema may be present, as well as rectal and vaginal prolapse. If clinical signs consistent with acute gastric dilatation are noted, a veterinarian should be contacted immediately. Treatment consists of passing a stomach tube to relieve gastric gas and excess fluid buildup. Supportive therapy, including fluid therapy, antibiotics, analgesics, and corticosteroids for shock, should also be initiated. The occurrence of bloat can be decreased by limiting feed intake after fasting and anesthesia, changing diets gradually, feeding animals multiple times during the day, and the judicious use of broad-spectrum antibiotics that affect the gut flora.

Scurvy121 Nonhuman primates develop scurvy if they do not receive adequate amounts of dietary vitamin C. Typical clinical signs include joint swelling and joint pain, lameness, anemia, and gingival and subcutaneous hemorrhage. Cephalohematoma is a common finding in squirrel monkeys with scurvy. Treatment for vitamin C deficiency consists of ascorbic acid injections; 25 mg/kg given intramuscularly twice daily for 5 days. Vitamin C deficiency can be prevented by feeding an appropriately stored commercial nonhuman primate diet within 3 months of milling.120

anesthesia and analgesia The principles of anesthesia utilized in other species can be applied to nonhuman primates. The purpose of this section is to briefly outline the principles of anesthesia, peri-anesthestic management, and information on the use of anesthetics and analgesics commonly used in nonhuman primates. Readers are strongly encouraged to consult with a veterinarian who has experience in primate anesthesia and to utilize veterinary or human anesthesiology texts to augment their understanding of the information provided in this condensed format.122, 123

General Principles Definitions • Anesthesia is described as the loss of sensation to the entire or

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any part of the body. It is induced by drugs that depress the activity of nervous tissue locally, regionally, or within the central nervous system. • Analgesia is defined as the insensibility to pain without loss of consciousness. • Sedation is a calm state usually accompanied by drowsiness. • Tranquilization refers to a state characterized by calmness without drowsiness or unconsciousness, where analgesia is not usually provided. Anesthetics may be used simply for chemical restraint in nonhuman primates to minimize risk to personnel and distress or potential injury to the animals. Many anesthetic regimens are able to provide pain relief in addition to the unconsciousness, which is important for surgeries and other potentially painful procedures. Analgesics are commonly used pre-emptively prior to surgery to reduce post-operative pain and are usually continued after animals recover from anesthesia to extend pain relief. Sedatives and tranquilizers are often used to reduce the amount of general anesthetics needed, as well as to counteract some of their negative physiologic effects. It is the responsibility of those planning to use nonhuman primates to consult with a veterinarian to select the appropriate anesthestics and analgesics to ensure that pain is alleviated. To withold the use of anesthetics and analgesics must be scientifically justified and approved by the facility’s Institutional Animal Care and Use Com66 mittee. Training in the use of anesthestics is very important for persons unfamiliar with their use. People who administer anesthetics should be familiar with the stages of anesthesia because these are used to determine whether an animal requires more or less anesthesia for a given procedure. Moreover, familiarity with the signs of anesthesia allows staff to provide a more consistent application of anesthesia. Veterinary support is often appropriate for anesthesia where surgery or prolonged recumbency is anticipated and, under special circumstances, may require the use of advanced skills and techniques beyond the scope of this chapter.

Stages of general anesthesia • Stage I: termed the stage of voluntary movement, it lasts from the initial administration of the anesthetic until the loss of con© 2002 CRC Press LLC

sciousness. Some analgesia may be present. This stage is the most variable. • Stage II: termed the stage of delirium or involuntary movement, it lasts from the onset of unconsciousness until the onset of regular breathing. Depression of the central nervous system causes the patient to lose voluntary control and reflexes become more primitive and exaggerated. In the nonhuman primate, excessive salivation may occur, and because the gag reflex is pronounced, vomiting may occur during this stage. It is important that this stage be short. • Stage III: termed the stage of surgical anesthesia. Patients are unconscious with progressive depression of reflexes. Muscles become relaxed, and swallowing and gag reflexes and response to painful stimuli are lost. • Stage IV: termed the stage of respiratory arrest. The central nervous system is very depressed. The heart will only keep beating for a short time after this stage begins unless immediate resuscitative steps are taken.

Characteristics of anesthesia • Respiration. Breath holding can occur in Stages I and II, and breathing is irregular in Stage II although it becomes more regular as Stage III is entered. As the depth of anesthesia increases in Stage III, the muscles used to expand the chest and diaphragm weaken. This causes respiration to become shallow and, consequently, the respiratory rate increases. In late Stage III, the abdominal muscles are responsible for respiration. Respiration stops in Stage IV. • Circulation. Heart rate, blood pressure, and capillary refill time (CRT) are the parameters used to assess the circulatory system during anesthesia. CRT is the amount of time it takes a capillary bed in the gums, conjunctiva, or tongue to refill after blanching when digital pressure is applied. Normal CRT is less than 1 second. It is often monitored in the absence of more sophisticated methods such as arterial pressure. Arterial pressure is the most 124 reliable indicator of the adequacy of circulation. During Stages I and II, the pulse is strong and accelerated, and arrhythmias may occur in Stage II. In Stage III, the pulse rate is regular although accelerated slightly, and pain stimulation can cause the © 2002 CRC Press LLC

pulse rate to increase or become irregular. As anesthesia deepens, blood pressure declines and the pulse weakens. • Ocular signs. These signs include eyeball position and movement; pupillary size and light reflex; lacrimation; and palpebral, corneal, and conjunctival reflexes. While ocular signs can be helpful indicators of depth of anesthesia, they tend to be variable. Injectable anesthestics such as ketamine and tiletamine influence these signs. Because these two anesthetics produce muscle contraction, nonhuman primates anesthetized with these drugs have eyelids that do not blink even when the cornea or the conjunctiva is touched, and their pupils are dilated and unresponsive to light. In nonhuman primates, eyeball position, pupillary size and light reflex, and palpebral reflexes are often monitored, especially when patients are anesthetized with gaseous anesthetic agents. In Stage III, the globe of the eye is usually rotated down, pupils are constricted and unresponsive to light, and the palpebral reflex is absent or sluggish. Late Stage III can be characterized by dilated pupils that are unresponsive to light. As anesthesia is reduced, the palpebral and pupillary light reflexes return, and in Stage II the globes turn upward. • Pharyngeal and upper airway reflexes. Coughing (or gagging) and laryngospasm are usually lost in early Stage III, allowing endotracheal intubation by experienced personnel. Attempts to perform endotracheal intubation before this reflex is lost can result in vomiting. The use of neuromuscular blocking drugs to assist with intubation is not routinely necessary. Special care must be taken when attempting to intubate animals that have not been fasted in order to avoid the potential of aspiration of vomitus. • Other signs. Muscle tone progressively declines after Stage II, and its return during recovery from anesthesia is a signal of reduced depth of anesthesia. Loss of jaw tone, muscle tone in the limbs and anal sphincter, and a change in the character of respirations from a thoracic to abdominal pattern are signs that indicate deepening anesthesia. Response to painful stimuli such as the digital reflex (pinching the web between fingers or toes) usually returns in early Stage III anesthesia, but can be influenced by the type of anesthetic being used.

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Signs of pain Lack of response to painful stimuli provides a means to determine the extent of analgesia present in a nonhuman primate. Signs of pain are variable among individuals and species. Conscious nonhuman primates frequently mask signs of pain.125 Observers familiar with the animals may detect subtle behavioral changes that may be the result of pain; however, the frequency of analgesic administration is often determined by the perceived level of discomfort a human might be expected to feel. Signs of pain in conscious nonhuman primates include: • Decreased activity and appetite, accompanied by weight loss • Lack of interest in their environment • Guarding the area that is painful (i.e., abdominal splinting, favoring a limb, not eating hard food) • Inflamed (red and swollen) and/or exudative lesions During anesthesia, it is possible to utilize physiologic indicators as listed in Table 4.4 to assess pain. Personnel administering anesthetics can make adjustments utilizing these indicators in conjunction with their assessment of the signs of anesthesia to provide animals with optimum anesthesia while animals are unconscious.

Peri-Anesthetic Management Anesthetic selection The selection and use of anesthetic agents is best done in consultation with veterinary professionals who can utilize their professional expertise to develop the most appropriate anesthetic protocol for a given situation. A careful pre-anesthesia assesssment of the animal to determine drugs and dosages needed, while considering drug effects that may impact either the animal or the research objective, will help TABLE 4.4: PHYSIOLOGICAL INDICATIONS

OF

PAIN

↓ peripheral circulation ↑ or ↓ respiration ↑ or ↓ body temperature ↑ heart rate ↑ or ↓ blood pressure Deviations in WBC, circulating cortisol, catecholamines Note: ↓, decreased; ↑, increased.

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prevent complications and confounding results. The choice of anesthetic protocol should be one that is safe for the patient based on: • Patient factors likely to influence the uptake, distribution, and elimination of the anesthetic • The physical status of the patient • The specific needs of the species • The specific needs of the case, including the relative requirements for sedation, immobilization, analgesia, relaxation, and safety • The experience of the personnel administering the anesthestic Ideally, the anesthetic protocol should be designed so that the procedure can be accomplished efficiently, thus ensuring patient comfort as a result of consistent application. It may be necessary to perform a pre-study anesthestic trial to provide personnel with adequate training to consistently administer the anesthesia and to ensure that the anesthestic protocol is adequate for the intended use.

Patient care prior to anesthesia Patient evaluation The health status of the animal prior to anesthesia can affect the outcome of an anesthetic procedure. History and physical examination are the best ways to assess for the presence of disease. Laboratory tests are only necessary based on the patient’s history and physical examination results. Aged or debilitated animals and those with experimentally induced disease can require special care for successful anesthetic outcomes, and anesthesia should be done under the supervision of an experienced veterinarian. Anemia, hypoproteinemia, and decreased hemoglobin oxygen saturation concentration are examples of preexisting physiological conditions that may require additional treatment and support prior to and during an anesthetic procedure. Patient preparation To avoid complications resulting from vomiting during anesthesia, food is usually withheld from animals to allow the stomach to empty. Small marmosets are typically fasted 6 to 8 hours, while the larger species are fasted 12 hours. Hypoglycemia may result if small or young animals are fasted for prolonged periods. It is generally not necessary to withhold water and it is contraindicated in patients with © 2002 CRC Press LLC

kidney disease. For surgeries that involve the lower intestinal tract, pre-operative bowel cleansing may need to be done beginning up to 48 hours prior to anesthesia through the use of electrolyte enemas (GoLYTELY®, Braintree Laboratories, Braintree, MA) or cathartics used for human patients. Where the risk of infection is high, prophylactic antibiotics may also be administered prior to surgery. Preoperative analgesics should be administered in cases where post-procedural pain is anticipated. Consultation with a veterinarian prior to the administration of drugs for these purposes is strongly advised because their use may be contraindicated in individual animals or research studies.

Patient care during anesthesia Airway An unobstructed airway must be maintained. Cheek pouches of Old World species should be checked immediately after sedation to remove any food or other objects that might be aspirated, because fasting these animals will not ensure that cheek pouches are empty. In the event salivation or vomition occurs during anesthesia, suction equipment is useful to prevent aspiration. Endotracheal intubation can help prevent breathing impairment during anesthesia; however, endotracheal tubes can become obstructed and personnel monitoring intubated animals must be alert to recognize this complication. Position Normal breathing and circulation should be maintained during anesthesia by placing the animal on its side or back, restraining the limbs with padded ties, and positioning the head to allow saliva to run out of the mouth rather than pool in cheek pouches or accumulate in the back of the mouth where it might be aspirated. Prevention of hypothermia/hypovolemia An anesthetized animal loses the ability to maintain body temperature and can quickly develop hypothermia if supportive measures are not used to maintain body heat. Warmed IV fluids and circulating warm water blankets are commonly used, and circulating warm air units are very useful for animals undergoing prolonged anesthesia. Protecting animals from metal surfaces that conduct heat away from the body should be routinely done. The use of heat lamps may produce thermal burns in anesthetized animals, so they must only be used with extreme caution and only when animals are closely monitored.

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Eye protection Ophthalmic lubricant should be used routinely in anesthetized animals to protect the corneas from dessication. The eyes of patients anesthetized with ketamine or tiletamine/zolazepam remain open, thereby increasing the potential for dessciation or irritation. Urine collection Catheterization of the bladder, especially for animals undergoing abdominal surgeries or prolonged procedures, is routinely performed to prevent urination during surgery. This helps to prevent contamination of the surgical site, provide better visualization of the abdomen, and prevent heat loss through moisture evaporation. Patient monitoring The goal of good anesthesia should be to provide a level of anesthesia sufficient to perform the procedure. Reflexes are involuntary, purposeful and orderly responses to a stimulus. They are lost as brain function is depressed by general anesthesia. Reflexes commonly assessed to monitor depth of anesthesia are found in Table 4.5. Monitoring these reflexes requires access to the head and/or limb of the animal. The extent of monitoring necessary should be determined when the procedure requiring anesthesia is planned. Table 4.6 lists commonly used monitoring parameters and methods. Due to the small size of marmosets and squirrel monkeys, devices developed for rodents may be better suited for these species. Many anesthetic procedures that involve surgery require that much of the animal be covered from view by surgical drapes. This can prevent the use of direct visualization methods to monitor depth of anesthesia such as respiratory rate, reflexes, eye position, and muscle tone. Physiological monitoring methods can assist with patient monitoring in these circumstances, but require specialized monitoring equipment as detailed in Table 4.6. At a minimum, for short surgeries or procedures involving only anesthesia, heart rate, respiratory rate, CRT, and jawtone are adequate to assess anesthesia in nonhuman primates. Prolonged anesthesia or complicated procedures may require more sophisticated monitoring to ensure that patients remain physiologically stable. Equipment designed for use in small animal veterinary practice or for infant humans can often be used on nonhuman primates; however, some electrical monitoring devices may have limits on their ability to detect parameters that fall outside the normal limits of humans.

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TABLE 4.5: REFLEXES COMMONLY USED TO MONITOR EXTENT ANESTHESIA Righting ability

Swallowing/gag/cough reflex

Palpebral reflex

Withdrawal reflex

Pupillary reflex

OF

When lost, the animal is unable to assume normal postures. This is the first reflex to be lost during anesthesia. When lost, the placement of an endotracheal tube is possible. Dissociative anesthetics, such as ketamine, may not eliminate this reflex. When lost, the eyelids will not move when the corner of the eye is lightly touched. Dissociative anesthetics, such as ketamine, interfere with the interpretation of this reflex. When lost, a toe that is firmly squeezed will not produce limb withdrawal. Anesthesia is sufficient to perform painful procedures when this reflex is lost. When lost, pupils do not constrict in response to light. Pupillary dilatation and lack of light response are indicative of dangerously deep anesthesia.

Recordkeeping Procedures that require the use of anesthesia generally involve drugs 126 with usage controlled by the Drug Enforcement Administration. To obtain such drugs, appropriate licensure is required by state and federal authorities, and drugs must be maintained in double-locked, secured cabinets with records of usage scrupulously maintained. Individual anesthesia records should document the procedure being performed, specific drug used, dosage, time of administration, monitoring parameters (usually 10 to 15-minute intervals for uneventful procedures), and other pertinent observations with respect to the animal and procedure as they occur. Carefully maintained anesthetic records can alert staff to trends that may require action to avert a negative anesthetic outcome. Retrospectively, these records document adequate care and use of the animals, and provide investigative personnel with information pertinent to their research. © 2002 CRC Press LLC

TABLE 4.6: ANESTHESIA MONITORING PARAMETERS Parameter

AND

METHODS

Noninvasive Method

Invasive Method

Heart rate

Stethoscope, palpation, esophageal stethoscope, pulse oximeter, ECG

Swan–Ganz catheter

Blood pressure

Capillary refill time, palpation, indirect blood pressure cuff

Catheterization of vessel

Cardiac output

Ultrasonography

Swan–Ganz catheter

Cardiovascular function

Left atrial pressure

Swan–Ganz catheter

Blood flow

Doppler

Waveform

ECG

Transducer

Respiratory function Rate

Esophageal stethoscope, ventilator, direct visualization, capnography

Tidal volume/minute volume

Ventilator, anesthestic machine

End tidal CO2

Capnometer

Inhaled gas concentration

Anesthetic machine

Metabolic status Oxygenation

Capillary refill time, pulse Arterial blood gas, oximeter, tissue appearance hemoximeter

Electrolyte analysis Acid-base Body temperature

Blood collection, arterial blood gas, lactate analysis Rectal thermometer

Biochemical profile

Swan–Ganz catheter Blood collection/analysis

Hematologic status Level of hydration

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Skin turgor, capillary refill time

Hematocrit, central venous catheter

Patient care post-anesthesia Environmental considerations Nonhuman primates recovering from anesthesia should be kept warm. Metal cages will conduct body heat away from the animal, possibly leading to hypothermia. Placing recovering animals in cages in rooms with increased ambient temperatures, the judicious use of supplemental heat devices such as heat lamps, chemical heating pads, and laying animals on disposable underpads to protect them from contact with bare metal are all methods that can be used to maintain normothermia. The administration of warmed IV fluids can also be used during recovery from anesthesia. Recovery areas should be quiet and preferably separate from species such as dogs and swine because nonhuman primates may become frightened if they awaken in the presence of perceived predator species. Position Animals should be maintained in a position that allows normal breathing and minimizes swelling and soilage of surgical sites. Monitoring Vital signs (i.e., heart rate, respiratory rate, rectal temperature) should be checked until animals start to return to consciousness. Once animals exhibit voluntary movement and can swallow, visual checks may be all that is safe and prudent for the caregiver to perform. Animals can occlude their airways while still in Stage I anesthesia; therefore, they should be monitored closely until they can maintain an upright posture. Treatments These are generally determined by the reason for the anesthesia rather than the anesthesia itself. However, hypoglycemia is a common problem in animals that have been fasted for anesthesia. Treat with dextrose (25% dextrose at 2.5 ml/kg IV in a slow bolus or an oral gavage of Ensure® 10 to 20 ml/kg) in the post-anesthetic period. This will hasten recovery of New World species. If nonhuman primates show prolonged return to consciousness following discontinuation of anesthesia, a veterinarian should be consulted to assess the animal and institute appropriate post-anesthetic treatment.

Anesthetic Agents Injectable anesthetics are often used to initiate anesthesia in nonhuman primates because they can be given to animals while they are manually restrained or in their home cages through the use of

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squeeze-back restraint devices. Sedatives and tranquilizers supplement anesthetics and are usually used to augment an anesthetic by countering some of its negative effects. Although anticholinergic drugs, sedatives, and tranquilizers are considered separate pharmacologic agents from anesthetics, they are included in the following list because they are used in conjunction with anesthetics. Table 4.7 provides dosages, routes of administration, and duration of effect for the pre-anesthetics and anesthetics commonly used in nonhuman primates. Several references are provided to direct readers to sources of more in-depth information on pre-anesthetics, anesthetics, and analgesics.127, 129, 131–136 TABLE 4.7: INJECTABLE PRE-ANESTHETIC

AND

ANESTHETIC DRUG DOSAGES

Agent

Dosage, Route (speciesa)

Approx. Duration

Ref.

Atropine

0.05 mg/kg IM

60 min

127

Glycopyrrolate

0.005–0.01 mg/kg IM

120 min

127

Ketamine

5–25 mg/kg IM

Variable

128

Tiletamine/zolazepam 4–6 mg/kg IM (Mm)

45–60 min

127

10 mg/kg IM (Ss)

Light–mod. anes.

127

5 mg/kg IM (Cj)

15 min light anes.

127

Medetomidine

100 mcg/kg + Ketamine 5 mg/kg IM (Pa), (Mf)

127

Propofol

2.5–10 mg/kg IV bolus induction (Mf)

127

5 mg/kg IV bolus induction (Pa)

127

0.4–0.6 mg/kg/min maintenance, continuous IV admin (Pa), (Mf)

105

2.0 mg/kg IV bolus, then 0.2 mg/kg/min continuous IV admin (Mm)

129

Ketamine/xylazine

5–10 mg/kg/1–2 mg/kg IM (Mm, Pa)

30–60 min

127

Ketamine/diazepam

15 mg/kg/1 mg/kg IM

Variable

128

Thiopental

15–20 mg/kg IV bolus to effect

5 min

128

Pentobarbital

5–15 mg/kg IV

30–60 min

128

a

Ss–Saimiri sciureus; Mf–Macaca fascicularis; Mm–Macaca mulatta; Pa–Papio anubis; Cj–Callithrix jacchus jacchus.

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Pre-anesthetic agents Pre-anesthetic agents often used in nonhuman primates include: • Atropine is used to prevent slowing of heart rate and minimize the production of saliva and bronchial secretions. It produces effects for approximately 1 hour in most species. • Glycopyrrolate has actions similiar to atropine but its effects can last at least twice as long as those of atropine. • Tranquilizers include drugs such as diazepam and midazolam, which are controlled substances. These agents provide muscle relaxation without analgesia and they may decrease heart and respiratory rates. Flumazenil reverses the effects of these drugs. • Sedatives include drugs such as xylazine and medetomidine. These agents provide sedation, muscle relaxation, and analgesia. They lower heart rate and blood pressure and produce peripheral vasoconstriction. The use of these sedatives is contraindicated in aged or sick individuals. Reversal agents, such as yohimbine, atipamazole and tolazoline, competitively bind to the same receptors.

Injectable anesthetics Injectable anesthetics used in nonhuman primates include: • Ketamine is a controlled substance that is most commonly used for chemical restraint via intramuscular (IM) injection. It is a dissociative anesthetic and has minimal negative cardiovascular effects. With ketamine, the eyes of the patient remain open and muscle rigidity is marked when it is used alone. Ketamine does not provide sufficient analgesia of the patient for most surgical procedures unless it is used in combination with other pre-anesthetic drugs such as xylazine. Animals often maintain the swallow/gag reflex when anesthetized with ketamine. Recovery may take several hours after ketamine anesthesia. • Tiletamine/zolazepam is a combination of a dissociative anesthetic and tranquilizer. It is a controlled substance. Tiletamine/zolazepam has a longer duration of effect than ketamine in larger nonhuman primates, and it is most often used instead of ketamine for chemical restraint and short procedures on small monkeys (marmosets, tamarins, squirrel monkeys, and capuchin monkeys) and chimps.

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• Barbiturates include drugs such as thiopental and pentobarbital, which are controlled substances. Thiopental is short-acting, must be given intravenously (IV), and is often used to facilitate endotracheal intubation prior to the use of an inhalant gas anesthetic. Pentobarbital is long-acting, produces dose-dependent cardiovascular depression, and the effects can be unpredictable. Prolonged, violent recoveries discourage its routine use for survival anesthetic protocols. • Propofol is a sedative hypnotic drug that is ultra-short-acting and provides minimal analgesic effects; Fentanyl may be used with it to provide intraoperative analgesia. It must be given IV, either as a bolus or continuous infusion. Apnea can occur after induction of anesthesia using propofol. When used for chemical restraint, patients anesthetized with propofol require close monitoring, because return to consciousness can occur within minutes after propofol is stopped. • Local anesthetics include lidocaine and bupivicaine. They provide local analgesic effects with few systemic effects when used to infiltrate tissue. Small quantities of lidocaine can also be used topically to help prevent laryngospasm during attempts at endotracheal intubation. Direct application to the vocal cords of several drops of anesthestic, using a tuberculin syringe with a short catheter attached to it, is a safe administration method. Bupivicaine has a longer duration of effect than lidocaine and is often used as an intra-operative adjunct to general anesthesia for long acting pain control when nerves are cut.

Inhalant anesthetics Inhalant anesthetics used in nonhuman primates include: • Halothane requires a calibrated, precision vaporizor to administer safely. Reaching a desired depth of anesthesia in animals anesthetized with halothane takes longer as compared to isoflurane. Intracranial pressure increases under halothane anesthesia due to increased cerebral blood flow. Halothane can sensitize the heart muscle to the effects of catecholamines, which can result in cardiac arrhythmias. Monitoring heart rhythm when halothane is administered is recommended. • Isoflurane requires a calibrated, precision vaporizor to administer safely. The animal’s depth of anesthesia is quickly adjusted © 2002 CRC Press LLC

because isoflurane is eliminated almost solely through respiration instead of metabolism. Isoflurane causes minimal cardiovascular depressant effects. Cerebral blood flow is increased in animals anesthetized with isoflurane; however, unlike halothane, preanesthetic hyperventilation will prevent this effect. • Nitrous oxide is used only to reduce the amount of other inhalant anesthetic gases in order to reduce their negative effects. It produces minimal respiratory depression and has a slight cardiovascular sparing effect when used in conjunction with more potent inhalant anesthestics. Nitrous oxide produces less anesthesia in nonhuman primates than in humans; consequently, it is never used alone. Nitrous oxide administration must be stopped prior to discontinuing oxygen administration to allow re-equilibration between nitrogen gas and nitrous oxide present in the blood and gas compartments of the body.

Analgesic Agents Table 4.8 provides information on dosage, route of administration, and duration of effect of the analgesics commonly used in nonhuman TABLE 4.8: COMMON ANALGESIC DRUGS Drug

Dosage/Route

NONHUMAN PRIMATES Ref.

Duration (hr)

Species

6–8

Mm, Pa, Cj 128

0.015 mg/kg

6–8

Ss

Morphine

1–2 mg/kg SC, IM

4

Mm, Pa, Ss 137

Oxymorphone

0.15 mg/kg IM, SC, IV

4–6

Mm, Pa

138

0.075 mg/kg IM, SC, IV

4–6

Ss, Cj

138

5-10 mcg/kg or 10–25 mcg/kg/hr IV

Intra-operatively

Mm, Pa

127

Intra-operatively

Mm, Pa

127

20 mg/kg PO

6–8

Pa

128

125 mg/5 kg rectal suppository

<24

Mm, Pa

103

Mm, Pa

103

Buprenorphine 0.01 mg/kg IM, IV

Fentanyl

infusion Aspirin

Ketorolac a

FOR

15–30 mg/kg IM

a

Ss–Saimiri sciureus; Mm–Macaca mulatta; Pa–Papio anubis; Cj–Callithrix jacchus jacchus. © 2002 CRC Press LLC

103

primates. Opioid analgesics such as morphine, buprenorphine, oxymorphone, and fentanyl are often used to supplement anesthetics that do not provide adequate analgesic effects needed to perform surgery. Oftentimes, to control pain post-operatively, it is necessary to administer an opioid such as buprenorphine as a pre-anesthetic agent so that pain is not perceived after it occurs. It is possible to reverse the effects of opioid drugs through the use of opioid antagonist drugs such as naloxone. All opioid drugs have a potential for abuse and are thus controlled substances. Nonsteroidal antiinflammatory drugs (NSAID) such as aspirin and ketorolac are used to minimize pain caused by inflammation. They do not block pain receptors as the opioid drugs do and, hence, they are not used to supplement anesthesia in the same manner as opioid drugs. In nonhuman primates, NSAIDs are less commonly used because many are oral medications and are not uniformly accepted by all animals. Adequate dosages of NSAIDs may not be ingested by oral administration following surgery. • Morphine is an opioid drug that can be used to provide analgesia during operative procedures. Its side effects include vomition, constipation, and respiratory depression. It has no significant effect on cardiac output and can be reversed by naloxone. • Oxymorphone is an opiod drug that is approximately 10 times more potent than morphine as an analgesic agent. It produces minimal respiratory depression and is usually used to augment injectable anesthetics. Barbiturate dosages can be reduced by two thirds when oxymorphone is used as a pre-anesthetic agent. Sensitivity to loud noises can be appreciable when this drug is used. Naloxone will reverse oxymorphone. • Fentanyl is an opioid drug that is approximately 250 times more potent than morphine as an analgesic agent. It has a very short duration of effect, approximately 30 minutes, when administered IV or IM and it is most often given to nonhuman primates as IV boluses or continuous IV infusions. Like oxymorphone, sensitivity to loud noises is marked in unanesthetized animals. Respiratory depression, apnea, or panting can occur; thus, it is best used when animals are intubated and ventilatory support can be provided. Atropine or glycopyrrolate should be used to offset the bradycardia fentanyl produces. Naloxone will reverse fentanyl. • Buprenorphine is approximately 30 times more potent than morphine as an analgesic agent. It is widely used for post-operative © 2002 CRC Press LLC

analgesia in nonhuman primates because it has a longer duration of effect than other opiod drugs (up to 12 hours). Because it has a slow onset of action, administration as a pre-anesthetic can ensure that pain is controlled in the early post-operative period. It can cause respiratory depression. Naloxone will reverse buprenorphine’s effects. • Aspirin is an NSAID used to control mild to moderate pain produced from inflammation. Oral and rectal suppository formulations are available. Drug absorption is affected by the size of the gastrointestinal tract and stomach emptying when oral formulations are used. Consequently, wide variations in plasma concentration can occur. Side effects include gastric and intestinal ulceration. Renal disease can occur if aspirin is used in patients with hypovolemia, congestive heart failure, or other cardiac impairment. Flavored children’s formulations may be better accepted by nonhuman primates and are available in lower concentrations that are safer to administer to small animals. • Ketorolac is an NSAID that is available as an injectable formulation. Its side effects are similiar to those of aspirin. It can be used to control moderate to severe pain caused by inflammation.

Principles of Inhalation Anesthesia Inhalant anesthetics are often used on nonhuman primates to perform operative procedures. These anesthetics are volatile liquids and can be administered with an anesthetic machine. For safety, these agents should only be administered using equipment that has been designed precisely for the specific agent, and by personnel with the knowledge to do so. Anesthetic machines and associated vaporizors require regular maintenance to ensure that these agents can be safely administered. It is necessary to have compressed oxygen available to deliver an inhalant anesthetic to the patient, and waste anesthetic gases must be collected or directly vented into nonrecirculating air exhaust systems. The equipment used for administration of these agents to humans and animals can be adapted for safe use on nonhuman primates. It is beyond the scope of this manual to describe in detail the use of these systems; consultation with a veterinarian who has experience anesthetizing nonhuman primates using gaseous anesthetics is recommended for people desiring to purchase and use these systems in Old or New World species. A reference is provided for the reader needing more information.139

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Endotracheal Intubation Endotracheal intubation is routinely performed to ensure a patent airway in anesthetized animals, and to administer a gas anesthetic while preventing environmental contamination of the surrounding room with the agent. Intubation is also necesssary to assist with ventilation in the event an animal should stop breathing. The technique used on nonhuman primates is similiar to the techniques 140 described elsewhere for small animals. Supplies used for endotracheal intubation include: • Endotracheal tube; cuffed models are preferred except for very small nonhuman primates and infants; tube sizes range from 2 mm inner diameter to 8 mm for most species of nonhuman pri141 mate • Laryngoscope with a blade long enough to reach into the pharynx of the animal; straight or curved blades may be used • Cotton gauze or umbilical tape to secure the tube in position • Sterile lubricant • Cotton gauze sponges • Syringe to fill the cuff • Stylet to fit into the tube • Ambu bag • Topical anesthetic, tuberculin syringe, and catheter for its administration • Face mask and inhalation anesthestic machine with oxygen • Emergency drugs (see Table 4.9)

Procedure Endotracheal intubation of nonhuman primates is similiar to that of other mammalian species. The position the animal is placed in before attempting the procedure is often based on personal preference. The following steps describe the procedure that many people use to intubate these species: 1. Prior to attempting intubation, personnel should dress in appropriate personal protective clothing, including protective eyewear. 2. Check the endotracheal tube for leaks by inflating the cuff with air and letting it sit for 5 minutes. Withdraw the air from the cuff prior to attempting intubation. © 2002 CRC Press LLC

TABLE 4.9: EMERGENCY DRUGS a

Drug

Indication for Use

Dosage, Route

Species

Atropine

Bradycardia

0.05 mg/kg IV

Any

142

Dopamine

Bradycardia after arrest

10 mcg/kg/min IV

>3 kg

142

Doxapram

Respiratory arrest

2 mg/kg IV

Any

103

Epinephrine

Cardiac arrest

0.2–0.4 mg/kg diluted in 5 ml sterile water; give intratracheally >3 kg or 1:10,000 dilution; 0.5–1.0 ml IV

Ephedrine

Hypotension

1.25–2.5 mg/kg IV

Any

127

Furosemide

Pulmonary edema

1–2 mg/kg IV

Any

127

Lidocaine

Premature ventricular contractions

1–2 mg/kg IV

Any

122

Naloxone

Opioid reversal

0.1–0.2 mcg IV; repeat as needed

Any

127

Norepinephrine Hypotension

0.05–0.1 mcg/kg/min, IV infusion Any

127

Phenylephrine

Hypotension

1–2 mcg/kg IV bolus, then 0.5–1.0 Any mcg/kg/min, IV infusion

127

Atipamazole

Medetomidine reversal

0.2 mg IV; 0.3 mg/kg IV

127

a

Ref.

142

Ss, Pt

Ss–Saimiri sciureus; Pt–Pan troglodytes.

3. Lubricate the end of the tube and cuff with sterile lubricant. 4. The nonhuman primate should be anesthetized to a level of early Stage III anesthesia. 5. Position the animal in dorsal or lateral recumbency (righthanded persons usually prefer the animal’s right side down). 6. Open the animal’s mouth and grasp the tongue with a cotton gauze sponge and apply outward traction on the tongue to maintain the mouth in an open position to allow the introduction of the laryngoscope blade into the mouth. 7. The end of the laryngoscope blade is placed at the base of the animal’s epiglottis (Figure 4.11). Sometimes it is necessary to slightly hyperextend the animal’s neck by exerting pressure on

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Fig. 4.11 Placement of an endotracheal tube in a baboon in dorsal recumbency. Outtake is a cross-sectional view of the glottis and endotracheal tube with soft palate pushed downward.

the laryngoscope blade to straighten out the curve between the animal’s jaw line and neck in order to see the glottis. The animal’s soft palate may prevent good visualization of the animal’s glottis until it can be gently pushed out of the way with the endotracheal tube. 8. Once the laryngoscope is in position, it is steadied with the hand not used to handle the endotracheal tube. 9. The endotracheal tube is passed along the blade of the laryngoscope and through the arytenoid cartilages into the trachea with the beveled end of the tube parallel to the glottis. 10. Time the passage of the endotracheal tube with the animal’s inspiration to facilitate placement. A stylet may be placed inside the endotracheal tube to assist with intubation. If a stylet is used, it should be removed immediately following placement of the endotracheal tube to allow normal breathing.

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11. It is normal for the animal to exhibit a strong gag reflex when the tube has been correctly positioned in the trachea (i.e., below the larynx and above the branching point of the major bronchi). Care must be taken to insert the tube so that it will not be pushed out by the force of the animal’s gag reflex. 12. Position the end of the endotracheal tube in the trachea below the larynx. Premarking the tube prior to placement may help identify when the tube is in the correct position and prevent complications from slippage of the tube during transport or positioning of the animal. 13. Fill the cuff on the tube with enough air to prevent escape of expired air around the tube when the animal is ventilated gently with the Ambu bag. 14. Auscultate each side of the animal’s chest for lung sounds; if lung sounds can only be heard on one side of the animal’s chest, withdraw the tube slightly and reauscultate. (Note: If lung sounds can only be heard on one side of the chest, the endotracheal tube has been placed into a mainstem bronchi and only one side of the lungs is being ventilated.) 15. Secure the tube in place with umbilical tape or cotton gauze tied around the tube first, and then draw the ends of the ties to the back of the animal’s head and secure them. 16. When recovering an intubated animal from anesthesia, the tube is removed when the animal’s gag reflex returns. The ties holding the tube in place are cut and the cuff is deflated using an empty syringe prior to withdrawal of the tube.

aseptic surgery Aseptic techniques must be used when performing survival surgery. This requires the use of procedures to prevent the introduction of pathogenic organisms. To perform aseptic surgery, appropriate facilities, sterile instruments, trained and properly garbed personnel, pre-operative preparation of the patient, and adherence to the principles of asepsis throughout the surgery are required. Following any survival surgery, it is necessary to provide post-operative care to ensure that healing proceeds normally and that any post-operative complications are addressed promptly. Recordkeeping to detail anesthesia, surgery, and post-operative care are required and become © 2002 CRC Press LLC

part of the animal’s individual health record. References provided are not intended to be comprehensive, but rather to introduce readers to several sources for additional information.143–145

Facilities/Features/Equipment The Guide suggests that for most surgical programs, the functional components of an aseptic surgery area should include areas for surgical support, animal preparation, surgeon’s scrub, operating room, and postoperative recovery.54 Separation of functional areas is best achieved by physical barriers, although it might be possible to achieve separation by timing activities and appropriate cleaning/disinfection between these activities. Below is a list of facilities, facility features, and equipment necessary to support aseptic surgery involving nonhuman primates: • Single-use operating room with limited access • Ventilation to surgery rooms should be positive to surrounding areas • Surgical lights that can be focused on the patient from two directions • Numerous electrical outlets • Minimal nonessential equipment and storage in the operating area to allow for easy clean-up and thorough sanitization • Separate surgical pack preparation/sterilization area with steam sterilizer • Surgeon’s prep area with foot, knee, or electric eye surgical sinks (usually separate from the storage and pack prep area) • Patient prep area (containing oxygen source, sink, clippers, vacuum cleaner, suction set-up, anesthesia supplies, and drug storage cabinet) • Dressing room area or multipurpose room that can be used for personnel to change clothes • Adjustable-height surgical tables, kick buckets, vacuum lines for suction • Oxygen source, either in-line and serviced from a central location, or from individual compressed gas tanks • IV stands, IV fluid warming units, circulating warm water or air blankets

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• Inhalant anesthestic machine with exhaust gas scavenging device • Anesthesia monitoring equipment such as pulse oximeter, capnometer, arterial blood pressure, rectal temperature, and ECG • Post-operative recovery room that allows close patient monitoring without physically disturbing them, with the capability to increase ambient room temperature or provide safe supplemental heat sources for animals in recovery cages

Personnel Surgeon and assistant (if necessary). Proper surgical attire includes scrub clothes, eye protection, disposable cap, and mask. Watches, jewelry, and nail polish should be removed beforehand. A surgical scrub using a povidone-iodine scrub solution or chlorhexidine scrub solution should be performed. These solutions are usually provided in sterile packages containing a plastic scrub brush and nail cleaner. First, the area under each nail is inspected and cleaned with the nail cleaner. The scrub is performed by starting at the fingertips under the nails and working toward the hand and forearm to include the elbow. Each surface is scrubbed a minimum of 10 times, rinsed and repeated for a total of three times on each hand and arm, and for a minimum of 10 minutes. The hands are then dried on a sterile towel, which is usually provided with the gown. Gowns are often packed inside-out to allow the surgeon to pick-up and put on the gown without touching the outside surfaces that are to remain sterile. Sterile surgical gloves must be put on in a manner to prevent contamination of the surgical gown, and several techniques may be used according to the surgeon’s preference and availability of personnel to assist 144 them. Once the gowns and gloves are on, the hands should be kept above the waist and in sight at all times. If powdered gloves are used, the powder should be wiped off with a sterile cotton gauze sponge and sterile saline prior to starting surgery. Surgeons and their assistants must be vigilant about not handling or coming into contact with any nonsterile item or surface throughout the operation. Person administering anesthesia. Attire includes scrub clothes, eye protection, cap, mask, and examination gloves. If this is the same person who preps the animal pre-operatively, he/she should wear a lab coat or disposable isolation gown over his/her scrub clothes while clipping and handling the animal outside the surgery room.

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This outer wear is discarded before the animal is prepped in the surgery room. Circulating personnel. Attire is the same as for the person administering anesthesia. Assistants who must provide equipment and supplies to surgeons must utilize techniques to ensure that the operating field and instrument table also remain sterile throughout the operation.

Pre-Operative Preparation Nonhuman primates undergoing surgery require a pre-operative assessment that includes evaluation for the anesthetic, analgesic, and any prophylactic medication requirements. A veterinarian who has experience with nonhuman primates should be consulted by the investigative staff planning experimental surgeries to ensure that animals are appropriately prepared. Personnel supporting the surgery need to be aware of the type of preparation required on the animal, the position required during surgery, the types of anesthestic, analgesics, and other drugs that are to be administered, and surgical instrumentation to be used. If special equipment is needed either for the surgery or supporting the animal during the procedure, staff need to know how to use it and it should be available for use.

Patient pre-operative preparation Patient pre-operative preparation includes: • Pre-anesthesia preparation (see previous section) • Endotracheal intubation (see previous section) • The surgical site must be clipped sufficiently to ensure that contamination will not occur intraoperatively. One should avoid unnecessarily removing too much hair on small animals because hypothermia can result. • Pre-operative antibiotics and analgesics should be given after consultation with a veterinarian experienced with nonhuman primates. • IV catheterization should be performed to permit the administration of parenteral fluids and facilitate emergency treatment. Usually, the cephalic or saphenous veins are used. • The animal is positioned appropriately on the surgery table, and anesthesia monitoring devices such as ECG, thermometer, pulse oximeter, etc., are put in place prior to initiation of the procedure. © 2002 CRC Press LLC

• When electrosurgical devices are to be used, it is necessary to make sure that there is good contact between the animal’s skin and the grounding devices. Electrosurgical gels for this specific purpose are available. One should not substitute ultrasound or ECG gels. Severe thermal burns can result from improper grounding of electrosurgical devices.146

Surgical instruments and drapes Surgical instruments and drapes must be sterilized pre-operatively. Various methods of sterilization are available, and whichever method is used, one should implement appropriate quality assurance methods to ensure that sterilization is consistently achieved. Methods of sterilization include: • Steam sterilization (250°F, with the time dependent on the type of pack being sterilized) is a very dependable method. Sterilization by this method depends on the ability of the steam to reach the items to be sterilized. Pressure steam sterilizers and vacuum steam sterilizers are commonly used for sterilizing surgical instruments. Each of these units is a type of autoclave, because the door of the sterilization chamber is held closed by the pressure within the chamber. They differ in the method that is used to evacuate the air from the sterilization chamber. Pressure steam sterilizers often utilize gravity to displace air from the chamber, while vacuum is used in vacuum steam sterilizers. The disadvantage of steam sterilization is that fine instruments, devices that cannot be disassembled for adequate penetration by the steam, and many heat-sensitive synthetic materials cannot be sterilized using steam. • Ethylene dioxide gas sterilization is safe for nearly all types of materials; however, it is flammable and explosive except when mixed with carbon dioxide. Consequently, it is expensive to maintain sterilizers that utilize this method of sterilization and a high volume of sterilization must be done to offset their cost. Research facilities often arrange to have their sterilization done through formal contractual agreements. This method of sterilization is especially useful for sterilization of items that cannot tolerate steam sterilization. Pre-planning is necessary when utilizing ethylene dioxide sterilization because sterilization cycles require many hours and equipment may not be available for quick turnaround

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or surgeries scheduled on short notice. For this reason, ethylene dioxide should not be relied upon as a sole method of sterilization. ®

• Liquid sterilants such as 2% glutaraldehyde (Cidexplus , Advanced Sterilization Products, Irvine, CA) can be used for specific equipment, but should not be routinely relied upon as the sole method of sterilization for most surgical instruments. Instruments treated with liquid sterilants must be thoroughly rinsed with sterile irrigation saline prior to use. • Gamma-irradiation is another sterilization method used by some facilities. It is usually used for pharmaceuticals and hospital and surgical materials that are heat sensitive.

Operating Room Procedures Once the animal is moved into the surgery room, it is necessary to prepare the surgical site for surgery. This involves the use of the aseptic techniques described below. Surgeons and all operating room personnel should be trained and should adhere to aseptic technique rigorously throughout the surgery.

Aseptic technique To prevent the introduction of pathogenic organisms during a surgical procedure, the surgical site on the patient must be prepared by cleansing and disinfecting the skin. Aseptic technique involves disinfection of the area to reduce any pathogens present. This includes the application of an antiseptic to inhibit or prevent the growth of bacteria during surgery after the site is disinfected, and the utilization of techniques to prevent contamination by microorganisms of the site. The attendant doing the preparation should wear scrub clothes, cap, mask, protective eyewear, and gloves. Surgical preparation of the patient involves scrubbing in a circular pattern, starting at the center of the intended incision site and gradually working outward, never going back over the previously scrubbed area (Figure 4.12). A povidone-iodine based scrub solution diluted with sterile irrigation saline and applied with sterile cotton gauze sponges should be used. New sponges should be used for each scrub cycle. After each scrub, 70% ethyl alcohol is applied in the same circular, concentric manner as used for the scrub. The area is cleansed a minimum of three times and then povidone-iodine antiseptic solution is applied to the surgical site and allowed to dry prior to surgery. To reduce patient © 2002 CRC Press LLC

Fig. 4.12 Scrubbing method, starting at the incision site and moving outward. hypothermia, the amount of saline and alcohol that are used during the scrub should be kept to a minimum. Sterile surgical drapes are used to prevent contamination of the surgical site by contact with nonsterile areas, persons, and instruments. These may be disposable drapes with an adhesive strip on them or reuseable towels. If towels are used, they should be clamped to the skin with sterile towel clamps. A fenestrated sterile drape is used (over the disposable adhesive drapes or towels) to isolate the surgical site from nonsterile areas.

Intra-operative monitoring Anesthesia monitoring during surgery is usually performed by technical support personnel with special training in veterinary anesthesia. In addition, a veterinarian with experience in nonhuman primate anesthesia should be available to assist the support personnel when surgeries on nonhuman primates are scheduled. Monitoring parameters and methods have been previously outlined in Table 4.6. Additional information can be obtained from the references.142, 147

Post-Operative Care Post-operative care begins when the surgery ends and concludes when the surgical incision has healed. Patient monitoring during the

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immediate recovery period (from the end of surgery until the patient regains consciousness) is similiar to that used intra-operatively. Fewer parameters can be monitored once the animal is extubated and placed into a recovery cage. Parameters to be evaluated during the immediate recovery period include: • Temperature, pulse rate, and respiratory rate • Mucous membrane color and capillary refill time • Return of reflexes, including withdrawal (pedal), palpebral, shivering, jawtone, and gag Several other factors to address during the immediate recovery period include: • Position the animal to maintain an open airway and minimize swelling from fluid accumulation by gravity around incision • Provide a warm, quiet, preferably darkened recovery area • Provide preferred foods such as fruit, and position watering device for easy access • Monitor for signs of pain and consult with a veterinarian if animals appear painful Additional information regarding post-surgical considerations is 148–149 provided for readers in the references.

Post-operative records Parameters monitored should be recorded until the animal is conscious. Once the animal is able to move around normally, it can be returned to the home cage, although group housed nonhuman primates should remain singly housed until the veterinarian releases them to return to their group. Inspection of animals should occur daily and the veterinarian should examine any animals that appear sick, painful, do not eat, or have unusual redness, swelling, or discharge from the surgical incision. Post-operative records should indicate the date, an assessment of the animal’s behavior, appetite, fecal output, hydration, and the condition of incision site, and all treatments. In cases in which the investigative staff may be administering treatments, it is important that the veterinarian communicate regularly with these staff and that records be assessible for examination by all. Post-operative records are complete when the incisions are

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healed, sutures have been removed, and animals are ready to return to group housing, if appropriate.

euthanasia Euthanasia is the act of inducing a humane death in an animal. The American Veterinary Medical Association (AVMA) has published recommended methods of euthanasia for animals, including nonhuman primates.150 Nonhuman primates are usually sedated with an anesthetic such as ketamine and subsequently euthanatized via intravenous administration of a saturated sodium pentobarbital solution. For some research applications, to preserve tissues for histological examination or other in vitro techniques, it may be necessary to anesthetize the animal with an injectible anesthetic such as sodium pentobarbital to then perfuse the animal’s tissues. In either case, both methods are approved by the AVMA.

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5 experimental methodology introduction Numerous experimental techniques and methods have been applied to the use of nonhuman primates in biomedical research. It is impossible to cover all the techniques, methods, and variations reported in the literature. Therefore, this chapter presents those techniques and methods most frequently encountered in a research environment. Personnel handling and/or performing experimental manipulations on nonhuman primates must be properly trained. Training should include a review of the institution’s occupational health and safety program; a review of zoonotic concerns; a review of appropriate protective equipment; a review of proper methods for handling biological samples; a review of the behavioral aspects of the species utilized; and a demonstration of the technique(s) to be employed.

restraint The restraint of nonhuman primates for manipulation can be divided into three general categories: (1) physical restraint methods, (2) chemical restraint methods, and (3) operant conditioning. Whatever general method of restraint is utilized, it is of the utmost importance that the nonhuman primate be handled and restrained in a manner that ensures the well-being and health of the animal as well as the safety of the personnel involved.151 This can be achieved, in part, by acclimating the animal to the restraint method, training

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animal carestaff in the proper use of the restraint method, and wearing appropriate personal protective equipment. Personnel handling nonhuman primates should wear, at a minimum, a mask, protective eye wear, disposable procedure gloves, and a long-sleeved gown/coveralls or laboratory coat. The authors recommend double-gloving when handling or manipulating macaques. If handling will involve the restraint or manipulation of an awake nonhuman primate, then the protective equipment should also include leather gloves. The length and thickness of the gloves can vary, depending on the size and species of animal being manipulated. Leather gloves decrease the potential for injury; however, they are not impervious to puncture from sharp canine teeth. Additional protective measures include wearing stainless-steel mesh or Kevlar® (E.I. du Pont de Nemours and Company, Inc., Wilmington, DE) gloves inside the leather gloves (Figure 5.1). These protective devices reduce the chance of deep puncture wounds and lacerations; however, care must still be taken when handling nonhuman primates as these devices will not fully protect fingers from crushing injuries. In addition, rubber, Tyvek®, and Kevlar® (E.I. du Pont de Nemours and Company, Inc., Wilmington, DE) protective sleeves are available. These sleeves, if worn on the arm above the leather gloves, can significantly decrease the chance of animal care personnel being scratched during manipulation of an awake animal (Figure 5.2).

Physical Restraint Methods Squeeze-back cages Cages with squeeze-back mechanisms are the most commonly used restraint device for nonhuman primates. Squeeze-back cages provide a safe, rapid, and relatively stress-free method of restraint for such routine procedures as cursory examinations and injections. These cages are used in the restraint of a variety of species, ranging from marmosets and tamarins to chimpanzees.151 Squeeze-back cages come with either a manual or mechanical restraint system. In general, manual squeeze-back cages consist of either a movable panel or basket (squeeze-back) attached to either one or two pull-bars, respectively. In the non-engaged position, the squeeze-back is located at the back of the cage. Lock(s) on the pullbar(s) ensure that the animal cannot move the squeeze-back forward when not in use. To engage a manual squeeze-back cage, the technician releases the lock(s) on the pull-bar(s) and pulls the squeezeback and the animal to the front of the cage. The pull-bar(s) have © 2002 CRC Press LLC

Fig. 5.1 Leather gloves for handling nonhuman primates (left and right), and stainless-steel (left center) and Kevlar® (right center) mesh glove inserts for additional protection.

Fig. 5.2 Tyvek® (top) and Kevlar® (bottom) protective sleeves.

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notches located so that the squeeze-back, as it is pulled forward, will lock in place, thereby holding the animal against the front of the cage. Manual squeeze-back cages are ideal for restraining animals up to the size of an adult male rhesus macaque. Mechanical squeeze-back cages consist of a movable panel attached to either a gear-box or screw mechanism. To engage a mechanical squeeze-back cage, the technician must use a hand crank or electric drill to move the panel and animal to the front of the cage. In either case, the movable panel locks in place as soon as the technician stops operating the hand crank or drill. Mechanical squeeze-back cages are ideal for large nonhuman primates such as baboons and chimpanzees. When using a squeeze-back cage system for restraint, the following points should be remembered: • The ideal position in which a nonhuman primate should be restrained is with his/her side facing the front of the cage (Figure 5.3). In such a position, the force of the squeeze-back is applied

Fig. 5.3 Mechanical squeeze-back cage with clear side panel demonstrating the proper position in which a nonhuman primate should be moved to the front of the cage (side of animal facing front of cage). © 2002 CRC Press LLC

to the shoulders. This position allows for relatively comfortable yet tight restraint of the animal. Moreover, this position provides the technician with ready access to the major muscle groups for injection. • Not all animals readily assume the ideal position. This may mean that the technician will have to release and re-tighten the squeeze-back to get the animal in a position with its side facing the front of the cage. • Care should be taken when using squeeze-back cages not to entrap the tail, fingers, or toes of an animal at the junction of the squeeze-back and floor. • The type and placement of enrichment devices within squeezeback cages should be taken into account because they can jam the squeeze-back mechanism. Toys hung from the top of squeeze-back cages should be placed near the front of the cage to minimize potential obstruction of the squeeze-back mechanism.

Transfer chutes and cages Transfer chutes and cages are devices that allow for the simple transfer of animals of various sizes from one cage to another without sedation or physical contact. Moreover, chute systems are often incorporated into gang cage or corral systems to facilitate the segregation, removal, and manipulation of group housed animals.152, 153 These devices can be purchased commercially, although many facilities use devices that are customized to their respective caging system.154–156 A transfer chute is a tunnel that connects two cages. The chute can be removable or permanent. Transfer cages, in their simplest form, consist of a cage or box that attaches to the door of a cage or, in the case of marmosets and tamarins, a nesting box with a door that can be closed. Transfer cages can be used to transfer animals between cages. They also provide a convenient method to weigh conscious animals. In some cases, wheels and squeeze mechanisms are added to the transfer cage to facilitate the movement and restraint of larger animals (Figure 5.4). Whether using a transfer cage or chute, there should be a mechanism to securely attach the device to the cage to prevent escape. Most animals can be readily trained to use tunnels or transfer cages by using food treats as positive reinforcement and taking into account the animal’s line of sight. To facilitate transfer during training, it is advisable for the technician to stand off to the side of the © 2002 CRC Press LLC

Fig. 5.4 A mobile transfer cage designed to transfer animals between stacked cages. cage opening and avoid direct eye contact with the animal. Additionally, if the caging system allows, the placement of a technician behind the cage will often result in the movement of the animal to the front of the cage and into the chute or transfer cage. When using transfer chutes or cages, the following points should be remembered: • The door to the animal room should always be closed when moving an animal into a chute or transfer cage. • The transfer chute or cage should always be securely attached to the cage. • When moving an animal into a transfer cage, the door of the transfer cage should be opened prior to opening the door to the animal’s cage because a well-conditioned animal will sometimes dart out of his/her cage without checking to see if the transfer cage door is open. • The reverse is true when moving an animal from a transfer cage back into the animal’s cage. In this case, the door to the animal’s cage should be opened prior to opening the door of the transfer cage. © 2002 CRC Press LLC

• During the transfer process, care must be taken to avoid dropping cage doors onto the animal’s tail, hands, and feet.

Nets Nets are primarily used to capture escaped nonhuman primates or group housed animals. In addition, because nets allow the technician to maintain a safe distance from the animal, they can be safely and effectively used to direct or shepherd the movement of a group housed animal or escapee toward a chute or open cage. Although nets have been used effectively to capture smaller nonhuman primates housed in groups, it is less stressful to the animal to use chutes or transfer cages/boxes to segregate group housed animals. Lightweight, smallgauge mesh nets (scissor nets) are best used to capture smaller nonhuman primates. The small mesh minimizes potential injury to the animal by decreasing the likelihood that the animal will entrap a body part in the mesh. Heavier, large-gauge mesh nets are used to capture larger nonhuman primates, up to 15 kg. When netting nonhuman primates, the following points should be remembered: • Safe netting of nonhuman primates requires experienced personnel wearing appropriate personal protective equipment. • For Old World and large New World species, safe netting requires at least two technicians. • Once the animal is netted, it is best to allow the animal time to calm down prior to attempting removal. This will minimize stress, struggling, and potential injury. • Captured animals are removed from a net either awake or after sedation with ketamine. The decision to use sedation to remove the animal from the net is determined, in part, by the reason the animal was netted and the potential for injury to the animal and the animal handler.

Pole and collar restraint system The pole and collar restraint system provides the animal care technician with a method to restrain, manipulate, and move conscious animals in a manner that minimizes potential stress and injury to both the animal and the animal handler. This system of restraint is used to handle a variety of species, including macaques, baboons, © 2002 CRC Press LLC

157–162

and squirrel monkeys. Most frequently, this system is used to move a conscious animal from his/her cage to a restraint device (stock or chair), whereupon the animal undergoes short-term manipulations such as blood collection or intravenous drug administration. Collars are composed of lightweight metal or sturdy plastic and contain two rings located opposite each other to which metal catch poles can be attached. Collars come in a variety of diameters and must be fitted to the animal. Poles come in lengths of 2 and 3 feet. Both collars and catch poles are commercially available (see Chapter 6). The procedure to remove an animal from a cage using the pole and collar system typically requires two animal care technicians. Procedure 1. The first technician places the catch pole through the cage bars at a location other than the door and clasps one of the two rings on the collar. Depending on the disposition and conditioning of the animal, the squeeze-back mechanism of the cage may need to be used to control and position the animal. 2. Once the animal is secured by the first technician, the second technician opens the cage door enough to advance a second pole into the cage to clasp the other ring. 3. The first technician then releases the collar, removes the pole from between the bars, places the pole through the open cage door, and clasps the empty ring on the collar. At this time, the door is fully opened and the animal is removed from the cage by both technicians and directed to a restraining device (Figure 5.5). 4. The animal is then placed in the restraining device, which in most cases is equipped with a grooved yoke that can accept the collar. 5. The collar is secured to the yoke of the restraining device. 6. For smaller and/or well-conditioned nonhuman primates, the second technician may remove the animal from the cage immediately after the first technician releases his catch pole. For safety reasons, two technicians and two poles are recommended when working with baboons and adult macaques.

Conditioning/Training The pole and collar restraint system is particularly advantageous because animals can be readily conditioned and trained to this system. Macaques and baboons have been trained in as few as four © 2002 CRC Press LLC

Fig. 5.5 Juvenile baboon being moved using a pole and collar restraint system.

or five 10-minute sessions,157 although the authors prefer at least a 10-session training period prior to initiation of a study. When conditioning nonhuman primates to the pole and collar restraint system, the following points should be remembered: • To successfully condition and train animals to the pole and collar system, the same person(s) should work with the animal(s) during the entire training period. • The trainers should be knowledgeable in the use of the pole and collar restraint method as well as the species behavioral repertoire. • During the first five training sessions (5- to 10-minute daily sessions) the animal is conditioned to the catch pole and being caught. At first, the animal may resist the catch pole and, in some cases, the squeeze-back mechanism of the cage might need to be used to control and position the animal.

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• During the second five training sessions, the animal is removed from the cage, taught to walk under the control of the pole, and placed in the restraining device. • After ten sessions, most animals readily accept being caught, removed from their cage, and placed into a restraint device. • The trainers should be patient, persistent, and consistent in their training, and should provide immediate rewards, (e.g., food treats), when the animal meets various training objectives.

Restraint stocks and tubes Restraint stocks and tubes are designed to restrict the movement of a conscious animal so that a technician can easily and safely perform such procedures as drug administration, blood collection, and physical examination. These devices are most often constructed of plastic materials. They can be purchased commercially, although many facilities have designed their own devices to meet the needs of their management practices and research projects.156, 162–166 Stocks are most frequently used to restrain Old World species such as rhesus and cynomolgus monkeys (Figure 5.6). Animals are usually moved from their cage and placed into the stock using the pole and collar restraint system. Tubes are most frequently used to restrain smaller New World species, such as marmosets and tamarins (Figure 5.7). Marmosets and tamarins are usually hand caught and placed into the restraint tube. Restraint stocks and tubes are designed to securely restrict an animal’s ability to move. Thus, an animal should not be maintained in these devices for extended periods of time. To minimize stress to the animal and the technician, it is advisable to condition animals to the restraint device prior to initiation of the study. Nonhuman primates should not be left unattended in a restraint stock or tube.

Restraint chairs Restraint chairs are used in the research laboratory environment for a variety of purposes, including drug administration, blood collection, and the collection of other biologic samples. These restraint devices are designed to allow for convenient and safe access to the animal while the animal is restrained in a comfortable position. Most chairs are adjustable so that they can accommodate anatomical differences between species and individual animals (Figure 5.8).

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Fig. 5.6 A plastic restraint box for macaques.

Fig. 5.7 A restraint tube for marmosets and tamarins. © 2002 CRC Press LLC

Fig. 5.8 A cynomolgus monkey in a restraint chair. (Courtesy of Primate Products, Inc.)

Restraint chairs are constructed of plastic or lightweight metals. They can be purchased commercially, although many facilities have designed or modified chairs to meet the needs of their specific research projects.151, 167, 168 Restraint chairs are used successfully to restrain a variety of species, including squirrel monkeys, marmosets, macaques, African green monkeys, and baboons.151 Old World species are usually moved from their cage to the restraint chair using the pole and collar restraint system; whereas New World species are usually handcaught and placed into the restraint chair. Most animals readily acclimate to a restraint chair. The acclimation process involves placing the animal in the chair for short periods of time. During this acclimation period, the time that the animal is left in the restraint chair is increased in an incremental manner. The use of positive reinforcement rewards such as food treats and fruit juices facilitates acclimation. Because restraint chairs readily accommodate a more natural postural position, these devices are often used for procedures (i.e., pharmacokinetics studies) that require an extended period of restraint. It should be noted that the standards set by the Animal Welfare Act specifically address the use of restraint devices with nonhuman primates. © 2002 CRC Press LLC

Briefly, the standards set forth by the Animal Welfare Act state that nonhuman primates should be restrained for the shortest period of time possible. In instances where long-term (defined as more than 12 hours) restraint is required, the nonhuman primate must be provided the opportunity daily for unrestrained activity for at least one continuous hour during the period of restraint, unless continuous restraint is required by the research proposal and approved by the Institutional Animal Care and Use Committee.66 Moreover, nonhuman primates should never be left unattended in a restraint chair.

Tether system The tether system was developed more than 20 years ago as a less restrictive form of restraint for chronically instrumented nonhuman primates. It is specifically designed to permit the continuous or intermittent administration of agents and the collection of biologic samples and physiologic data without imposing constraints on the activity and position of an animal in a cage. In general, tether systems consist of three basic components: a jacket or backpack, that protects the catheters as they exit the animal; a flexible metal cable, that protects the catheters as they travel between the animal and the cage exterior; and a swivel at the interface of the cable and cage that prevents the 169 catheters from twisting on themselves (Figure 5.9). Since the

swivel

Fig. 5.9 Diagrammatic representation of a tether system with a side-mounted swivel. © 2002 CRC Press LLC

inception of the tether system, numerous variations have been described and successfully applied to a variety of species.57, 170–178 Two important considerations in managing an animal on tether are acclimation and monitoring. It is recommended that nonhuman primates be acclimated to the tether system for at least 7 days prior to surgical instrumentation.177 Most animals readily acclimate to the tether system; however, those animals that do not acclimate should not be instrumented nor considered for projects involving tethering. Animals on tether require vigilant monitoring to ensure the physical integrity of the system. Failure of a tether system will not only affect the research project but could also jeopardize the health and wellbeing of the animal.

Manual restraint Manual restraint is considered to be a relatively inexpensive and quick method to manipulate conscious nonhuman primates for procedures that require short periods of restraint. Such methods are commonly used to manipulate New World species (marmosets, tamarins, and squirrel monkeys) and, to a lesser extent, smaller Old World species (cynomolgus and rhesus monkeys).33, 151, 162, 168 The decision to use manual restraint methods should take into account the potential risk of physical injury to the technician and the animal. Manual restraint of a conscious nonhuman primate by definition involves direct contact between the technician and the animal. During this period of direct contact, the technician is at an increased risk of being bitten, scratched, and/or exposed to bodily excretions and secretions from the animal. In the case of a macaque, this means the technician is at risk of being exposed to B virus. Moreover, during manual restraint, the nonhuman primate is at risk of physical injury should a technician react to a struggling animal in an overzealous manner. Finally, manual restraint can be very stressful to the animal and can result in alterations in a variety of physiologic 32, 151, 179–181 parameters. Because of these concerns, operant conditioning and chemical and physical restraint methods that minimize direct contact with the animal should be considered before a decision is made to use manual restraint methods. Other factors that should be considered as part of the decision process to use manual restraint methods include the species, size, sex, age, health status, disposition, personnel training, B virus status, study needs, and the presence of canine teeth.

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Hand-catching and manual restraint of conscious animals has been reported in nonhuman primates up to 10 kg.104 These authors use a 5-kg upper limit for hand-catching young healthy nonhuman primates. Only experienced, well-trained, technicians wearing appropriate personal protective equipment should attempt to hand-catch nonhuman primates. In general, callitrichids do not like being handled or restrained; however, they can be acclimated to such procedures with time. Trained personnel can restrain callitrichids by holding them in the palm of their hand such that the thumb and fingers are below the animal’s arms while encircling the animal’s chest (Figure 5.10). When restraining callitrichids with this method, care should be taken to not restrict the animal’s ability to breathe. Some facilities advocate the use of rubber or lightweight cloth gloves to catch marmosets because their teeth are delicate and can be damaged by biting leather gloves. Tamarins, however, should only be handled with leather gloves because they have long upper canines and can inflict 33 painful bites. The procedure to hand-catch and remove an Old World or a large New World species from a cage is as follows:

Fig. 5.10 Manual restraint of a common marmoset. © 2002 CRC Press LLC

Procedure 1. Appropriate personal protective equipment is worn, including leather gloves and protective arm sleeves. 2. The door to the animal room is always closed before attempting to hand-catch an animal. Ideally, a second technician is present to provide assistance should it be necessary. 3. The technician responsible for catching the animal uses the cage’s squeeze-back mechanism to bring the animal to the front of the cage. 4. The cage door is opened just enough for the technician to place his/her arm through the opening. 5. The technician reaches through the opening with the back of a closed fist presented to the animal. This technique significantly reduces the likelihood that the animal will be able to inflict a crushing injury (bite) to a fingertip. 6. The technician grabs one of the animal’s arms above the elbow. Ideally, the technician grabs the same arm of the animal that they are using to catch the animal (i.e., a right-handed technician should attempt to grab the animal’s right arm). This subtle step greatly facilitates the hand catching process by ensuring that the animal, upon removal from the cage, is in a position so that when the technician grabs the nonhuman primate’s free arm, the animal is facing away from the technician’s body. 7. Once a secure hold is established on the animal’s arm, the cage door is completely opened, the animal is removed from the cage, and the animal’s free arm is grabbed above the elbow. Grabbing the animal’s free arm can be facilitated by swinging the animal toward the front of an adjacent cage. As the animal reaches out to grab the cage with its free arm, the technician grabs the arm above the elbow. At this point, the animal should be facing away from the technician. 8. The nonhuman primate is now manually restrained by holding its arms behind its back (Figure 5.11). In some cases, it may be necessary for a second technician to assist with the restraint process by also holding the animal’s legs. 9. During manual restraint, care is taken to provide sufficient restraint without injuring the animal’s arms or shoulders.

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Fig. 5.11 Manual restraint of a macaque (note arms held behind animal’s back).

The procedure to return a manually restrained Old World or a large New World species to a cage is as follows: Procedure 1. The door to the animal room is always closed before attempting to return a manually restrained animal to its cage. Ideally, a second technician is present to provide assistance should it be necessary. 2. The cage door is placed in an open position and the squeezeback mechanism is placed at the back of the cage. The presence of a second technician facilitates having the door and squeezeback mechanism in the appropriate positions. 3. The technician restraining the animal places the animal through the door opening and releases the animal while a second technician closes the cage door. Most animals readily retreat to the back of the cage before turning around, thus providing sufficient time to shut the door.

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Chemical Restraint When it is compatible with the experimental study and the health status of the nonhuman primate, strong consideration should be given to the use of chemical restraint. Chemical restraint facilitates handling and reduces the potential for injury of animals and personnel.85, 182 The most frequently used drug for chemical restraint in the research environment is ketamine. During chemical restraint, care must be taken to monitor the physiologic status of the animal until recovery is apparent. (See Chapter 4 for more specific information on the use of chemical restraint agents and monitoring.) Chemical agents used for restraint can be delivered via several methods. The most common delivery method is the hand-held syringe used in conjunction with a squeeze-back cage. Pole syringes are useful if an animal is confined to a small area such as a cage, transfer box, or chute. Pole syringe systems come in a variety of lengths and many are equipped with safeguards to control the depth of needle penetration. Remote injection systems, such as capture pistols, rifles, and blow pipes, are useful in delivering immobilization agents to animals in large cages, pens, or free-range environments. However, they are seldom if ever needed in the typical research setting. Capture pistols and rifles have an approximate maximum range of 150 to 300 feet, respectively. Blow pipes are used to accurately deliver an immobilization agent up to 30 to 60 feet. Although blow pipe systems have a limited range, they are in many ways preferred over capture pistols and rifles because they are inexpensive, easier to maintain, quieter, and less likely to injure the animal and/or the user.183 Because remote injection systems can injure both the animal and the user, they should only be used by individuals trained and skilled in their use.

Operant Conditioning and Training Methods The use of operant conditioning and training methods, even in their simplest form, can be very important in the general management of nonhuman primates in a research setting. Many nonhuman primates can be trained to participate in routine procedures, thus reducing the stress associated with physical and chemical restraint. Nonhuman primates have been trained to participate in a variety of techniques including, but not limited to, venipuncture, injections, topical application of agents, fecal collection, perineum examination, vaginal cytology assessment, and transfer through cages and chutes 184–187 (Figure 5.12). © 2002 CRC Press LLC

Fig. 5.12 Stump-tailed macaque trained to present head for topical drug application. The initial training of animals can be labor intensive, taking weeks to train an animal to reliably perform a complex task. However, the benefits of conditioning are significant. The use of conditioning methods not only minimizes stress to the animal, but also decreases the likelihood of injury to the animal and the technician. In addition, the use of such methods eliminates the need to anesthetize an animal for a procedure that takes only a few seconds to perform, reduces the time required to obtain a sample, and reduces the use of pharmacologic restraint agents, while at the same time giving the animal a 184 greater degree of control over its environment. When conditioning nonhuman primates, the following points should be remembered: • Depending on the task, the initial training of the animal(s) can be labor intensive. • The training should be conducted under the supervision of someone familiar with the respective species’ behavioral repertoire and the basic principles of training. • To successfully condition and train animals, the same person(s) should work with the animal(s) during the entire training period. © 2002 CRC Press LLC

• The trainers should be patient, persistent, and consistent in their training of the animal and should provide immediate rewards, such as food treats, when the animal meets various training objectives. • Complex tasks must often be broken down into a sequence of events that must be learned in a step-wise manner. For more information on the principles associated with training nonhuman primates to assist with routine procedures, refer to Laule188, 189 and/or consult a behaviorist.

sampling techniques Blood Collection The volume of blood and the frequency of collection depend on the animal’s size, health status, and the needs of the experimental study. To minimize the detrimental effects of blood loss (i.e., hypovolemia and anemia), every effort should be made to collect the smallest volume necessary to meet the needs of the study. This is of particular importance with nonhuman primates that weigh less than 3 kg.190 There are two general guidelines for determining the maximum volume of blood that can be safely collected as a one-time sample.191 The first guideline recommends that a maximum one-time sample not exceed 15% of the animal’s blood volume (estimated to be 7% of an animal’s bodyweight).33 The second guideline recommends using the 10%–10% rule, which states that the maximum blood sample is 10% of an animal’s blood volume, which is estimated in this case to be 10% of the animal’s bodyweight. In essence, this means that the maximum one-time blood sample volume that can be collected is 1% of an animal’s bodyweight. Both guidelines yield the same approximate blood volume recommendation for a one-time sample collection. However, the 10%–10% or 1% bodyweight rule is the simplest and quickest method to determine the maximum one-time blood volume that can be collected from a nonhuman primate. To determine the maximum one-time volume of blood (in milliliters) that can be collected using this method, multiply the animal’s bodyweight (kg) by 1% (0.01), followed by 1000 (for conversion purposes), which is the same as multiplying an animal’s bodyweight (kg) by 10. Using this method, the maximum one-time blood volume that should be collected from a 5-kg rhesus macaque would be 50 ml (i.e., 5 × 10). © 2002 CRC Press LLC

Animals from which blood samples are repeatedly taken should have their hematocrit or CBC checked periodically for evidence of anemia. It is recommended that squirrel monkeys, which become anemic (defined as packed cell volume less than 40%) from repeated blood sampling, be supplemented with iron dextran (50 mg intramuscularly, once); folic acid 2.5 mg (subcutaneously, twice weekly); and vitamin B complex (subcutaneously, twice weekly, dose dependent on formulation) until the hematocrit reaches 40% or greater.32 In the case of callitrichids, some authors recommend that no more than 15% of the animal’s total blood volume be taken over the course of a month and that animals undergoing repeated or large sample collection be supplemented with iron.33 In the case of baboons, the authors supplement animals that undergo repeated or large blood sample collection with iron dextran (200 mg intramuscularly, once); vitamin B complex (subcutaneously, once, dose dependent on formulation); and/or chewable children’s vitamins administered daily throughout the course of the study. When collecting blood, a variety of blood collection vials can be used. Vials containing the anticoagulant EDTA (lavender top tubes) are used when collecting whole blood for cellular analysis, such as for a complete blood count (CBC). Vials containing the anticoagulants heparin (green top tubes) and citrate (light-blue top tubes) are used to collect whole blood if plasma is needed for analysis. Red top tubes do not contain an anticoagulant and are used to obtain serum for chemistry analysis. Some red top tubes contain a serum separator, which helps separate upon centrifugation the blood cells from the serum. A number of collection devices are used to obtain blood. Syringes and needles are most frequently used for venipuncture. Vacutainer ® blood collection systems (Becton-Dickson, Rutherford, NJ) are used with larger nonhuman primates (animals weighing more than 5 kg); however, the negative pressure created by large collection tubes (5 to 10 ml) can collapse veins in smaller animals. Butterfly needles are advantageous for saphenous venipuncture because the needle and hub can lie flat to the surface of the leg. Placement of indwelling 190 or vascular access ports can greatly facilitate repeated catheters blood sampling. Percutaneous blood sampling should follow these general guidelines: • A disposable needle and syringe should be used for blood collection. © 2002 CRC Press LLC

• Needles and syringes should be placed in a plastic container designated for sharp objects after use. Needles should not be recapped prior to disposal.80 • Smaller diameter needles should be used in smaller diameter vessels. • The size of the syringe to be used is dependent on the size of the animal and vessel, and the sample volume to be collected. • Syringes (3 to 10 ml) and needles (20 to 23 gauge) are typically used in macaques and baboons weighing more than 3 kg. • Syringes (1 to 3 ml) and needles (23 to 27 gauge) are typically used in adult callitrichids and squirrel monkeys.

Blood collection sites The most common site for the collection of large volumes of blood in the nonhuman primate is the femoral vein. Other collection sites include the cephalic, saphenous, coccygeal, and jugular veins.32, 33, 168, 190, 192–194 When repeated sampling is necessary, the collection site should be rotated. Nonhuman primates can be restrained for blood collection using physical or chemical restraint methods. In addition, nonhuman primates can be trained to present their arms or legs for venipunc180, 190 ture. Positioning of the animal for blood collection is dependent, in part, on the blood collection site as well as the restraint method. The femoral vein and artery are used to obtain relatively large quantities of blood. Venous blood is preferred for most samples; arterial blood is obtained for blood gas assessment. Procedure 1. The animal is placed in dorsal or dorsal lateral recumbency with the hindlimbs in extension. 2. The skin over the femoral triangle (junction of the muscles of the upper inner thigh with the lower abdomen) is cleaned with 70% ethyl alcohol. 3. A finger is placed in the femoral triangle and a pulse signifying the location of the femoral artery is identified. 4. A needle is inserted at a 45 to 60° angle to the skin, with the syringe barrel held so that the needle bevel is up. For an arterial blood sample, the needle is inserted directly over the palpable pulse. For a venous sample, the needle is inserted just medial to the palpable pulse (Figure 5.13). © 2002 CRC Press LLC

Fig. 5.13 Blood collection from the femoral vein of a nonhuman primate (note vein is medial to the artery).

5. Slight negative pressure is applied to the syringe barrel as the needle is advanced. 6. The needle is advanced into the vessel until a flash of blood occurs in the hub. 7. The blood sample is gently aspirated, the needle is removed, and firm pressure using gauze or a cotton ball is applied to the puncture site for 30 to 60 seconds. 8. For arterial samples, pressure is applied for 3 to 5 minutes.195 9. Arterial samples are differentiated from venous samples by color and pressure. Arterial blood is bright red and the pressure is usually sufficient in larger animals to begin to push back the syringe plunger; venous blood is dark red and the pressure is insufficient to push back the syringe plunger. The cephalic and saphenous veins are used to obtain small to moderate amounts of blood. The technique that is used to obtain blood from these vessels applies to all superficial veins.

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Procedure 1. The animal is usually placed in either ventral or lateral recumbency with the forelimb (cephalic vein access) or hindlimb (saphenous vein access) in extension. 2. The hair over the respective vessel is clipped and the skin is cleaned with 70% ethyl alcohol. 3. The respective vein is distended by compressing the vein closer to the heart than the venipuncture site. Compression is applied by a tourniquet or manually by an assistant. 4. For the cephalic vein, compression is usually applied at the crux of the elbow or just above the elbow (Figure 5.14). For the saphenous vein, compression is applied at the level of the knee or just above the knee (Figure 5.15). 5. The vein is visualized and/or palpated following distension. 6. The needle is inserted with the bevel up at an approximate 20 to 30° angle to the skin over the vessel. 7. In most cases, the needle can be inserted into the vessel in one motion. In some instances, the vessel will move or roll under the skin as the needle is inserted. In such cases, a thumb is placed along side the vessel and the needle is inserted under the skin to the side of the vessel (opposite of and angled toward the thumb). 8. Slight negative pressure is applied to the syringe barrel as the needle is advanced. 9. The needle is advanced until a flash of blood occurs in the hub. 10. Blood is aspirated by withdrawing the syringe plunger in a steady, gentle manner. If the flow of blood stops during the collection process, the needle is rotated slightly within the vein or the animal’s foot or hand is gently squeezed to stimulate venous blood flow. 11. Upon collection of the sample, compression of the vessel is released and the needle is removed. Digital pressure is applied to the venipuncture site for approximately 30 seconds.

Urine Collection The collection of urine from nonhuman primates can be accomplished using several methods. The method used is dependent, in part, on the volume needed, the sample type (sterile) needed, the © 2002 CRC Press LLC

Fig. 5.14 Blood collection from the cephalic vein of a nonhuman primate.

Fig. 5.15 Blood collection from the saphenous vein of a nonhuman primate.

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period of time that the urine is to be collected, and the species of nonhuman primate. Urine samples should be refrigerated until analyzed to prevent bacterial overgrowth.

Free-catch method The free-catch method is the simplest and least invasive way to obtain a urine sample. The primary disadvantage of this method is that the samples are almost always contaminated with food, water, and/or feces. To collect urine using the free-catch method, the animal is housed individually or confined to a trap area of a gang cage. A clean litter tray is placed under the cage or trap area. The tray is sloped to one corner to facilitate pooling and decrease loss due to evaporation. Screen material is placed over or incorporated into the dependent corner of the tray to separate food and feces from the urine. Modified rodent metabolic cages have been used to obtain 24-hour urine samples from marmosets and owl monkeys.190

Urethral catheterization Urethral catheterization is used to obtain uncontaminated urine samples. Urethral catheterization is possible in both female and male 190 macaques, guenons, baboons, and chimpanzees. Animals are typically catheterized while using chemical restraint methods or appropriate restraint devices. Placement of a urethral catheter requires aseptic technique to minimize the introduction of bacteria into the urinary tract. Appropriate aseptic technique includes the use of sterile gloves, catheter, and lubricant, as well as cleansing the area around the vulva or glans penis with a povidone-iodine surgical scrub followed by a sterile saline rinse. Experienced personnel may be able to aseptically advance a catheter out of its sterile package in a manner that does not require the use of sterile gloves. When placing a urethral catheter, the following points should be remembered: • Urethral catheters made of soft/flexible materials (rubber or vinyl) should be used because they cause less trauma to the urethra and bladder wall. Urethral catheters made of polypropylene are not recommended in male nonhuman primates because they 190 are too inflexible to pass around the ischial arch.

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• The ability to pass a catheter beyond the level of the ischial arch in male macaques, even when using urethral catheters composed of flexible materials, is variable and operator dependent.190 • The volume of urine collected is dependent on the last time the animal voided. Because many animals void upon being restrained for the administration of an anesthetic agent, the volume collected may be relatively small. • Typically, 3.5 to 9.0 French catheters are used for urethral catheterization of macaques and baboons.190, 196, 197 Procedure 1. Female animals are best positioned in ventral recumbency with their legs suspended over the edge of an examination table. 2. The animal’s tail is held to the side. 3. The region around the vulva is cleaned. 4. Sterile lubricant is applied to the tip of the catheter. 5. The urethral papilla is located on the ventral midline of the vulvar opening. In many cases, it can be readily visualized without the use of a speculum. Individual variation, traumatic injuries to the perineum, and changes in tumescence can alter the location and the ability to visualize the urethral papilla. In such circumstances, a vaginal or small nasal speculum with light source is used to facilitate visualization of the papilla. 6. The papilla is identified and a lubricated catheter is passed aseptically through the urethral orifice and advanced into the bladder. The length of catheter needed to enter the bladder is estimated as the distance between the perineum and the anterior aspect of the pubis. 7. Urine will usually appear in the catheter upon entering the bladder. 8. If no urine appears, a syringe is placed on the end of the catheter and gentle negative pressure is applied. If urine still does not appear upon aspiration, the catheter is either advanced another inch or slowly withdrawn while aspirating. The female nonhuman primate urethra is relatively short and it is not uncommon to advance too much catheter into the bladder, which results in kinking and obstruction. 9. The urine sample is collected, and the catheter is kinked and removed by applying gentle traction. Care is taken to control the tip of the catheter upon exiting the urethra to prevent splashing. © 2002 CRC Press LLC

Procedure 1. Male animals are best positioned in dorsal or dorsal lateral recumbency with their legs in extension. 2. The glans region of the penis is cleaned. 3. Sterile lubricant is applied to the tip of the catheter. 4. The length of catheter needed to enter the bladder is estimated by holding the catheter in the approximate position it will traverse to enter the bladder (i.e., from the tip of the penis and around the ischial arch to the anterior aspect of the pubis). 5. The penis is grasped and extended to erectile length, and a lubricated catheter is passed aseptically through the urethral orifice and advanced into the bladder. Slight to moderate resistance may be encountered as the catheter passes over the ischial arch. 6. Urine will usually appear in the catheter upon entering the bladder. 7. If no urine appears, a syringe is placed on the end of the catheter and gentle negative pressure is applied. If urine still does not appear upon aspiration, the catheter is either advanced 1 to 2 inches, or slowly withdrawn while aspirating in case the catheter is kinked within the bladder. 8. The urine sample is collected, and the catheter is kinked and removed by applying gentle traction. Care is taken to control the tip of the catheter upon exiting the urethra to prevent splashing.

Cystocentesis Cystocentesis involves the percutaneous placement of a needle into the bladder to obtain a urine sample. This technique is used to obtain uncontaminated urine samples and has been used successfully to collect urine from macaques, squirrel monkeys, and baboons.168, 198 It can be technically difficult to perform because, in many species, a significant portion of the bladder is located within the pelvic canal and many animals void upon being restrained — both of which make it difficult to palpate the bladder. The use of ultrasound is useful in training staff how to perform this technique, because it can provide the trainees with a visual image of the bladder and its location within the pelvic canal. When obtaining a urine sample via cystocentesis, the following points should be remembered: © 2002 CRC Press LLC

• The volume of urine collected is dependent on the last time the animal voided. Because many animals void upon being restrained for the administration of an anesthetic agent, the volume collected may be relatively small. • In macaques and baboons, a 1.5- to 2.0-inch, 20 to 22 gauge needle is typically used to obtain urine via cystocentesis. • In squirrel monkeys, a 0.5- to 0.75-inch, 20 to 22 gauge needle is used to obtain urine via cystocentesis.168 • Cystocentesis should not be performed in animals suspected of having endometriosis. Procedure 1. Animals are chemically restrained and positioned in dorsal recumbency. 2. The hair anterior to the pubis is shaved if warranted (many nonhuman primates have little hair in this region), and the skin is aseptically prepared. 3. An appropriate-sized sterile needle is inserted on the midline, 0.5 to 1.0 inch cranial to the anterior aspect of the pubis, and directed ventral and caudal at an approximate 45° angle to the abdomen (Figure 5.16).

Fig. 5.16 Diagrammatic representation of cystocentesis in a nonhuman primate. © 2002 CRC Press LLC

4. The needle is advanced until urine appears in the hub. The sample is aspirated, negative pressure is released, and the needle is withdrawn. 5. If blood, green material, or brown “chocolate-like” fluid is encountered at any time during the sample collection process, the needle is immediately withdrawn and replaced prior to reattempting sample collection. In the case of green material or brown “chocolate-like” fluid, a veterinarian should be contacted prior to re-attempting sample collection.

Bone Marrow Aspiration and Biopsy Bone marrow aspiration and biopsy techniques involve the introduction of a rigid, hollow needle into the marrow-containing cancellous part of either long or flat bones. A number of collection sites have been reported in the larger nonhuman primate species, including the 199 199, 200 dorsal anterior aspect of the iliac crest, the ischial tuberosity, 201 202 the trochanteric fossa of the femur, the tibial crest, and the proximal humerus. In general, collection sites in nonhuman primates are 190, 203–205 similar to those described in the dog. Animals undergoing bone marrow aspiration or bone biopsy should be appropriately anesthetized. Collection of a bone marrow aspirate or biopsy requires the use of aseptic technique. This includes shaving the skin over the collection site, preparing the collection site with a povidone-iodine surgical scrub, draping the site, and the use of sterile gloves. Special needles incorporating a central stylet are required for bone marrow collection. Several different styles are available (e.g., Rosenthal, Klima, Illinois sternal/iliac aspiration, and Jamshidi® (Baxter Healthcare Corp., Deerfield, IL); these needles vary in length, gauge (13 to 18), bevel, and handle style (Figure 5.17). In smaller New World species, 20gauge, 1.5-inch spinal needles can be used for bone marrow collection. Bone marrow core biopsies are best performed using a Jamshidi needle, minimum 14 gauge. When performing a bone marrow aspiration the following points should be remembered: • Syringes and needles should be washed with heparin prior to use. • Collection sites should be alternated in animals undergoing repeated sampling. • Analgesics should be used post-procedure. © 2002 CRC Press LLC

Fig. 5.17 Jamshidi® (left) and Illinois sternal/iliac (right) bone marrow aspiration needles.

The iliac crest is an ideal site for obtaining large volumes of bone marrow in the baboon and adult macaque (Figure 5.18).

Fig. 5.18 Diagrammatic representation of a macaque pelvis and femur demonstrating common bone marrow aspiration sites. © 2002 CRC Press LLC

Procedure 1. The animal is appropriately anesthetized and positioned in ventral recumbency. 2. The hair over the iliac crest (dorsum of the pelvis) is shaved, the skin is prepared with a povidone-iodine surgical scrub, and the site is draped. 3. The iliac crest is palpated and a small incision is made with a scalpel blade through the skin over the anterior aspect of the iliac crest. 4. The bone marrow needle is advanced through the incision until it rests upon the dorsal anterior aspect of the iliac crest. 5. The needle is then advanced ventrally into the iliac crest by applying firm, steady pressure while rotating the needle backand-forth. The needle should remain parallel to a sagittal plane of the iliac crest as it is advanced. 6. As the needle penetrates the cortex of the bone, it will meet less resistance and it will become well anchored (the needle will not wobble if it is rocked back-and-forth). 7. The stylet is removed from the needle and a 20-ml syringe is attached. Negative pressure is applied by forcefully pulling back on the syringe plunger, and bone marrow should begin to fill the syringe. 8. If no bone marrow is aspirated, the needle is rotated 90 to 180° and aspiration is re-attempted. If bone marrow still cannot be aspirated, the stylet is placed back in the needle, and the needle is either advanced or withdrawn slightly and aspiration is reattempted. 9. A bone marrow sample is dark red and more viscous than blood, and it contains pale bone spicules. 10. For diagnostic specimens, less than 1 ml is required. If larger volumes are needed, multiple sites should be used (3 to 5 ml per site can be collected before excessive dilution with blood occurs). 11. Upon collection of the bone marrow sample, the needle is removed and the small incision is closed with tissue glue or a suture. The ischial tuberosity is a useful collection site for small diagnostic bone marrow samples in macaques (Figure 5.18). This technique © 2002 CRC Press LLC

is, in essence, the same as that described for the iliac crest with the exception of the approach. For this technique, the animal is positioned in either lateral or ventral recumbency, with the hind legs flexed and drawn close to the animal’s body. The ischial tuberosity is palpated under the animal’s ischial callosity (pad), and the bone marrow needle is inserted through the pad toward the animal’s head and into the ischial tuberosity. The sample is collected as described above. The trochanteric fossa of the femur is a useful collection site in small animals (less than 5 kg; Figure 5.18). This technique is, in essence, the same as that described in the iliac crest with the exception of the approach. For this technique, the animal is positioned in lateral recumbency and the collector holds the proximal thigh in the palm of his/her hand with the thumb lying along the lateral aspect of the animal’s femur. The trochanteric fossa, which is located medial and slightly distal to the greater trochanter of the femur, is palpated. The needle is directed into the trochanteric fossa toward the shaft of the femur and into the medullary cavity. The sample is collected as described above. The proximal humerus is also a useful collection site in small animals (Figure 5.19). This technique is, in essence, the same as that described for the femur trochanteric fossa with the exception of the approach. For this technique, the animal is positioned in lateral

Fig. 5.19 Anterior view of the proximal humerus demonstrating bone marrow collection site via penetration through the intertrabecular groove (arrow). © 2002 CRC Press LLC

recumbency and the collector holds the proximal upper forelimb in the palm of his/her hand with the thumb lying along the lateral aspect of the animal’s humerus. The intratubercular groove, which is located between the greater and lesser tubercle of the humerus is palpated. The needle is directed into the groove toward the shaft of the humerus and into the medullary cavity. The sample is collected as described above. Core bone biopsies204 are obtained from the same sites from which bone marrow is aspirated. The primary difference between a bone biopsy and a bone marrow aspirate lies in what is done with the needle after it is firmly embedded in the bone. In the case of a bone marrow aspirate, the stylet is removed, a syringe is attached, and negative pressure is applied. In the case of a core bone biopsy, the stylet is removed and the needle is advanced further into the bone (1 to 2 cm) by gentle rotation and steady pressure. The needle is then loosened by both rotating and rocking the needle back-and-forth, and then the needle is partially withdrawn and re-introduced into the bone. The repositioning of the needle ensures that a larger sample is obtained and that the biopsy specimen is severed from the bone. To remove the biopsy sample, the needle is rotated and rocked back-and-forth and then withdrawn with steady traction and rotation. A wire or expeller is used to displace the bone biopsy specimen from the needle.

Cerebrospinal Fluid Collection Cerebrospinal fluid (CSF) is obtained from nonhuman primates via either cisternal (suboccipital) or lumbar puncture.190 Animals undergoing CSF collection are appropriately anesthetized and positioned in accordance with the site of collection. CSF collection requires the use of aseptic technique. This includes shaving the skin over the collection site, preparing the collection site with a povidone-iodine surgical scrub, draping the site, and the use of sterile gloves. When collecting CSF the following points should be remembered: • Most nonhuman primates have 12 thoracic vertebrae (T) and 7 lumbar vertebrae (L); however, the number of lumbar vertebrae present in the great apes and some New World monkeys can vary.40 • The spinal cord in nonhuman primates ends at T12. Therefore, lumbar puncture can be performed caudal to L1 without traumatizing the spinal cord.40

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• Differences have been reported in the composition of CSF taken from a lumbar site when compared to a suboccipital site.190 For purposes of comparison, samples should be collected from the same site. • In macaques and baboons a 1.5 to 2.0 inch, 18 to 22 gauge spinal needle is typically used to obtain CSF from a lumbar or suboccipital collection site. In squirrel monkeys, a 5/8 inch, 25 gauge needle is used to obtain CSF from a suboccipital collection site.32 Procedure 1. The animal is appropriately anesthetized and positioned in lateral recumbency. 2. To widen the intravertebral space and thus facilitate placement of the spinal needle, the back is flexed by drawing the animal’s hindlimbs forward toward the umbilicus, while at the same time drawing the forelimbs backward toward the umbilicus (Figure 5.20). 3. The hair over the lumbar spine is shaved, the skin is prepared with a povidone-iodine surgical scrub, and the site is draped. 4. An intervertebral space between the dorsal processes of two lumbar vertebrae is palpated (usually L2–L3 or L3–L4).

Fig. 5.20 Diagrammatic representation of a baboon positioned to increase the intervertebral space for lumbar puncture. © 2002 CRC Press LLC

5. The spinal needle is inserted into the center of the intervertebral space just behind the anterior vertebral spinous process. The needle is directed anteriorly at an approximate 70° angle from the long axis of the back (Figure 5.21). 6. The needle is advanced through the skin and underlying tissue. As the needle is advanced, care is taken to ensure that the tip of the needle does not deviate off midline. 7. If bone is encountered as the needle is advanced, the needle is redirected either anteriorly or posteriorly. 8. As the needle is advanced, the stylet is removed periodically to see if CSF appears in the needle hub. Entry into the subarachnoid space (location of the CSF) is usually sensed by a sudden loss in resistance against the needle (a “pop”).

Fig. 5.21 Diagrammatic representation demonstrating placement of a spinal needle into the subarachnoid space. © 2002 CRC Press LLC

9. Once the needle is in the subarachnoid space, CSF should flow freely through the needle. CSF can be collected via gravity flow into a tube, or by placing a tuberculin syringe next to the spinal needle hub (not attached) and drawing back on the syringe plunger as the fluid wells up in the hub. If blood is encountered, the needle is removed and a new needle is used to attempt CSF collection from a different intervertebral space. 10. Upon collection of the sample, the needle is removed and direct pressure is applied to the puncture site. Procedure 1. The animal is appropriately anesthetized and positioned in either lateral recumbency or an upright sitting position. 2. The hair over the back of the head and the anterior cervical spine is shaved, the skin is prepared with a povidone-iodine surgical scrub, and the site is draped. 3. An assistant flexes the neck so that the animal’s chin nearly touches its chest. This increases the space used to access the cisterna magna with the needle. During flexion of the neck, care is taken not to obstruct the animal’s airway. 4. The occipital protuberance on the back of the skull and the wings of the atlas (C1) are palpated. 5. The spinal needle is inserted through the skin and underlying tissue on the midline at a point midway between the occipital protuberance and C1. 6. If bone is encountered, the needle is redirected either anteriorly or posteriorly. 7. Entry into the cisterna magna is usually sensed by a sudden loss in resistance against the needle (a “pop”). If the brain or spinal cord is penetrated, the animal will jerk or twitch. 8. Once the needle is in the subarachnoid space, CSF should flow freely through the needle. CSF is collected via gravity flow into a tube or by placing a tuberculin syringe next to the spinal needle hub (not attached) and drawing back on the syringe plunger as the fluid wells up in the hub. 9. Upon collection of the sample, the needle is removed and direct pressure is applied to the puncture site.

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Semen Collection Semen has been collected from a variety of nonhuman primate species using several techniques. The technique used to collect a sample can have a marked effect on semen morphology, viability, motility, numbers, and function, and therefore should be thoroughly investigated prior to choosing a method. Rectal probe electro-ejaculation methods are successfully used to obtain semen from callitrichids, squirrel monkeys, macaques, baboons, and great apes. These techniques require repetitive electrostimulation of the prostate and accessory sex glands, and are performed under anesthesia.206–211 Direct penile electrostimulation methods are used successfully to obtain semen from macaques. These methods require the direct application of repetitive electrostimulation to the penis through electrodes placed around the circumference of the penis near the glans and at the base of the penis. The use of non-metal electrodes minimizes potential burns to the skin of the penis. This technique is performed with animals that are chair restrained or under light ketamine sedation.211–214 Vibratory stimulation methods are also used in squirrel monkeys to obtain semen samples. These methods of collection produce superior semen samples in the squirrel monkey when compared to samples collected by rectal probe electro-ejaculation methods, and do not require anesthesia.207 Other methods used to obtain semen samples in nonhuman primates include the use of vaginal washing techniques in cal206 and the training of great apes to masturbate or use an litrichids artificial vagina.210

Amniotic Fluid Collection Amniotic fluid samples are obtained by placing a needle into the fluid-filled compartment in which the embryo/fetus is suspended. The technique to collect amniotic fluid is referred to as amniocentesis; it has been successfully performed in both macaques and 216–218 baboons. There are two percutaneous transabdominal amniocentesis techniques reported in the literature. One technique uses ultrasound to assist in needle placement,215 whereas the other employs a “blind stick” method.216, 217 For both techniques, the animal should be appropriately anesthetized and positioned in dorsal recumbency. © 2002 CRC Press LLC

Animals in the last trimester are rotated slightly to the left to minimize compression of the vena cava by the uterus. Collection of amniotic fluid requires the use of aseptic technique. This includes shaving the skin over the caudal abdomen, preparing the skin with a povidone-iodine surgical scrub, draping the site, and the use of sterile gloves. In the case of the ultrasound assisted technique, sterile ultrasound gel is used on the abdomen, and the scan head of the ultrasound probe is covered with ultrasound gel and placed into a sterile glove or sheath. Volumes of 1 to 10 ml can be safely collected from macaques between gestational days 60 and 150, respectively. Various needle sizes are used to collect amniotic fluid, including 1.0–1.5-inch, 22–23-gauge needles, and 3.0–4.0-inch, 22–25-gauge spinal needles.215, 217 For the ultrasound-assisted technique, the ultrasound probe is placed on the abdomen and the uterus is scanned to determine the location of the placenta, fetus, and bladder. A needle is inserted through the skin of the abdomen over an area of the uterus devoid of placenta. The needle is advanced through the muscles of the abdomen, the uterus, and into the amniotic sac. Care is taken not to hit the fetus with the advancing needle. Biopsy guides can be attached to the ultrasound probe to facilitate needle placement; however, the use of a “free-hand” technique provides greater flexibility and allows for imaging in multiple planes during the advancement of 7, 74 the needle. Amniotic fluid is obtained by gently aspirating with a 5- to 10-ml syringe. The fluid should be clear. Upon collection of the sample, the negative pressure on the syringe plunger is released and the needle is withdrawn. Amniotic fluid can be obtained using this technique as early as day 60 of gestation. The “blind stick” method is used to obtain amniotic fluid from macaques and baboons between days 80 and 156 of gestation.216, 217 This technique is most easily performed after day 110 of gestation because the fetus can be readily palpated. With one hand, the sample collector palpates the caudal abdomen (lower uterine segment) and moves the fetus and/or fetal parts toward the mother’s head to create an area within the uterus devoid of the fetus. This is done by gently pressing down on the uterus just anterior to the pubis and then moving the hand toward the mother’s head. The fetus is held in this anterior position while the collector inserts a needle with the other hand into the animal’s midline, 2 cm anterior to the pubis, and directed anteriorly at a 45° angle. The amniotic fluid is collected as described in the ultrasound-assisted method. Although few complications with © 2002 CRC Press LLC

this technique have been reported in the literature, the lack of visualization of the uterus and fetus increases the potential risk of trauma to the placenta and fetus.

compound administration Compounds are administered to nonhuman primates via both parenteral and oral methods. Parenteral refers to the administration of drugs by means other than through the gastrointestinal system — usually via injection. Parenteral techniques allow the compound to enter the vascular system more directly than it would through the gastrointestinal system. The decision regarding which administration method to use depends on several factors, including the compound’s pH, solubility, absorption rate, effect on tissue, and the volume to be administered. In addition, the size, health status, and disposition of an animal, as well as the available housing and restraint systems and the technical skill required to deliver an agent via a specific route, also play an important role in determining which route is used. Drug formularies and package inserts of therapeutic agents should be consulted to determine an agent’s optimal route of administration. Recommendations for injection site volumes and needle sizes are found in Table 5.1.218

TABLE 5.1: RECOMMENDED INJECTION VOLUMES Species

IV

IP

IM

AND

NEEDLE SIZESa SC

ID

Marmoset

Lateral tail vein,0.5–1 10–15 ml, ml (slowly), ≤23 gauge ≤23 gauge

Anterior thigh/post- Scruff, 5–10 0.05–0.1 ml, erior thigh, ml, ≤23 ≤25 gauge 0.3–0.5 ml, gauge ≤23 gauge

Baboon

Cephalic vein, saphenous 50–100 ml, vein, 10–20 ≤21 gauge ml, ≤20 gauge

Anterior thigh/postScruff, erior thigh, 10–30 ml, triceps, 1–3 ≤20 gauge ml, ≤21 gauge

a

0.05–0.1 ml, ≤25 gauge

Note: a 23-gauge needle has a smaller diameter than a 20-gauge needle (≤ sign refers to needle diameter.) © 2002 CRC Press LLC

Parenteral Administration Methods Intravenous administration In the laboratory environment, the intravenous (IV) route is frequently used in nonhuman primates to administer specific antibiotics, anesthetics (such as propofol and barbiturates), and various compounds as part of pharmacokinetic studies. In addition, this is the most common and effective route by which fluid therapy is administered. Compounds can be delivered IV via a number of delivery systems, including a syringe and needle, a percutaneous peripheral catheter, or a surgically placed central venous line or vascular access port. A needle and syringe is typically used for the one-time IV administration of a compound. Percutaneously placed peripheral catheters are used for the intermittent or continuous IV administration of compounds over a short period of time. Surgically implanted central venous lines are used in the chronic continuous IV administration of compounds. A description of such delivery systems can be found in the tether system references under restraint in this chapter. Surgically implanted vascular access ports are used in the chronic intermittent IV administration of compounds. A detailed description of such a delivery system can be found at the end of this section under vascular access ports. The three basic types of IV catheters used to administer compounds through peripheral vessels are the over-the-needle, the through-the-needle, and the butterfly. With the over-the-needle type, the catheter passes over the needle. With both the over-the-needle and the through-the-needle type catheters, the needle is removed following penetration of the vessel. The butterfly catheter consists of a needle with plastic flaps permanently attached to a flexible tube. With a butterfly catheter, the needle remains in the vessel during the intravenous administration of a compound.219 When using percutaneously placed peripheral catheters for the IV administration of compounds, the following points should be remembered: • Peripheral catheters are preferred over a syringe and needle for the IV administration of compounds that are irritating to tissues. • Peripheral catheters used to administer an irritating compound should be flushed with saline prior to removal.

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• Over-the-needle and butterfly catheters are easier to place than through-the-needle catheters. • Butterfly catheters are more likely to traumatize the inner wall of a vessel than over-the-needle or through-the-needle catheters. • Peripheral catheters are difficult to maintain in a nonhuman primate for any length of time unless the animal is in a restraint box or chair, anesthetized, sick, or has a protective cast over the catheter site.220 • Peripheral catheters should be removed within 72 hours of placement.221 The peripheral vessels most frequently used for the IV administration of compounds in nonhuman primates are the saphenous and cephalic veins. The saphenous vein is ideal for IV injections and catheterization. It is large, easily visualized, and distant from the animal’s head and mouth. Vessels should be alternated in animals undergoing frequent IV administration of compounds. The technique to administer a compound IV using a syringe and needle is essentially the same as the technique to collect blood from a peripheral vessel, with the exception that compression of the vessel is released prior to administration. In addition, the IV administration of a compound should be stopped if swelling develops at or near the puncture site during the injection process. This indicates that the needle is no longer in the vessel and that the compound is being administered into the perivascular space. The perivascular administration of a compound may warrant the infiltration of the injection site with saline and a local anesthetic to minimize inflammation and discomfort. Following is a description of the procedure to place an over-theneedle catheter in a peripheral vessel. Procedure 1. The animal is appropriately restrained in a position that allows access to the respective vein. 2. The hair over the vein is clipped and the skin cleaned with 70% ethyl alcohol. If the catheter is to be maintained for an extended period of time, the site is prepared with a povidone-iodine surgical scrub. 3. The vein is distended by compressing the vein closer to the heart than the catheter entry site. Compression is applied with a tourniquet or manually by an assistant.

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4. For the cephalic vein, compression is usually applied at the crux of the elbow or just above the elbow. For the saphenous vein, compression is applied at the level of the knee or just above the knee. 5. The vein is visualized and/or palpated following distension. 6. The catheter is inserted with the bevel up at an approximate 20 to 30° angle to the skin over the vessel. In most cases, the catheter can be inserted into the vessel in one motion. In some instances, the vessel will move or roll under the skin as the catheter is inserted. In such cases, a thumb is placed along side the vessel and the needle is inserted under the skin to the side of the vessel (opposite of and angled toward the thumb). 7. The catheter is advanced into the vein (approximately 1/4 to 1/2 inch) beyond the point where blood first began to flow into the hub of the needle/stylet. 8. The needle/stylet is held stationary as the catheter is slowly advanced into the vessel until the catheter hub reaches the skin puncture site. 9. Compression of the vein is released, and the needle is withdrawn while holding the catheter hub stationary. 10. An injection cap is placed on the catheter, and the catheter is secured to the limb with tape and flushed with heparinized saline. If the catheter is to be maintained for an extended period of time, antimicrobial ointment and sterile gauze are placed over the puncture site and the entire limb is bandaged. 11. The compound to be given IV is administered by a syringe and needle through the injection cap. 12. If irritating compounds are administered through the catheter, the catheter is flushed with saline prior to removal. 13. Upon removal of the catheter, pressure is applied to the exit site for approximately 30 seconds.

Intramuscular administration The intramuscular (IM) administration of compounds is the most common parenteral administration route used in nonhuman primates. Squeeze-back cages facilitate the IM administration of agents in the laboratory environment. Antibiotics, analgesics, and anesthetics are commonly administered via IM injection. © 2002 CRC Press LLC

A number of muscle groups are suitable for intramuscular injection (Figure 5.22). The muscle group used is dependent, in part, on the size of the animal, the limitations of the available restraint system, and the prior conditioning of the animal. Muscle groups should be alternated in animals receiving frequent IM injections. The following muscles/muscle groups are suitable for IM injection: • Anterior muscles of the thigh. This muscle group is the most frequently recommended IM injection site in the literature.190, 221, 222 • Hamstring. This muscle group, which includes the large muscles of the caudal thigh, is a frequently used IM injection site in nonhuman primates,190 although this site is not recommended in the veterinary literature.221, 222 There have been reports of lameness, paralysis, and self-mutilation resulting from the administration of compounds near the sciatic nerve, which runs along the posterior 223 aspect of the femur. If the hamstring is to be used as an injection site, care must be taken not to deposit the compound near the caudal aspect of the femur (location of the sciatic nerve). To minimize injecting near the sciatic nerve, the point of the needle should always be directed away from the caudal aspect of the femur.

Fig. 5.22 Common intramuscular injection sites: A = anterior muscles of the thigh; G = gluteal muscles; H = hamstring; T = triceps. © 2002 CRC Press LLC

• Triceps. This muscle serves as an alternative site for IM injections in larger species.190 • Gluteal muscles. This muscle group, which includes the muscles of the buttocks, also serves as an alternative site for IM injections in larger species.190 Procedure 1. The animal is appropriately restrained. 2. The muscle group is visualized and the needle is quickly inserted through the skin into the muscle group at a 45 to 90° angle. 3. Before injecting, it is recommended that the plunger of the syringe be retracted to determine that the needle is not in a blood vessel. If blood appears in the needle hub, the needle is withdrawn and a different injection site is selected. It should be noted that the available restraint system and the disposition of the nonhuman primate often make it impractical to swab the injection site with 70% ethyl alcohol and retract the plunger prior to injection. 4. The compound is injected in a steady motion and the needle is withdrawn.

Subcutaneous administration The subcutaneous (SC) administration route is used most frequently to deliver fluids when intravenous administration is not practical or critical. The skin over the dorsum (shoulders and back) is relatively loose, making it an ideal site for the SC administration of compounds, particularly large volumes of fluids. Procedure 1. For an animal restrained in a squeeze-back cage, an SC injection is administered through the cage bars by inserting a needle, without picking up a skin fold, at a 10 to 20° angle through the skin and into the subcutaneous space. The compound is injected in a steady motion and the needle is withdrawn. 2. For a sedated animal or an animal in a restraint box/device, a fold of skin is typically picked up over the animal’s back, a needle is inserted at a 45° angle into the base of the skin fold, and the compound is injected in a steady motion and the needle withdrawn. © 2002 CRC Press LLC

3. Before injecting, it is recommended that the plunger of the syringe be retracted to verify that the needle is not in a blood vessel. If blood appears in the needle hub, the needle is withdrawn and a different injection site is selected. It should be noted that the available restraint system and the disposition of the nonhuman primate often make it impractical to swab the injection site with 70% ethyl alcohol and retract the plunger prior to injection. 4. If saline is to be administered via gravity (with a fluid administration set and bag), the placement of the needle within the subcutaneous space may need to be manipulated in order to maximize the flow rate. Large volumes of saline should be administered in multiple sites.

Intradermal administration Intradermal (ID) injections are most frequently used for testing purposes. In nonhuman primates, tuberculin is administered via the ID administration route. Tuberculin is most frequently administered into the skin of the upper eyelid or ventral abdomen. For more information on tuberculin testing, see Chapter 4. Procedure 1. The animal is appropriately restrained so that access to the intradermal (ID) injection site is readily available. 2. If applicable, the skin over the injection site is shaved. 3. The needle, with the bevel up, is inserted at a 10 to 20° angle until the bevel is completely enclosed within the layers of the skin (Figure 5.23a). Typically, 1/2 to 5/8 inch, 25 to 27 gauge needles are used for ID injections. 4. Proper placement of the needle is determined by lifting up on the needle tip. If properly placed, the metal of the needle tip is just visible through the skin. 5. A small volume of the compound is slowly injected into the intradermal site. If the procedure is performed correctly, a small translucent “bleb” will appear at the injection site (Figure 5.23b). The typical intradermal injection volume for a compound is 0.05 to 0.1 ml. 6. The use of soaps or disinfectants to clean intradermal injection sites can irritate the skin and interfere with interpretation of the 222 test of interest. © 2002 CRC Press LLC

Fig. 5.23 (a) Proper position of a syringe and needle for the intradermal administration of tuberculin into an eyelid. (b) Bleb that appears at the intradermal injection site if tuberculin is administered appropriately.

Vascular access port Implantable vascular access ports (VAPs) are used to deliver compounds intravenously in a variety of nonhuman primates, including macaques, squirrel monkeys, and marmosets.224–228 VAPs are ideal for the chronic intermittent intravenous administration of cytotoxic or irritating compounds. These systems can also be used to collect blood and monitor blood pressure. VAP systems consist of an indwelling catheter attached to a rigid puncturable port. The port consists of a rigid base and a silicone rubber window (septum) (Figure 5.24). The rigid base is designed so that it can be anchored to the underlying subcutaneous tissues and be easily located under the animal’s skin. The rigid base also prevents needle penetration through the port into the animal. The septum, through which the needle penetrates to access the port, is designed so that the penetration site closes up after the needle is removed. © 2002 CRC Press LLC

Fig. 5.24 Vascular access port without catheter (bar = 1.0 cm).

This feature allows VAPs to be accessed up to 2000 times before failure. The number of times a VAP can be accessed depends on the specific composition of the material that comprises the septum. VAP systems must be surgically implanted using strict aseptic technique. For intravenous compound administration, the catheter is typically placed in the jugular or femoral vein and the access port is implanted into the subcutaneous tissue over the animal’s back. If the primary use of the VAP is to collect blood or monitor blood pressure, the catheter is typically implanted into the femoral artery. VAP systems can remain patent for long periods of time. The duration of patency depends on the frequency with which the system is accessed and the maintenance program to prevent thrombosis. Many facilities have developed their own VAP flushing/locking solutions and maintenance protocols. Examples of locking solutions include heparinized saline solutions ranging in concentration from 100 to 500 U/ml, and heparinized dextrose solutions containing antibiotics (50% dextrose, 100 U/ml heparin, and 1 mg/ml vancomycin).229, 230 Current recommendations on the frequency with which VAPs should 229 be flushed is every 2 to 3 weeks. Things to remember when using a VAP: • The volume of the catheter and port reservoir should be determined prior to implantation. © 2002 CRC Press LLC

Fig. 5.25 (a) Diagrammatic representation of the tip of a standard needle, (b) and a noncoring Huber point needle.

• Only a Huber point needle, which is designed to preserve the integrity of the septum, should be used to access the port (Figure 5.25).224 • Strict aseptic technique should be used when accessing a VAP. • The septum should not be depressed unless accessing the port. Depression of the septum will displace a small amount of the locking solution from the catheter, thereby increasing the potential for catheter obstruction.231 • The skin over the VAP should be moved so that the needle penetrates a different site each time the port is accessed. Rotation of the skin-puncture site minimizes tissue irritation and necrosis. • The saline flush, locking solution, and the compound to be administered should not precipitate when mixed. Procedure 1. The animal is appropriately restrained and positioned so that access to the subcutaneous port can be readily obtained. 2. The hair over the access port is shaved and the skin is aseptically prepared using a povidone-iodine surgical scrub. 3. The port is located by palpation and the edges of the base are stabilized with a thumb and index finger. 4. A noncoring needle (Huber point) is pushed firmly into the VAP. The needle is in place when the tip reaches the bottom of the port and a “click” is felt. 5. An injection cap or stopcock is attached to the hub of the needle to facilitate the aseptic changing of syringes during compound administration and blood withdrawal. © 2002 CRC Press LLC

6. To verify placement and patency, an empty syringe is attached to the needle through the injection cap or the stopcock, and negative pressure is applied. 7. For compound administration, aspiration is stopped as soon as blood is observed in the syringe. The port is flushed with saline, at least two times the volume of the VAP, to flush out any residual locking solution and ensure patency. The syringe with the compound to be administered is attached to the needle and the agent is slowly infused. 8. For blood withdrawal, 1.5 to 2.0 times the volume of the VAP is aspirated to remove the locking solution. Following removal of the locking solution/heparinized blood, an empty syringe is attached to the needle and the blood sample is collected. 9. After both compound administration and blood withdrawal, the VAP is flushed with saline. The VAP is flushed, using pulsatile bursts, with at least two times the volume of the system. The pulsatile bursts create turbulence, which more effectively purges the VAP than does a continuous infusion. 10. After flushing, the VAP is infused in a pulsatile manner with a locking solution (the amount depends on the volume of the port and catheter). 11. As the appropriate volume to be infused is reached, the Huber needle is removed from the VAP while positive pressure is applied to the syringe plunger. This prevents reflux of blood into the catheter tip and potential occlusion. Note: Occlusion is the most common problem encountered with VAPs. If the VAP appears to be occluded, contact the attending veterinarian. Do not try to clear a VAP by over-pressurizing the system as this could result in catheter rupture and embolization.

Osmotic pumps Implantable osmotic pumps are used to continuously deliver drugs, hormones, and other agents at controlled rates for up to 4 weeks without the need for external connections or frequent handling of the animal (Figure 5.26). Osmotic pumps are usually implanted subcutaneously or intraperitoneally. A catheter or cannula can be attached to a pump to administer a compound via very specific routes (e.g., 232–236 The intravenous, intracerebral, intrathecal, or intraoviduct). pumps are placed surgically or via a large-bore needle using aseptic © 2002 CRC Press LLC

Fig. 5.26 Disassembled and assembled osmotic minipumps (bar = 1.0 cm). technique. The degree of technical difficulty to implant a pump is dependent on the size of the pump and the route of administration. For example, in the case of an intracerebral administration route, the pump is placed in a subcutaneous location over the back of the animal’s neck; however, a cannula from the pump would have to be surgically placed within the appropriate anatomical location of the brain.

Oral Administration Methods Self-administration The use of methods that promote self-administration are the easiest and least stressful way to orally administer compounds to nonhuman primates. Many medicants — such as vitamins, anti-inflammatories, anti-histamines, and even some antibiotics — come in children’s formulations that nonhuman primates often find quite palatable. Some companies (see Chapter 6) market a variety of medicated treats for nonhuman primates. These companies can often custom-formulate medicated treats to meet the needs of the investigator, research project, and/or the facility’s management practices. When using an oral self-administration method, the following points should be remembered: © 2002 CRC Press LLC

• There is minimum control over whether an animal will receive the entire dose of a compound. • Many animals undergoing treatment are sick and, consequently, have decreased appetite. • If it is important that a group of animals be treated the same (individual animals have different taste preferences) or that an animal receive a known therapeutic dose, other delivery methods should be considered. • The volume, form, solubility, and taste of the compound, as well as the taste preference and disposition of the animal, must be taken into account in determining the appropriate method of selfadministration. • If noncommercial self-administration methods are used, it is important to ensure that the compound is equally distributed throughout the palatable material. Noncommercial self-administration methods are only limited by the imagination and creativity of the animal carestaff administering the agent, and the creativity of the nonhuman primate to circumvent the delivery system. Some examples of noncommercial self-administration methods include: • Compounds can simply be administered in a piece of the animal’s favorite fruit. • Compounds can be administered in a palatable paste spread on bread, examples include jelly, Marmoset Jelly®, mashed banana and honey, peanut butter, and yogurt. • Compounds can be administered in the form of a frozen yogurt cup or a Jell-O® jiggler (Kraft Foods, Inc., Rye Brook, NY). • Water-soluble compounds can be administered in a palatable liquid delivered through a water bottle or by moving an animal to the front of the cage with the squeeze-back mechanism and squirting the liquid into the animal’s mouth. Examples of palatable liquids ® ® include Tang (Kraft Foods, Inc., Rye Brook, NY), Crystal Light ® (Kraft Foods, Inc., Rye Brook, NY), Kool-Aid (Kraft General Foods, Inc., White Plains, NY), Gatorade®, Prang, Ensure®, and fruit juices.

Nasogastric and orogastric gavage The administration of compounds via nasogastric and orogastric gavage is indicated when an unpalatable or large volume of a compound © 2002 CRC Press LLC

must be delivered orally. These techniques are also indicated when animals are anorexic or receiving a compound as part of a research protocol that requires verification that the entire dose is received. Nasogastric and orogastric gavage can be performed in awake or sedated animals. Awake animals should be placed in a restraint chair or box prior to gavage. If chemical restraint is used, light sedation (ketamine, 5 to 10 mg/kg) is preferred because it does not completely eliminate the animal’s swallow reflex, thus facilitating placement of the tube in the stomach. Orogastric gavage methods are used in smaller New World species such as marmosets and tamarins, while either orogastric or nasogastric methods are used in larger New World and Old World species. Five to eight French stomach tubes are typically used for the orogastric administration of compounds in marmosets, tamarins, and squirrel monkeys, whereas 16 French tubes are used in adult macaques and baboons. For the nasogastric administration of compounds, five French stomach tubes are typically used in squirrel monkeys and 12 to 16 French tubes are used in adult macaques and baboons. A reasonable maximum volume to administer into the stomach of a nonhuman primate is 10 to 20 ml/kg of bodyweight. Procedure 1. The animal is appropriately restrained. 2. The length of stomach tube to insert is estimated by measuring, with the tube, the distance between the animal’s mouth and last rib. 3. A dab of lubricating jelly is placed on the tip of the stomach tube. 4. With one hand, the technician holds the animal’s head in either a normal postural position or in extension while, with the other hand, the technician inserts the tube into a nostril in a ventral medial direction. 5. The tube is advanced through the nasal cavity until the appropriate distance, as determined in step 2 above, is reached. 6. If resistance is encountered in the nasal cavity, the tube is removed and re-inserted. If resistance continues to be encountered, the tube is removed and inserted into the other nostril, or a smaller diameter tube is used. If the tip of the tube is beyond the nasal cavity and the animal begins to cough or gag, the tube is removed and re-inserted. © 2002 CRC Press LLC

7. Placement of the tube into the stomach is confirmed by attaching a 10- to 20-ml syringe to the tube and applying negative pressure. If the tube is properly placed, stomach contents will appear in the tube (Figure 5.27). Proper placement of the tube can also be confirmed by rapidly instilling 5 to 10 ml air into the stomach tube while auscultating the animal’s upper-left abdominal quadrant. If the tube is in the stomach, gurgling sounds will be evident. Compounds should not be administered through a stomach tube without prior confirmation that the tube is in the appropriate location. 8. The compound is slowly administered through the stomach tube. If the animal begins to cough or exhibit signs of respiratory distress during the administration of the compound, administration is stopped, the tube is kinked to prevent backflow, and the tube is removed.

Fig. 5.27 Aspiration of stomach contents from a sedated macaque to verify proper placement of stomach tube. © 2002 CRC Press LLC

9. Upon administration of the compound, water or air (approximately two times the volume of the tube) is used to flush residual compound from the tube into the stomach. 10. The tube is then kinked to prevent backflow and is removed in one motion. To prevent splattering, care should be taken to control the tip of the tube as it exits the nostril. 11. The stomach tube is discarded after gavage. Procedure237 1. The animal is appropriately restrained. 2. The length of stomach tube to insert is estimated by measuring, with the tube, the distance between the animal’s mouth and last rib. 3. A dab of lubricating jelly is placed on the tip of the stomach tube. 4. With one hand, the technician holds the animal’s head in either a normal postural position or in extension, while with the other hand the technician places a speculum into the animal’s mouth. 5. A second technician inserts the stomach tube into the animal’s mouth and over or through the speculum, depending on the type used (Figure 5.28). The tube is advanced until the appropriate distance, as determined in step 2 above, is reached.

Fig. 5.28 Example of a simple mouth speculum (dog chew bone) over which a stomach tube is passed (top) and a mouth speculum237 through which a stomach tube is passed (bottom). © 2002 CRC Press LLC

6. Confirmation of placement, administration of the compound, and removal of the stomach tube are the same as described for nasogastric gavage. 7. The stomach tube is discarded after gavage and the mouth speculum is thoroughly disinfected prior to use in another animal.

Orogastric gavage of capsules Some study designs may require the orogastric gavage of capsules in nonhuman primates to simulate the manner in which the compound is administered to humans. Capsules can be administered to nonhuman primates using a technique similar to the orogastric administration of fluids. Orogastric capsule delivery is best performed using a Sovereign® rubber feeding tube (Kendall Co., Mansfield, MA.) Procedure 1. The animal is appropriately restrained and the length of tube is measured similar to that for nasogastric gavage. 2. The capsule is placed snugly into the dosing end of the stomach tube and the tip of the other end of the tube is cut off so that a 10- to 20-ml syringe can be attached. 3. A thin film of vegetable oil is placed on the capsule. The vegetable oil facilitates passage of the tube while protecting the capsule from degradation by enzymes in the animal’s saliva. Water® soluble lubricants such as KY Jelly (Johnson & Johnson Co., Arlington, TX) are unacceptable because they will degrade gelatin capsules. 4. With one hand the technician holds the animal’s head in either a normal postural position or in extension, while with the other hand the technician places a speculum into the animal’s mouth. 5. A second technician inserts the end of the stomach tube with the capsule into the animal’s mouth and over the speculum. The tube is advanced until the appropriate distance, as previously determined, is reached. 6. The presence of the capsule in the end of the tube precludes verification of proper tube placement prior to administration as described for nasogastric and orogastric gavage of liquids. 7. A 10- to 20-ml syringe filled with air is attached to the end of the stomach tube and the capsule is dislodged by depressing the syringe plunger. © 2002 CRC Press LLC

8. Proper placement of the tube can be confirmed at the time the capsule is administered by auscultating the animal’s upper-left abdominal quadrant as the plunger is depressed. A “pop” will be heard as the capsule is dislodged from the tube by the air. 9. The tube is removed as described for nasogastric gavage. 10. The stomach tube is discarded after gavage and the mouth speculum is thoroughly disinfected prior to use in another animal.

miscellaneous procedures Disarming Canine Teeth To minimize potential physical harm to both personnel and animals, it is often necessary to disarm the canine teeth of male nonhuman primates. Several canine disarming techniques have been described in nonhuman primates, including extraction of canines,238–240 crown reduction followed by a mucoperiosteal flap,241 crown reduction fol242 lowed by a root canal procedure, and crown reduction followed by 243 Crown reduction followed by pulpal a pulpal capping procedure. capping has many advantages over the other techniques because it is less invasive, quicker to perform, requires less technical expertise, and is associated with minimal post-procedural care and complications. Additional information on canine disarming can be found in the veterinary dental literature.244 Equipment needed to disarm canine teeth using a crown reduction and a pulpal capping technique includes: • Mouth gag or dental bite block • Cotton pellets or paper points • Front surface mirror, Williams perio-probe, and self-locking forceps • Double-ended amalgam carrier, amalgam plugger, and amalgam well • Mixing spatula and mixing pad • Restorative liners (such as calcium hydroxide and varnish) • Restorative material (such as amalgam) • An amalgam shaker • Diamond burrs, including a tapered, cylindrical, and inverted © 2002 CRC Press LLC

cone (size of burrs depends on the size of the animal) • A dental unit with a high-speed, water-cooled hand piece, and a 3-way syringe for irrigation and air drying Procedure 243, 244 1. Appropriate protective clothing should be worn during dental procedures. For macaques, this includes a Tyvek® gown/coveralls, two pairs of gloves, goggles, face shield, and a NIOSH 95 particulate respirator. A PAPR (Powered Air Purifying Respirator) could be used in lieu of the goggles, face shield, and respirator. 2. The animal is appropriately anesthetized. 3. The crown of the canine is amputated with a tapered diamond burr at the level of the occlusal surface of the incisors and premolars (Figure 5.29). 4. The viability of the pulp is assessed. The presence of blood or pink coloration within the pulp indicates a viable pulp and the pulpal capping procedure can proceed. The absence of blood/pink coloration and/or the presence of necrotic tissue indicates the pulp is dead and a root canal procedure must be

Fig. 5.29 Amputation of the canine crown at the level of the premolar and incisor occlusal surface.

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Fig. 5.30 Cross-sectional (A) and sagittal (B) views of the cavity created in the pulpal chamber. Sagittal view (C) of the restorative materials placed over the exposed pulp of the canine tooth.

performed. 5. If the pulp is viable, an access cavity is created within the pulpal chamber, approximately 4 to 5 mm in depth and 2 to 3 mm in diameter, using a cylindrical diamond burr (Figure 5.30 A & B). 6. Bleeding from the pulp is controlled by lavaging with a 1:40 Nolvasan/sterile water solution, pressure with cotton pellets and paper points, and gentle air-drying. In addition, rinsing the pulpal access cavity with a solution of 2% lidocaine and 1:100,000 epinephrine can be used to control bleeding. 7. After the bleeding is stopped, the pulpal access cavity is airdried and packed with calcium hydroxide paste. 8. The calcium hydroxide is allowed to dry and the excess is removed from the cavity using an inverted cone burr. A 1- to 2-mm base of calcium hydroxide is left over the previously exposed pulp. In addition to removing the excess pulp, the inverted cone diamond burr is used to undercut the pulpal access cavity, thus creating a retention shoulder for the amalgam. 9. Water and air are used to clean the cavity, and a drop of varnish © 2002 CRC Press LLC

(a sealant) is placed on the remaining calcium hydroxide. The varnish is gently dried with air. 10. Amalgam is packed in small increments into the cavity using an amalgam carrier and plugger until the amalgam is even with the cut surface of the tooth. The plugger is used to burnish (smooth) the surface of the amalgam (Figure 5.30C). 11. A cylindrical or taper diamond burr is used to round off any sharp angles on the cut surface of the tooth. 12. Post-procedure, animals are administered analgesics and fed a soft diet for 2 days.

Bimanual Rectal Palpation35 Bimanual rectal palpation has been used to detect early pregnancy in macaques and baboons, although ultrasonography has supplanted the use of this technique at many facilities. This technique requires well-trained experienced staff to interpret the findings. Procedure 1. The animal is appropriately sedated and placed in either lateral or ventral recumbency. 2. Wearing appropriate personal protective equipment, the examiner inserts a lubricated middle finger into the rectum of the animal. The finger is inserted as far as possible and then pressed ventral or toward the abdominal wall. 3. With the other hand, the examiner pushes the caudal abdomen toward the tip of the finger inserted into the rectum (Figure 5.31). 4. The examiner then palpates the uterus with the fingertip. A pregnant uterus is soft or fluctuant upon palpation, whereas a nonpregnant uterus is turgid. 5. For cynomolgus or smaller species, the little finger may have to be used instead of the middle finger.

Necropsy Necropsy is defined as the postmortem examination of a body, including the internal organs and structures after dissection, so as to determine the cause of death or the natural pathologic changes.245 In the biomedical research environment, necropsies are performed at © 2002 CRC Press LLC

Fig. 5.31 Sagittal representation of bimanual rectal palpation technique for the detection of pregnancy.

the conclusion of specific research projects and for diagnostic purposes. Necropsy of nonhuman primates should be performed by a veterinarian or under the direct supervision of a veterinarian. There are inherent zoonotic risks associated with the necropsy of a nonhuman primate. These risks include exposure to tissues, body fluids, aerosols, and sharp contaminated instruments that might harbor zoonotic agents such as B virus or M. tuberculosis. For this reason, personnel should be properly trained in how to perform a necropsy and the potential health risks associated with performing a necropsy on a nonhuman primate. Necropsy of nonhuman primates should be performed in a room solely dedicated to postmortem examination and on a surface that readily allows the drainage of fluids and sanitization. The management practices, safety equipment, and facilities of a necropsy room for nonhuman primates should meet, at a minimum, the CDC criteria for a Biosafety Level 2 Laboratory.80, 82 The use of down-draft © 2002 CRC Press LLC

tables and ventilated workstations significantly reduces potential aerosol exposure and should be considered an essential component of a necropsy room. Appropriate personal protective equipment should be worn when performing a necropsy on a nonhuman primate. This should include a gown or coverall impervious to fluids, gloves, face mask, and appropriate protective eye wear. Studies involving certain specific infectious agents may require that additional precautionary measures be taken, and double-gloving should be considered when performing a necropsy on a macaque. Nonhuman primates should be necropsied as soon after death as possible. The carcass should be refrigerated if a period of time will lapse between death and the postmortem examination. The freezing of a carcass is generally not acceptable because it can cause significant postmortem changes. Most necropsies will result in the collection of tissues for microscopic examination and the agent most frequently used to preserve tissues is a 10% neutral-buffered formalin solution. To ensure appropriate tissue fixation, specimens should be 0.5- to 1.0-cm thick and placed into a specimen container containing a volume of formalin ten times the total volume of tissue. Formalin is considered a mucous membrane irritant and a carcinogen; therefore, special precautionary measures should be taken when handling this fixative. Specimen containers should be filled in a fumehood and these containers should remain covered at all times except when tissues are being placed in them. Rubber or nitrile gloves should be used when working with formalin. A consistent, systematic approach should be used when performing a necropsy on a nonhuman primate. The use of specific forms, records, and/or standard operating procedures ensure that staff conduct a necropsy in a uniform manner. A complete guide to the necropsy of large animals has been published elsewhere and can be used as a template in developing necropsy procedures for nonhuman primates.246 The basic equipment needed for a proper postmortem examination of a nonhuman primate includes: • Appropriate personal protective equipment • A small metric ruler • Toothed and serrated tissue forceps • Scalpel blades and handles, and/or knife

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• Shears, scissors (heavy and fine), and hemostats • Rib cutters, bone cutting saw, and/or bone rongeur • A dissecting board • An assortment of syringes (3, 5, 10, and 20 ml) and needles (18 to 23-gauge) • Sterile swabs for bacteriologic culture • Fixative, saline, specimen containers, tissue cassettes, and silk ties Procedure 1. An appropriate necropsy report form is used to record findings. 2. The nonhuman primate’s identification number is checked, the animal is weighed, and the data are recorded. 3. An external examination of the body is performed, including an assessment of the animal’s body condition, skin, lymph nodes, eyes, ears, nose, mouth, and anal and urogenital openings. 4. The animal is placed in dorsal recumbency. A scalpel blade or knife is used to incise the skin and muscles of the inguinal and axillary regions, and the arms and legs are reflected so that the animal lies squarely on its back. 5. A scalpel is used to incise the skin along the ventral midline from the chin to the pubis. A scalpel, knife, or tissue shears is used to reflect the skin laterally away from the underlying subcutaneous tissues and musculature. 6. The abdominal wall is lifted up with forceps and a stab incision is carefully made with a scalpel along the ventral midline. Using heavy scissors or shears, the incision is extended caudally to the pubis and cranially to the rib cage. To allow for greater exposure of the peritoneal cavity, the abdominal muscles along the caudal aspect of the rib cage are also cut. 7. The abdominal organs and the peritoneal surfaces are examined for any abnormalities and the findings recorded. Fluid present in the abdominal cavity in excessive amounts or with abnormal color are sampled for cytology and bacteriologic culture, and the volume and appearance of the fluid are recorded. 8. The intestines are lifted toward the prosector and the mesenteric attachments are cut, thereby freeing the intestinal tract from the abdominal cavity. An intestinal forceps is placed at the © 2002 CRC Press LLC

stomach/esophagus junction, and the stomach/esophagus junction is cut so that the forceps remains attached to the stomach. This prevents leakage of gastrointestinal fluids. 9. The gastrointestinal tract is removed from the abdominal cavity and placed to the side of the carcass to facilitate a thorough examination of the liver, gall bladder, adrenal glands, abdominal aorta, vena cava, and the organs of the urogenital system. The spleen and pancreas usually remain attached to the gastrointestinal tract. 10. To prevent contamination of instruments and tissues, the gastrointestinal tract is examined and cut open at the conclusion of the necropsy. 11. The salivary glands, mandibular lymph nodes, larynx, and trachea are exposed by lateral reflection of the skin and muscles of the neck. Care is taken not to damage the thyroid and parathyroid glands. 12. The thoracic cavity is exposed by incising the diaphragm and removing the ventral portion of the rib cage. This is done by using rib cutters and cutting along the costochondral junction on both sides of the sternum. The ribs can be cut from either the thoracic inlet to the caudal aspect of the thorax, or from the caudal aspect of the thorax to the thoracic inlet. 13. The thoracic organs and the pleural surfaces are thoroughly examined for any abnormalities and the findings are recorded. Fluid present in the thoracic cavity in excessive amounts or with abnormal color is sampled for cytology and bacteriologic culture, and the volume and the appearance of the fluid is recorded. 14. To facilitate further inspection, the thoracic viscera, along with the tongue, oropharynx, larynx, trachea, esophagus thyroid, and parathyroids, are removed en-bloc. This is done by incising the muscles along the ventral medial aspect of the mandible until the tongue is free. The tongue is pulled through the ventral opening between the two mandibles, and gentle traction is applied as the larynx, trachea, and esophagus are dissected free of underlying attachments. The dissection is continued into the thoracic cavity by elevating the trachea so that the attachments to the heart and lungs can be severed. 15. To access the brain, the head is removed from the neck by cutting through the atlanto-occipital joint with a scalpel or knife. © 2002 CRC Press LLC

The head is placed on its ventral surface and a midline skin incision is made over the cranium. The skin and muscles over the cranium are reflected laterally. Using a bone cutting saw, cuts are carefully made through the rostral, lateral, and caudal limits of the cranium. A bone rongeur can also be used to access the brain after a window is cut in the cranium using a bone cutting saw. 16. During each step of the necropsy, tissues are collected as specified in the research protocol or as determined necessary for diagnostic purposes. Tissues are gently rinsed with saline prior to immersion into the respective tissue fixative. In cases where small organs (e.g., thyroid) must be separated from larger organs, cassettes are used to prevent tissue loss. 17. At the conclusion of the necropsy, the carcass is placed into a plastic bag (two bags are preferable), and all instruments and soiled surfaces are thoroughly cleaned. The bagged carcass is placed in a freezer until it can be disposed of in a manner consistent with institutional policy and/or local and state regulations.

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6 resources Examples of vendors and organizations are included in this chapter to provide the user of this handbook with information regarding pertinent organizations, books, periodicals, and electronic resources, as well as sources of nonhuman primates, equipment, and materials. The lists are not exhaustive, nor do they imply endorsement of one vendor over others, but rather are meant to be used as a starting point for developing a database of resources.

organizations Many professional organizations exist that can serve as initial contacts for obtaining information regarding specific professional issues related to the care and use of nonhuman primates. Membership in these organizations allows the laboratory animal science professional to stay abreast of regulatory issues, improvements in methodology and procedures, management issues, and animal health issues. Relevant organizations include the following: • American Association for Laboratory Animal Science (AALAS): 9190 Crestwyn Hills Drive, Memphis, TN 38125 (Telephone: 901-754-8620; Web site: http://www.aalas.org/). AALAS serves a diverse professional group, including principal investigators, animal care technicians, and veterinarians. The journals Comparative Medicine and Contemporary Topics in Laboratory Animal Science and the quarterly newsletter, Tech Talk, are published by AALAS and serve to communicate relevant information to the investigative community. AALAS sponsors a program for

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certification of laboratory animal science professionals at three levels: Assistant Laboratory Animal Technician (ALAT), Laboratory Animal Technician (LAT), and Laboratory Animal Technologist (LATG). The AALAS-affiliated Institute for Laboratory Animal Management (ILAM) is a program designed to provide training in laboratory animal facility management. AALAS maintains a Web site with extensive links to a variety of subject material related to the field of laboratory animal science, including links to pertinent regulatory agencies and organizations. In addition, the association sponsors an electronic bulletin board for technicians (Tech Talk Online) and hosts COMPMED (Comparative Medicine Discussion List). AALAS also sponsors an annual meeting and there are local groups associated with AALAS known as branches. • American College of Laboratory Animal Medicine (ACLAM): ACLAM is an organization of laboratory animal veterinarians founded to encourage education, training, and research in laboratory animal medicine. ACLAM is recognized as a specialty of veterinary medicine by the AVMA and certifies veterinarians as Diplomates in laboratory animal medicine by means of examination. The group sponsors the annual ACLAM Forum as well as sessions at the annual AALAS and American Veterinary Medical Association (AVMA) meetings. ACLAM also sponsors the publication of texts containing detailed information on species used in biomedical research. Two of those texts, Nonhuman Primates in Biomedical Research: Biology and Management and Nonhuman Primates in Biomedical Research: Diseases, are excellent reference texts for individuals working with nonhuman primates. In addition, ACLAM has sponsored the development of a series of autotutorials on laboratory animals for training and education, including a series on nonhuman primates. These autotutorials are also available on CD-ROM. Current contact information can be obtained by accessing their Web site at http://www.aclam.org. • American Society of Laboratory Animal Practitioners (ASLAP): ASLAP Coordinator, 11300 Rockville Pike, Suite 1211, Rockville, MD 20852 (Tel: 301-231-6349; Web site: http://www.aslap.org). ASLAP is an association of veterinarians engaged in some aspect of laboratory animal medicine. The society promotes the acquisition and dissemination of knowledge and information among veterinarians and veterinary students having an interest in laboratory animal practice. The society publishes a newsletter (Laboratory Animal Practitioner) and sponsors a biennial meeting © 2002 CRC Press LLC

in conjunction with the annual AALAS meeting as well as sessions at the annual AALAS and AVMA meetings. • American Society of Primatologists (ASP): ASP is an education and scientific organization whose purpose is to promote and encourage the discovery and exchange of information on a variety of subjects regarding nonhuman primates. ASP members receive the ASP Bulletin, which is published quarterly and contains information on the activity of the organization and other items of interest concerning nonhuman primates. The organization sponsors an annual meeting and members have the opportunity to subscribe to the American Journal of Primatology at a reduced rate. The contact person for ASP changes annually with the election of officers. Current contact information can be obtained by accessing their Web site at http://www.asp.org. • Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC): Executive Director, 11300 Rockville Pike, Suite 1211, Rockville, MD 20852-3035 (Telephone: 301-231-5353; Web site: http//www.aaalac.org). AAALAC is a nonprofit organization that provides a mechanism for peer evaluation of laboratory animal care programs. Since its inception in 1965, AAALAC accreditation has become widely accepted as strong evidence of a quality research animal care and use program. The accreditation process includes an extensive internal review conducted by the institution applying for accreditation as well as a comprehensive assessment of the institution’s animal care and use program by AAALAC using the Guide for the Care and Use of Laboratory Animals as a guideline. • Association of Primate Veterinarians (APV): 624 Stone Road, Harleysville, PA 19438 (Tel: 215-256-8511; Web site: http//www. primate.wisc.edu/pin/idp/idp/entry/429). APV is an association of veterinarians whose purpose is to promote the dissemination of information related to the health, care, and welfare of nonhuman primates. The organization sponsors an annual workshop and publishes a quarterly newsletter. • Institute of Laboratory Animal Research (ILAR): NAS 347, 2101 Constitution Avenue NW, Washington, D.C. 20418 (Telephone: 202-334-2590; Web site: http://www4.nas.edu/cls/ ilarhome.nsf/web/homepage). ILAR functions under the auspices of the National Research Council to develop and make available scientific and technical information on laboratory animals © 2002 CRC Press LLC

and other biologic resources. A number of useful publications are available from ILAR, including the Guide for the Care and Use of Laboratory Animals, the ILAR Journal, and The Psychological WellBeing of Nonhuman Primates. • International Council for Laboratory Animal Science (ICLAS): ICLAS is an international scientific organization dedicated to advancing human and animal health by promoting the ethical care and use of laboratory animals in research. A primary aim of the organization is to promote and coordinate the development of laboratory animal science throughout the world, including international collaborations of laboratory scientists, humane animal care and use of research animals, and the monitoring of quality in research animals worldwide. The organization is composed of national, scientific, and scientific union members. The contact person for ICLAS changes annually with the election of officers. Current contact information can be obtained by accessing their Web site at http://www.iclas.org. • International Primatological Society (IPS): IPS is an association of scientists who do research on nonhuman primates. The society holds international meetings that alternate between nonhuman primate habitat and non-habitat countries. Meetings are held every 2 years during even-numbered years. The society also publishes a newsletter and the International Journal of Primatology. The contact person for IPS changes annually with the election of officers. Current contact information can be obtained by accessing their Web site at http://www.primate.wisc.edu/pin/ ips.html. • Laboratory Animal Management Association (LAMA): P.O. Box 877, Killingworth, CT 06419 (Web site: http://www.lamaonline.org/). LAMA is an organization dedicated to the exchange of information between individuals with management responsibilities for laboratory animal facilities. The group publishes the LAMA Review and sponsors an annual meeting and sessions at the annual AALAS meeting.

publications There are a number of published materials, including both books and periodicals, that contain information pertinent to the management, health and well-being of nonhuman primates. © 2002 CRC Press LLC

Books 1. Anesthesia and Analgesia in Laboratory Animals, edited by D. F. Kohn, S. K. Wixson, W. J. White, and G. J. Benson, 1997. Academic Press, Inc., 525 B Street, Suite 1900, San Diego, CA 92101-4495 (Tel: 1-800-321-5068; Web site: http://www.academicpress.com). 2. Biosafety in Microbiological and Biomedical Laboratories, 4th edition, CDC-NIH (Centers for Disease Control and Prevention — National Institutes of Health), 1999. HHS Publication No. (CDC) 93-8395, U.S. Government Printing Office, Washington, D.C. 20402 (Tel: 202-257-3318; Web site: http://www.nih.gov/od/ ors/ds/pubs/bmbl/index.htm; Stock Number: 017-040400547-4). This book can be read online for free. 3. The Care and Feeding of an IACUC, edited by M. L. Podolsky and V. S. Lukas, 1999. CRC Press, 2000 NW Corporate Blvd., Boca Raton, FL 33431-9868 (Tel: 1-800-272-7737; Web site: http://www.crcpress.com). 4. Chimpanzees in Research, NRC (National Research Council) Committee on Long-term Care of Chimpanzees, 1997. National Academy Press, 2101 Constitution Avenue NW, Lockbox 285, Washington, D.C. 20055 (Tel: 1-888-624-8373; Web site: http://www.nap.edu). This book can be read online for free. 5. Formulary for Laboratory Animals, 2nd edition, by C. T. Hawk and S. L. Leary, 1999. Iowa State University Press, 2121 S. State Avenue, Ames, IA 50014-8300 (Tel: 1-800-862-6657; Web site: http://www.isupress.edu). 6. Guide for the Care and Use of Laboratory Animals, 7th edition, NRC (National Research Council) Institute of Laboratory Animal Resources Committee to Revise the Guide for the Care and Use of Laboratory Animals, 1996. National Academy Press, 2101 Constitution Avenue NW, Lockbox 285, Washington, D.C. 20055 (Tel: 1-888-624-8373; Web site: http://www.nap.edu). 7. The IACUC Handbook, edited by J. Silverman, M. A. Suckow, and M. Sreekant, 2000. CRC Press, 2000 NW Corporate Blvd., Boca Raton, FL 33431-9868 (Tel: 1-800-272-7737; Web site: http://www.crcpress.com). 8. Laboratory Animal Anaesthesia, 2nd edition, by P. A. Flecknell, 1996. Academic Press, Inc., 525 B Street, Suite 1900, San Diego, CA 92101-4495 (Tel: 1-800-321-5068; Web site: http://www.academicpress.com). © 2002 CRC Press LLC

9. Nonhuman Primates I: Monographs on Pathology of Laboratory Animals, edited by T. C. Jones, U. Mohr, and R. D. Hunt, 1993. Springer-Verlag New York, Inc., 175 5th Avenue, New York, NY 10010 (Tel: 1-800-777-4643; Web site: http://www.springerverlag.com). 10. Nonhuman Primates II: Monographs on Pathology of Laboratory Animals, edited by T. C. Jones, U. Mohr, and R. D. Hunt, 1993. Springer-Verlag New York, Inc., 175 5th Avenue, New York, NY 10010 (Tel: 1-800-777-4643; Web site: http://www.springerverlag.com). 11. Nonhuman Primates in Biomedical Research: Biology and Management, edited by B. T. Bennett, C. R. Abee, and R. Henrickson, 1995. Academic Press, Inc., 525 B Street, Suite 1900, San Diego, CA 92101-4495 (Tel: 1-800-321-5068; Web site: http://www.academicpress.com). 12. Nonhuman Primates in Biomedical Research: Diseases, edited by B. T. Bennett, C. R. Abee, and R. Henrickson, 1998. Academic Press, Inc., 525 B Street, Suite 1900, San Diego, CA 921014495 (Tel: 1-800-321-5068; Web site: http://www.academicpress.com). 13. Occupational Health and Safety in the Care and Use of Research Animals, NRC (National Research Council) Committee on Occupational Safety and Health in Research Animal Facilities, 1997. National Academy Press, 2101 Constitution Avenue NW, Lockbox 285, Washington, D.C. 20055 (Tel: 1-888-624-8373; Web site: http://www.nap.edu). This book can be read online for free. 14. The Psychological Well-Being of Nonhuman Primates, NRC (National Research Council) Committee on Well-Being of Nonhuman Primates, 1998. National Academy Press, 2101 Constitution Avenue NW, Lockbox 285, Washington, D.C. 20055 (Tel: 1-888-624-8373; Web site: http://www.nap.edu). 15. The Veterinary Clinics of North America: Exotic Pet Medicine Volume II, edited by K. E. Quesenberry and E. V. Hillyer, 1993. W.B. Saunders Co., 6277 Sea Harbor Drive, Orlando, FL 32887-4800 (Tel: 1-800-654-2452; Web site: http://customerservice.wbsaunders.com).

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Periodicals 1. American Journal of Primatology. Published by Wiley-Liss, Inc., 605 Third Avenue, New York, NY 10158-0012 (Tel: 212-8506645; Web site: http://www.wiley.com/). 2. Comparative Medicine. Published by the American Association for Laboratory Animal Science. For contact information, see above listing for AALAS. Most recent issues are available online. 3. Contemporary Topics in Laboratory Animal Science. Published by the American Association for Laboratory Animal Science. For contact information, see above listing for AALAS. Most recent issues are available online. 4. ILAR Journal. Published by the Institute of Laboratory Animal Research, National Research Council. For contact information, see above listing for ILAR. 5. Journal of Medical Primatology. Published by Munksgaard International Publishers Ltd., Commerce Place, 350 Main Street, Malden, MA 02148-0518 (Tel: 781-388-8273; Web site: http://www.munksgaard.dk). 6. Laboratory Animals. Published by the Royal Society of Medicine Press, 1 Wimpole Street, London W1G OAE, UK (Tel: 02072902921; Web site: http://www.roysocmed.ac.uk/pub/ rspress.htm). 7. Lab Animal. Published by Nature Publishing Co., 345 Park Avenue South, New York, NY 10010-1707 (Tel: 212-726-9332; Web site: http://www.labanimal.com). 8. Laboratory Primate Newsletter. Edited by Judith E. Schrier, Psychology Department, Box 1853, Brown University, Providence, RI 29012 (Tel: 401-863-2511). Current issues are available at http://www.brown.edu/Research/Primate.

electronic resources Many online sources of information relevant to the care and use of laboratory animals, including nonhuman primates, are available. The list below is not meant to be all inclusive; however, it is meant to provide the user with a starting point from which one can readily obtain information as well as explore the Internet as it relates to the care and management of nonhuman primates. © 2002 CRC Press LLC

1. Comparative Medicine Discussion List (COMPMED). COMPMED is an electronic mailing list available through the Internet; it is designed to provide users with a means to quickly tap into the expertise of laboratory animal science professionals around the world. List membership is restricted. At the time of publication, those interested in using this resource should subscribe by sending an e-mail message to:[email protected] and in the body of the message, enter “sub COMPMED first name last name”. Additional information about COMPMED can be obtained at http://www.aalas.org/. 2. Infant Primate Research Laboratory: Research Protocol & Technician’s Manual. A Guide to the Care, Feeding and Evaluation of Infant Monkeys. This is an online manual written by staff at the Washington Regional Primate Research Center regarding the care and husbandry of infant nonhuman primates. This manual can be accessed at http://www.rprc. washington.edu/iprl/contents.htm. 3. Network of Animal Health (NOAH). NOAH is a commercial online service sponsored by the American Veterinary Medical Association (AVMA). NOAH was designed to connect veterinarians to colleagues, board-certified specialists, and a variety of online-interactive veterinary professional services. A number of forums cover a variety of topics, some of which would be of interest to those charged with the care and use of nonhuman primates. Additional information can be obtained from the AVMA (1931 N. Meacham Road, Suite 100, Schaumburg, IL 60173 (Tel: 1-800-248-2862; Web site: http://www.avma.org/ noah/noahlog.asp). 4. NetVet Veterinary Resources (NetVet) and Electronic Zoo. These Web sites contain comprehensive directories related to veterinary medicine and animal resources available on the Internet as well as links to these directories. NetVet and Electronic Zoo are licensed by the AVMA and are ideal for veterinarians, researchers, technicians, and students seeking information on animals, including nonhuman primates. Access to the NetVet and Electronic Zoo Web sites are, respectively: http://www.avma.org/netvet/vet.htm and http://netvet.wustl.edu/e-zoo.htm. 5. Primate Info Net (PIN). The PIN was developed as an Internet information resource for people with an interest in the field of © 2002 CRC Press LLC

primatology and is maintained and managed by the Wisconsin Regional Primate Research Center (WRPRC) Library and Information Service located at the University of Wisconsin–Madison. PIN contains details on the many additional Internet-based programs available through the WRPRC Library, including: AskPrimate, an e-mail-based reference service for questions pertaining to primates, primate organizations, or individuals in primatology; Audiovisual Archives, an electronic method to access the WRPRC Library’s nonhuman primate audiovisual archives; International Directory of Primatology, a directory of informational resources pertaining to nonhuman primates; Primate-Jobs, an international job listing service for people interested in working with nonhuman primates; World Directory of Primatologists, an Internet source of contact information for people working in the field of primatology. The PIN can be accessed at http://www.primate.wisc.edu/pin. 6. Working Safely with Nonhuman Primates (Video). This video was developed in 1999 by the NIH Office of Animal Care and Use. It provides an overview of nonhuman primate behavior and the proper use of personal protective equipment and may be viewed for free at http://www.grants.nih.gov/grants/olaw/primatevideo.cfm, or it can be purchased by contacting the Office of Laboratory Animal Welfare, NIH, RKL1, Suite 1050, MSC 7982, 6705 Rockledge Drive, Bethesda, MD 20892-7982.

primate sources Nonhuman primates can be obtained from a variety of commercial and noncommercial domestic and nondomestic sources, including primary importers, domestic breeding colonies, academic institutions, and industrial corporations. No matter what the source, animals should only be obtained from facilities that have in place a welldefined preventive health program, and are in good standing with the appropriate federal agencies and accrediting organizations. In addition to the commercial sources listed below, primate centers (see Primate Research Centers) and the Primate Supply Information Clearinghouse (PSIC) can serve as invaluable resources for obtaining nonhuman primates, tissues, and other biological samples. The PSIC was established with the purpose of providing information to the © 2002 CRC Press LLC

research community for the efficient sharing of laboratory primates by research institutions, including the sale, exchange, and/or transfer of animals between institutions. The PSIC publishes a bimonthly newsletter with listings of available nonhuman primates. For more information on the PSIC contact: Washington Regional Primate Research Center, Box 357330, University of Washington, Seattle, WA 98195-7730 (Tel: 206-543-5178; Web site: http://www.rprc.washington.edu/psic/).

Possible Commercial Sources of Nonhuman Primates Nonhuman Primate Species African greens Baboons (olive) Cebus Cynomolgus Marmosets Owl monkeys Rhesus Squirrel monkeys

Commercial Source 1, 8, 9, 10 1, 7, 8, 10 1, 7, 10 1, 2, 3, 5, 6, 7, 9, 10 1, 4, 7, 10 10 1, 3, 5, 6, 9, 10 1, 2, 7, 9, 10

Contact Information for Nonhuman Primate Sources 1. Buckshire Corporation, P.O. Box 155, 2025 Ridge Road, Perkasie, PA 18944 (Tel: 215-257-0116; Web site: http://www.buckshire-corp.com). 2. Charles River BRF, Inc., 305 Almeda-Genoa Road, Houston, TX 77047 (Tel: 713-433-5846; Website: http://www.criver.com). 3. Covance Research Products, Inc., P.O. Box 7200, Denver, PA 17517 (Tel: 1-800-345-4114; Web site: http://www.covance.com). 4. Highwater Farms, P.O. Box 97, S.R. #1403, Kipling, NC 27543 (Tel: 919-639-6458; e-mail: [email protected]). 5. LABS of Virginia, P.O. Box 557, 95 Caselle Hall Road, Yemassee, SC 29945 (Tel: 803-589-5190). 6. Oriental Scientific Instruments I/E Group, No. 52 Sanlihe Road, Beijing 100864 China (Tel: 86-10-6872-6607; Web site: http://www.osic.com.cn/eindex.html). 7. Osage Research Primates, 54 Hospital Drive, Osage Beach, MO 65065 (Tel: 573-348-8002; e-mail: [email protected]). 8. Paradise Exports, Box 7522, Arusha, Tanzania (e-mail: [email protected]; fax (UK): 44-870-134-9481).

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9. Primate Products, Inc., 7780 NW 53rd St., Miami, FL 33166 (Tel: 305-471-9557; Web site: http://www.primateproducts.com). 10. Three Springs Scientific, Inc., 1730 W. Rock Road, Perkasie, PA 18944 (Tel: 215-257-6055; e-mail: [email protected]).

nonhuman primate transportation resources Below is a list of organizations that can provide useful information pertaining to the transportation of animals, including nonhuman primates. 1. Animal Transportation Association (AATA), U.S. Business Office, 5521 Greenville Avenue, Suite 104-310, Dallas TX (Tel: 903-769-2207). 2. International Air Transport Association (IATA), 800 Place Victoria, P.O. Box 113, Montreal, Quebec, Canada H4Z 1M1 (Tel: 514-874-0202). Publication: IATA Live Animal Regulations.

nonhuman primate transportation services Most, if not all, commercial sources of nonhuman primates will provide transportation service to your facility. This service may be provided by either air or ground transportation carriers. In the event that a research institution is selling, buying, transferring, or exchanging nonhuman primates with another research institution, one may have to make transportation arrangements. Some airlines will transport nonhuman primates and the cargo department of the respective airlines servicing your area should be contacted to obtain more information. When using an air carrier, every effort should be made to use direct flights between destinations to avoid the problems associated with flight changes and layovers. Moreover, additional arrangements will need to be made to get the animals to and from the airport. For door-to-door delivery, ground transportation services should be considered. Below is a list of companies that specialize in both the domestic and international transportation of nonhuman primates: 1. Frames Animal Transportation Service, 1119 Haverford Road, Ridley Park, PA 19078 (Tel: 610-521-1123). Specializes in the domestic transportation of nonhuman primates. 2. Highwater Farms, P.O. Box 97, Kipling, NC 27543 (Tel: 919639-6458). Specializes in the domestic transportation of nonhuman primates. © 2002 CRC Press LLC

3. International Animal Exchange, Inc., 130 E. Nine Mile Road, Ferndale, MI 48220 (Tel: 248-398-6533). Specializes in the domestic and international transportation of nonhuman primates. 4. Kritter Krates, 5533-A Avanak, Spring, TX 77389 (Tel: 281-2880040). Specializes in the domestic and international transportation of nonhuman primates. 5. Primate Products, Inc., 7780 NW 53rd St., Miami, FL 33166 (Tel: 305-471-9557; Web site: http://www.primateproducts.com/). Specializes in the domestic and international transportation of nonhuman primates.

laboratory services The following laboratories specialize in DNA analysis, parentage analysis, and genetic profiles. 1. Genetics Laboratory for Typing Nonhuman Primates, Trinity University, 715 Stadium Drive, San Antonio, TX 78212-7200 (Tel: 210-999-8347; Web site: http://www.trinity.edu/departments/biology/gentics/). 2. Charles River Therion, Inc., 185 Jordan Road, Troy, NY 121807617 (Tel: 518-286-0016; Web site: http://www.theriondna.com). The following laboratories specialize in the serologic health assessment of nonhuman primates. In addition, these laboratories will perform viral isolation of certain specific viral agents indigenous to nonhuman primates. 1. NIH B Virus Resource Laboratory, Viral Immunology Center, Georgia State University, 50 Decatur Street, Atlanta, GA 30303 (Tel: 404-651-0808; Web site: http://www.gsu.edu/bvirus). 2. BioReliance, 14920 Broschart Road, Rockville, MD 20850-3349 (Tel: 301-738-1000; Web site: http://www.bioreliance.com). 3. Simian Retrovirus Laboratory, California Regional Primate Research Center, Road 98 at Hutchinson, University of California, Davis, CA 95616 (Tel: 530-752-5696; Web site: http://www.srl.ucdavis.edu/). 4. Virus Reference Laboratory, South Texas Medical Center, Suite 205, 7540 Louis Pasteur, San Antonio, TX 78229 (Tel: 210-6147350). © 2002 CRC Press LLC

The following laboratory performs adjunctive tests to assist in the diagnosis of tuberculosis, including tuberculosis culture/isolation, polymerase chain reaction, an enzyme linked immunosorbent assay, and an in vitro blood-based assay of cell mediated immunity that detects the presence of interferon-gamma. 1. United States Department of Agriculture, Animal and Plant Health Inspection Service, National Veterinary Services Laboratories, P.O. Box 844, 1800 Dayton Avenue, Ames, IA 50010 (Tel: 515-6637301; Website: http://www.aphis.usda.gov/vs/nvsl/index.html).

feed The dietary considerations of nonhuman primates can vary significantly between species (i.e., New World vs. Old World primates and leaf eaters vs. non-leaf eaters). Care should be taken to make sure the diet’s nutritional composition meets the dietary needs of the respective species. Below is a list of companies that specialize in the production of diets for nonhuman primates. 1. Bio-Serv, Inc., 1 Eighth Street, Suite 1, Frenchtown, NJ 08825 (Tel: 908-996-2155; Web site: http://www.bio-serv.com). Produces custom diets, palatable medicated treats, and a variety of enrichment food items for nonhuman primates, including marmosets. 2. Harlan Teklad, Inc., P.O. Box 44220, Madison, WI 53744-4220 (Tel: 1-800-483-5523 or 608-277-2070; Web site: http://www.harlan.com). Produces New and Old World primate diets, and custom diets. 3. PMI/Purnia Mills, Inc., P.O. Box 66812, St. Louis, MO 631666812 (Tel: 1-800-227-8941; Website: http://www.labdiet.com or http://www.mazuri.com). Produces New (including marmoset) and Old World primate diets, custom diets, and a diet designed to reduce dental calculus buildup in Old World primates. 4. Zeigler Bros. Inc., P.O. Box 95, Gardners, PA 17324 (Tel: 717677-6181 or 1-800-841-6800; Web site: http://www.zeiglerfeed.com). Produces New World primate and marmoset diets.

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equipment Sanitation Several sources of disinfectants and other sanitation supplies are listed below. 1. BioSentry, Inc., 1481 Rock Mountain Blvd., Stone Mountain, GA 30083-9986 (Tel: 1-800-788-4246; Web site: http://www.biosentry.com). 2. Pharmacal Research Labs, P.O. Box 369, 33 Great Hill Road, Naugatuck, CT 06770-0369 (Tel: 1-800-243-5350; Web site: http://www.pharmacal.com). 3. Rochester Midland, Corp., 333 Hollenbeck St., P.O. Box 31515, Rochester, NY 14603-1515 (Tel: 1-800-836-1627; Web site: http:// www.rochestermidland.com). 4. Steris Corporation, 5960 Heisley Road, Mentor, OH 44060-1834 (Tel: 1-800-548-4873; Web site: http://www.steris.com).

Cages, and Research and Veterinary Supplies Several sources for pharmaceuticals, hypodermic needles, syringes, surgical equipment, bandages, and other related items are provided below. Pharmaceuticals should be ordered and used only under the direction of a licensed veterinarian. Cages should meet the size requirements as specified by relevant regulatory agencies. Stainless steel is preferable to galvanized steel.

Possible Sources of Cages, and Research and Veterinary Supplies Item Cages and supplies Veterinary and surgical supplies Gas anesthesia equipment Restraint chairs Restraint equipmenta Tether systems (jackets and swivels) Enrichment devices/treats Capture systems (blow dart) Syringes and needles Vascular access equipment Osmotic pumps Necropsy tools a

Leather gloves, nets, pole and collars, boxes. © 2002 CRC Press LLC

Source 5, 9, 10, 19, 20, 25, 27 11, 12, 14, 15, 17, 22, 30 14, 17, 22, 28, 29, 30 14, 24, 25 7, 21, 25 3, 21 7, 18, 23, 25 2, 31 4, 11, 13, 15, 17, 26 1, 8, 14, 16, 21 6 4, 13, 30

Contact Information for Cages, and Research and Veterinary Supplies 1. Access Technologies, Division of Norfolk Medical, 7350 N. Ridgeway, Skokie, IL 60076 (Tel: 877-674-7131; Web site: http://www.norfolkaccess.com). 2. Addison Biological Laboratory Inc., 507 North Cleveland Avenue, Fayette, MO 65248 (Tel: 1-800-331-2530; Web site: http://www.addisonlabs.com). 3. Alice King Chatham Medical Arts, 11915 Inglewood Avenue, Hawthorne, CA 90250 (Tel: 310-970-1063). 4. Allegiance Health Care Corp., 1450 Waukegan Road, McGaw Park, IL 60085-9988 (Tel: 1-800-964-5227; Web site: http://www.allegiance.net/). 5. Allentown Caging Equipment, Inc., P.O. Box 698, Allentown, NJ 08501-0698 (Tel: 609-259-7951 or 1-800-762-2243; Web site: http://www.acecaging.com). 6. Alza Corporation, 1900 Charleston Road, P.O. Box 7210, Mountain View, CA 94039-7210 (Tel: 650-564-5000; Web site: http://www.alza.com). 7. Bio-Serv, Inc., 1 Eighth Street, Suite 1, Frenchtown, NJ 08825 (Tel: 1-908-996-2155; Web site: http://www.bio-serv.com). 8. Braintree Scientific, Inc., P.O. Box 850929, Braintree, MA 021850929 (Tel: 718-843-2202; Web site: www.braintreesci.com). 9. Britz-Heidbrink, Inc., P.O. Box 1179, Wheatland, WY 822011179 (Tel: 1-800-808-5609; Web site: http://www.cagesbh.com). 10. Bryan Research Equipment Company, P.O. Box 4232, Bryan, TX 77803 (Tel: 1-800-822-5609). 11. Butler Co., Inc., 5000 Bradenton Avenue, P.O. Box 7153, Dublin, OH 43017 (Tel: 1-800-848-5983; Web site: http://www.wabutler.com). 12. Burns Veterinary Supply, 1900 Diplomat Drive, Farmers Branch, TX 75234 (Tel: 1-800-922-8767; Web site: http://www.burnsvet.com). 13. Fisher Scientific, Inc., 711 Forbes Avenue, Pittsburgh, PA 15219-4785 (Tel: 1-800-766-7000; Web site: http://www3.fishersci.com). 14. Harvard Apparatus, Inc., 84 October Hill Road, Holliston, MA 01746 (Tel: 1-800-272-2775). © 2002 CRC Press LLC

15. Henry Schein, 135 Duryea Road, Melville, NY 11747 (Tel: 1-800666-8100; Web site: http://www.henryschein.com). 16. Instech Solomon Laboratories, Inc., 5209 Militia Hill Road, Plymouth Meeting, PA 19462 (Tel: 215-256-1839; Web site: www.solsci.com). 17. J.A. Webster, Inc., 86 Leominster Road, Sterling, MA 01564 (Tel: 1-800-225-7911; Web site: http://www.jawebster.com/). 18. K.L.A.S.S., Inc., 7955 San Miguel, Cyn Road, Box 410, Salinas, CA 95118 (Tel: 831-786-0956). 19. Lab Products, Inc., 742 Sussex Ave., P.O. Box 639, Seaford, DE 19973-0639 (Tel: 302-628-4300 or 1-800-526-0469; Web site: http://www.labproductsinc.com). 20. L.G.L. - Animal Care Products, Inc., 1520 Cavitt Street, Bryan, TX 77801 (Tel: 979-775-1776; Web site: http://www.lglacp.com). 21. Lomir Biomedical, Inc., 95 Huot Notre-Dame, Ile Perrot, PQ J7V 7M4 Canada (Tel: 514-425-3604; Web site: http://www.lomir.com). 22. NLS Animal Health, 11407 Cronhill Drive, Owing Mills, MD 21117 (Tel: 1-800-638-8672; Website: http://www.nlsanimalhealth.com). 23. Otto Environmental, 6914 N. 124th St., Milwaukee, WI 53224 (Tel: 414-358-1001; Web site: http://www.ottoenvironmental.com). 24. Plas-Labs, 917 E. Chilson Street, Lansing, MI 48906 (Tel: 517372-7177 or 1-800-866-7527; Web site: http://www.plaslabs.com). 25. Primate Products, Inc., 7780 NW 53rd St., Miami, FL 33166 (Tel: 305-471-9557; Web site: http://www.primateproducts.com). 26. Retractable Technologies, Inc., 622 Mill St., Lewisville, TX 75057 (Tel: 972-221-6644 or 1-888-703-1010; Web site: http://www.vanishpoint.com). 27. Suburban Surgical Company, Inc., 275 12th St., Wheeling, IL 60090 (Tel: 847-537-9320 or 1-800-323-7366 ext. 3484; Web site: http://www.suburban-surgical.com/). 28. SurgiVet/Anesco, N7 W22025 Johnson Road, Suite A, Waukesha, WI 53186 (Tel: 262-513-8500 or 1-888-745-6562; Web site: http://www.anesco-vet.com). 29. Veterinary Equipment Inc., P.O. Box 10785, Pleasanton, CA 94588 (Tel: 925-463-1828 or 1-800-466-6463; Web site: http://www.vetequip.com). © 2002 CRC Press LLC

30. Viking Medical, P.O. Box 2142, Medford Lakes, NJ 08055 (Tel: 609953-0138 or 1-800-920-1033; Web site: http://www.vikingmedical.com). 31. Animal Care Equipment & Services, Inc., P.O. Box 3275, Crestline, CA 92325 (Tel: 1-800-338-2237; Web site: http://www.animal-care.com/).

primate research centers NCRR-Supported Regional Primate Research Centers The National Center for Research Resources (NCRR) supports eight Regional Primate Research Centers (RPRC). These centers, which are affiliated with academic institutions, are located throughout the country to maximize their accessability to as many scientists as possible. The RPRC provide a wide variety of services to the scientific and educational communities. These services include specialized facilities, scientific and technical expertise, an appropriate environment for research on primates, the availability of a wide variety of nonhuman primate species, and access to biological specimens. Below is a list of the eight NCRR supported RPRC, their contact information, areas of research emphasis, and a general list of the primary animals maintained at the respective center.

1. California Regional Primate Research Center Contact information: California Regional Primate Research Center, 1 Shields Avenue, Davis, CA 95616-8542 (Tel: 530-7520447; Web site: http://www.crprc.ucdavis.edu/crprc/homepage.html). Research emphasis: behavioral biology, comparative primate biology, developmental and reproductive biology, virology and immunology, and the effects of environmental pollutants and disease on the pulmonary system. Primary species: macaques.

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2. New England Regional Primate Research Center Contact information: New England Regional Primate Research Center, One Pine Hill Drive, P.O. Box 9102, Southborough, MA 01772-9102 (Tel: 508-624-8002; Web site: http://www.hms.harvard.edu/nerprc/). Research emphasis: infectious diseases with a focus on simian lentivirus-induced disease, immunology, oncogenic herpesviruses, behavioral biology, neurodegenerative diseases, neurochemistry, and neuropharmacology. Primary species: marmosets, macaques, and squirrel monkeys. 3. Oregon Regional Primate Research Center Contact information: Oregon Regional Primate Research Center, 505 NW 185th Avenue, Beaverton, OR 97006-3499, Mail Code: L584 (Tel: 503-645-1141; Web site: http://www.ohsu.edu/orprc). Research emphasis: reproductive biology, neurobiology, immunology, and pathobiology. Primary species: macaques. 4. The Southwest Foundation for Biomedical Research Contact information: The Southwest Foundation for Biomedical Research, P.O. Box 760549, San Antonio, TX 78245-0549 (Tel: 210-258-9400; Web site: http://www.srprc.org). Research emphasis: genetics, virology, reproductive endocrinology and biology, cardiovascular diseases, and infectious diseases with a focus on AIDs and hepatitis. The facility has a biosafety level-4 laboratory for the study of highly contagious and dangerous pathogens. Primary species: baboons, macaques, and chimpanzees. 5. Tulane Regional Primate Research Center Contact information: Tulane Regional Primate Research Center, 18703 Three Rivers Road, Covington, LA 70433 (Tel: 504892-2040; Web site: www.tpc.tulane.edu). Research emphasis: infectious diseases with a focus on AIDS, virology, parasitology, urology, and gene therapy. Primary species: macaques.

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6. Washington Regional Primate Research Center Contact information: Regional Primate Research Center, University of Washington, Health Sciences I-421, Seattle, WA 981957330 (Tel: 206-543-1430; Web site: http://www.rprc.washington.edu). Research emphasis: neurologic sciences, cardiovascular function, disease models, developmental biology, endocrinology and metabolism, AIDS, immunogenetics, and virology. Primary species: macaques and baboons. 7. Wisconsin Regional Primate Research Center Contact information: The Wisconsin Regional Primate Research Center, 1220 Capitol Court, Madison, WI 53715-1299 (Tel: 608-263-3500; Web site: http://www.primate.wisc.edu). Research emphasis: reproductive and developmental biology, ethology, neurobiology, immunology, immunogenetics, virology, psychobiology, and aging, and metabolic diseases. Primary species: macaques and marmosets. 8. Yerkes Regional Primate Research Center Contact information: Yerkes Regional Primate Research Center, Emory University, 954 Gatewood Road, Atlanta, GA 30322 (Tel: 404-727-7732; Web site: http://www.cc.emory.edu/ whsc/yerkes/). Research emphasis: microbiology and immunology with a focus on AIDS, gene therapy, cardiovascular disease, reproductive disorders, neurobiology, psychobiology, and the visual system. Primary species: macaques and chimpanzees.

Other Primate Research Centers 1. Caribbean Primate Research Center Contact information: Caribbean Primate Research Center, University of Puerto Rico, Medical Sciences Campus, P.O. Box 1053, Sabana Seca, PR 00952-1053 (Fax: 787-795-6700; Web site: http://home.coqui.net/caribbeanprc).

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Research emphasis: behavioral, demographic, genetic, functional morphological and spontaneous diseases, and other primarily noninvasive research projects. Primary species: macaques. 2. Coulston Foundation Contact information: Coulston Foundation, White Sands Research Center, 1300 LaVelle Road, Alamogordo, NM 88310 (Tel: 505-434-1725; Web site: http://www.Coulston.org/). Research emphasis: infectious disease and vaccine development with a focus on AIDs and hepatitis, pharmaceutical research, aging, and reproductive biology. Primary species: macaques and chimpanzees. 3. Duke University Primate Center Contact information: The Duke University Primate Center, 3705 Erwin Road, Durham, NC 27705 (Tel: 919-489-3364; Web site: http://www.duke.edu/web/primate). Research emphasis: primate evolution, and prosimian biology, and conservation. Primary species: prosimians. 4. University of Oklahoma Health Sciences Center Contact information: Department of Microbiology and Immunology and Division of Animal Resources, University of Oklahoma Health Sciences Center, 940 S. L. Young Blvd., Oklahoma City, OK 73140 (Tel: 405-271-5185). Research emphasis: reproductive biology and behavior. Primary species: baboons. 5. University of South Alabama Primate Research Laboratory Contact Information: Primate Research Laboratory, Department of Comparative Medicine, University of South Alabama, Mobile, AL 36688 (Tel: 334-460-6239; Web site: http://www.saimiri. usouthal.edu/prl). Research emphasis: reproductive biology and endocrinology, medical primatology, behavior, and genetics. Primary species: squirrel monkeys. © 2002 CRC Press LLC

equivalents and conversions Weight Equivalents 1 1 1 1 1 1 1

lb oz kg gm mg mcg mcg

= 453.6 gm = 16 oz = 28.35 gm = 1000 gm = 2.2 lb = 1000 mg = 1000 µg = 0.001 gm = 0.001 mg = 0.000001 gm per gram or 1 mg per kg is the same as 1 ppm

Volume Equivalents HOUSEHOLD

METRIC

1 drop 15 drops 1 teaspoon (tsp.) 1 tablespoon (tbs.) 2 tablespoons (tbs.) 1 ounce (oz) 1 measuring cup (8 oz)

= = = = = = =

0.06 millimeter (ml) 1 ml 5 ml 15 ml 30 ml 30 ml 240 ml

Metric Apothecary 1 milligram (mg) = 1/60 grain (gr) 15.0 mg = 1/4 gr 30.0 mg = 1/2 gr 40.0 mg = 2/3 gr 50.0 mg = 3/4 gr 60.0 mg = 1 gr 1 gram = 15 gr

Temperature Conversion °Celsius = °Fahrenheit: 9 × °C = (5 × °F) – 160 °Celsius to °Fahrenheit: (°C × 1.8) + 32 = °F °Fahrenheit to °Celsius: (°F - 32) × 0.555 = °C

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references 1. Hill, B. F., The rhesus monkey (M. mulatta): history, management and scientific investigation, Charles River Digest, 16(1), 1, 1977. 2. Squirrel Monkey Breeding and Research Resource, Department of Comparative Medicine, College of Medicine, University of South Alabama, Mobile, AL, http://www.saimiri.usouthal.edu/ 3. Hearn, J. P., The marmoset and tamarin, in The UFAW Handbook on the Care & Management of Laboratory Animals, 6th edition, Poole, T., Ed., Longman Scientific & Technical, Essex, 1987, chap. 37. 4. Schnell, C. R., Haemodynamic measurements by telemetry in conscious unrestrained marmosets: response to social and nonsocial stress events, in Marmosets and Tamarins in Biological and Biomedical Research, Proc. of a workshop organized by the European Marmoset Research Group, Pryce, C., Scott, L., and Schnell, C., Eds., DSSD Imagery, Salisbury, U.K., 1997, 181. 5. American Association for Laboratory Animal Science Assistant Laboratory Animal Technician Training Manual, Lawson, P. T., Ed., Sheridan Books, Inc., Chelsea, MI, 1999, 181. 6. Canadian Council on Animal Care, Guide to the Care and Use of Experimental Animals, Volume 1, Ottawa, Ontario, Canada, 1980, Appendix III. 7. Hom, G. J., Comparison of cardiovascular parameters and/or serum chemistry and hematology profiles in conscious and anesthetized rhesus monkeys (Macaca mulatta), Contemporary Topics, 38, 60, 1999.

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8. Yarbrough, L. W., Serum biochemical, hematological and body measurement data for common marmosets (Callithrix jacchus jacchus), Lab. Anim. Sci., 34, 276, 1984. 9. Mansfield K., Personal communication, 2000. 10. Manning, P. J., Lehner, N. D., Feldner, M. A., and Bullock, B. C., Selected hematologic, serum biochemical, and arterial blood gas characteristics of squirrel monkeys (Saimiri sciureus), Lab. Anim. Sci., 19, 831, 1969. 11. Verlangieri, A. J., Normal serum biochemical, hematological, and EKG parameters in anesthetized adult male Macaca fascicularis and Macaca arctoides, Lab. Anim. Sci., 35, 63, 1985. 12. Hainsey, B. M., Clinical parameters of the normal baboons (Papio species) and Chimpanzees (Pan troglodytes), Lab. Anim. Sci., 43, 236, 1993. 13. McAulty, P. A., The relative merits of the marmoset in toxicological testing, in Marmosets and Tamarins in Biological and Biomedical Research, Proc. of a workshop organised by the European Marmoset Research Group, Pryce, C., Scott, L., and Schnell, C., Eds., DSSD Imagery, Salisbury, U.K., 1997, 185. 14. Clemons. D., Personal communication, 2000. 15. Carrol, R. M. and Feldman, E. B., Lipids and lipoproteins, The Clinical Chemistry of Laboratory Animals,1st edition, Loeb, W. F. and Quimby, F. W., Eds., Pergamon Press, Maxwell House, 1989, chap. 7. 16. Riley, J. H. and Cornelius, L. M., Electrolytes, blood gases and acid-base balance, The Clinical Chemistry of Laboratory Animals, 1st edition, Loeb, W. F. and Quimby, F. W., Eds., Pergamon Press, Maxwell House, 1989, chap. 15. 17. Kessler, M. J., The hemogram, serum biochemistry, and electrolyte profile of aged rhesus monkeys (Macaca mulatta), J. Med. Primatol., 12, 184, 1983. 18. Wolford, S. T., Reference range data base for serum chemistry and hematology values in laboratory animals, J. Toxicol. Environ. Health, 18, 161, 1986. 19. Fernie, S. and Wrenshall, E., Normative hematologic and serum biochemical values for adult and infant rhesus monkeys (Macaca mulatta) in a controlled laboratory environment, J. Toxicol. Environ. Health, 42, 53, 1994.

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20. Buchl, S. J., Hematologic and serum biochemical and electrolyte values in clinically normal domestically bred rhesus monkeys (Macaca mulatta) according to age, sex and gravidity, Lab. Anim. Sci., 47, 528, 1997. 21. Kapeghian, J. C., Changes in selected serum biochemical and EKG values with age in cynomolgus macaques, J. Med. Primatol., 13, 283, 1984. 22. Castro, M. I., Ketamine-HCl as a suitable anesthetic for endocrine, metabolic, and cardiovascular studies in Macaca fascicularis monkeys, Proc. Soc. Exp. Biol. Med., 168, 389, 1981. 23. Kakoma, I., Distribution characteristics and relationships between hematologic variables of healthy Bolivian squirrel monkeys, Lab. Anim. Sci., 37, 352, 1987. 24. Kakoma, I., Correlative clinical biochemistry and hematological profiles of laboratory-bred Bolivian squirrel monkeys (Saimiri sciureus), J. Med. Primatol., 16, 273, 1987. 25. Kakoma, I., Hematologic values of normal Bolivian squirrel monkeys (Saimiri sciureus): a comparison between wild-caught and laboratory-bred male animals, Folia Primatol., 44, 102, 1985. 26. Socha, W. W., Blood groups of apes and monkeys: current status and practical applications, Lab. Anim. Sci., 30, 698, 1980. 27. Socha, W. W., Rowe, A. W., Lenny, L. L., Lasano, S. G., and Moor-Jankowski, J., Transfusion of incompatible blood in rhesus monkeys and baboons, Lab. Anim. Sci., 32, 48, 1982. 28. Yong-Ye, Niekrasz, M., Kehoe, M., Rolf, L. L., Martin, M., Baker, J., Kosanke, S., Romano, E., Zuhdi, N., and Cooper, D. K. C., Cardiac allotransplantation across the ABO-blood group barrier by the neutralization of preformed antibodies: the baboon as a model for the human, Lab. Anim. Sci., 44, 121, 1994. 29. Smith, B. H., Crummett, T. L., and Brandt, K. L., Ages of eruption of primate teeth: a compendium from aging individual and comparing life histories, Yearbook Phys. Anthropol., 37, 177, 1994. 30. Hendrickx, A. G. and Dukelow, R. W., Reproductive biology, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 9.

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31. Williams, L. and Glasgow, M., Squirrel monkey behavior in research, Inst. Lab. Anim. Res. J., 41, 26, 2000. 32. Brady, A. G., Research techniques for the squirrel monkey (Saimiri spp.), Inst. Lab. Anim. Res. J., 41, 10, 2000. 33. Poole, T., Hubrecht, R., and Kirkwood, J. K., Marmosets and tamarins, in The UFAW Handbook on the Care and Management of Laboratory Animals, 7th edition, Poole, T., Ed., Blackwell Science, Oxford, 1999, chap. 35. 34. Mendoza, S. P., Squirrel monkeys, in The UFAW Handbook on the Care and Management of Laboratory Animals, 7th edition, Poole, T., Ed., Blackwell Science, Oxford, 1999, chap. 37. 35. Hendrickx, A. G. and Dukelow, R. W., Breeding, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 14. 36. Baskerville, M., Old world monkeys, in The UFAW Handbook on the Care and Management of Laboratory Animals, 7th edition, Poole, T., Ed., Blackwell Science, Oxford, 1999, chap. 39. 37. Hendrickx, A. G. and Kraemer, D. C., Reproduction, in Embryology of the Baboon, Hendrickx, A. G., Ed., University of Chicago Press, Chicago, 1971, chap. 1. 38. McMahan, C. A., Wigodsky, H. S., and Moore, G. T., Weight of the infant baboon (Papio cynocephalus) from birth to fifteen weeks, Lab. Anim. Sci., 26, 928, 1976. 39. Welshman, M., Breeding macaques in source countries, in The UFAW Handbook on the Care and Management of Laboratory Animals, 7th edition, Poole, T., Ed., Blackwell Science, Oxford, 1999, chap. 40. 40. Turnquist, J. E. and Hong, N., Functional morphology, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 3. 41. Visalberghi, E., Anderson, J. R., and Poole, T., Capuchin monkeys, in The UFAW Handbook on the Care and Management of Laboratory Animals, 7th edition, Poole, T., Ed., Blackwell Science, Oxford, 1999, chap. 38. 42. Stein, F. J., Sex determination in the common marmoset (Callithrix jacchus), Lab. Anim. Sci., 28, 75, 1978.

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177. Morton, W. R., Knitter, G. H., Smith, P. M., Susor, T. G., and Schmitt, K., Alternatives to chronic restraint of nonhuman primates, J. Am. Vet. Med. Assoc., 191, 1282, 1987. 178. Rogers, W. R., Lucas, J. H., Mikiten, B. C., Smith, H. D., and Orr, J. L., Chronically indwelling venous cannula and automatic blood sampling system for use with nonhuman primates exposed to 60 Hz electric and magnetic fields, Bioelectromag. Suppl., 3, 103, 1995. 179. Elvidge, H., Influence of handling and sedation on plasma cortisol in rhesus monkeys (Macaca mulatta), J. Endocrinol., 70, 325, 1976. 180. Rheinhardt, V., Refining the blood collection procedure for macaques, Lab Animal, 25, 32, 1996. 181. Rheinhardt, V., Liss, C., and Stevens, C., Restraint methods of laboratory nonhuman primates: a critical review, Animal Welfare, 4, 221, 1995. 182. National Research Council, Occupational Health and Safety in the Care and Use of Research Animals, National Academy Press, Washington, D.C., 1997, chap. 3. 183. Smith, E., Blowpipe simplifies injection, adds safety, Veterinary Product News, 1993, 35. 184. National Research Council, The Psychological Well-Being of Nonhuman Primates, National Academy Press, Washington, D.C., 1998, chap. 3. 185. Rheinhardt, V. and Rheinhardt, A., Social enhancement for adult nonhuman primates in research laboratories: a review, Lab Animal, 29, 34, 2000. 186. Rheinhardt, V. and Selig, D., Environmental Enhancement for Caged Rhesus Macaques: A Photographic Documentation, Animal Welfare Institute, Washington, D.C., 1998. 187. Rheinhardt, V., Training nonhuman primates to cooperate during handling procedures: a review, Anim. Technol., 44, 55, 1997. 188. Laule, G. E., Keeling, M. E., Alford, P. L., Thurston, R. H., Bloomsmith, M. A., and Beck, T. E., Positive reinforcement techniques and chimpanzees: an innovative training program, in Regional Proceedings American Association of Zoological Parks and Aquariums, Wheeling, WV, 1992, 713. 189. Laule, G. E., Alford, P. L., Thurston, R. H., Bloomsmith, M. A., and Beck, T. E., Training to reliably obtain blood and urine samples from a diabetic chimpanzee (Pan troglodytes), Zoo Biol., 15, 587, 1996. © 2002 CRC Press LLC

190. Dysko, R. L. and Hoskins, D. E., Medical management (Part B) collection of biological samples and therapy administration, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 13. 191. McGuill, M. N. and Rowan, A. N., Biological effects of blood loss: implications for sampling volume and techniques, ILAR News, 31, 5, 1989. 192. Bivin, W. S. and Smith, G. D., Techniques of experimentation, in Laboratory Animal Medicine, Fox, J. G., Cohen B. J., and Loew, F. M., Eds., Academic Press, Orlando, 1984, chap. 19. 193. Stickrod, G. and Pruett, D. K., Multiple cannulation of the primate superficial lateral cocygeal vein, Lab. Anim. Sci., 29, 398, 1979. 194. Goldsteen, D. L. and Hoyt, R. F., Multiple blood sampling sites enhance the use of small nonhuman primates on pharmacokinetic studies (abstract P22), Contemporary Topics, 38, 52, 1999. 195. Crow, S. E. and Walshaw, S. O., Injection techniques, in Manual of Clinical Procedures in the Dog, Cat and Rabbit, 2nd edition, Crow, S. E. and Walshaw, S. O., Eds., Lippincott-Raven, Philadelphia, 1997, chap. 2. 196. Bahnson, R. R., Ballou, B. T., Ernstoff, M. S., Schwentker, and Hakala, T. R., A technique for catheterization and cystoscopic evaluation of cynomolgus monkey urinary bladders, Lab. Anim. Sci., 38, 731, 1988. 197. Schaffer, N. E., Cranfield, M., Fazleabas, A. T., and Jeyendran, R. S., Viable spermatozoa in the bladder after electroejaculation of lion-tailed macaques (Macaca silenus), J. Reprod. Fert., 86, 767, 1989. 198. Keeling, M. E. and Wolf, R. H., Medical management of the rhesus monkey, in The Rhesus Monkey, Borne, G. H., Ed., Academic Press, New York, 1975. 199. Switzer, J. W., A new technique for sampling bone marrow in monkeys, Lab. Anim. Care, 17, 255, 1967. 200. Wisecup, W. G., Hodson, H. H., Jr., Lovett, R. L., Prine, J. R., and Hanley, W. C., Anemia in chimpanzees (Pan troglodytes) resulting from serial collection of blood, Am. J. Vet. Res., 29, 1823, 1968. 201. Hall, A. S. and Knezevic, A. L., Bone marrow sampling in monkeys, J. Am. Vet. Med. Assoc., 147, 1075, 1965. © 2002 CRC Press LLC

202. Porter, E. S., Fitch, C. D., and Dinning, J. S., Vitamin E deficiency in the monkey, further studies of the anemia with emphasis on bone marrow morphology, Blood, 20, 471, 1962. 203. Bistner, S. I. and Ford, R. B., Special diagnostic procedures, in Handbook of Veterinary Procedures & Emergency Treatment, 6th edition, Bistner, S. I. and Ford, R. B., Eds., W.B. Saunders, Philadelphia, 1995, 501. 204. Crow, E. S. and Walshaw, S. O., Bone marrow aspiration and biopsy, in The Manual of Clinical Procedures in the Dog, Cat, & Rabbit, Crow, E. S. and Walshaw, S. O., Eds., Lippincott-Raven, Philadelphia, 1997, chap. 29. 205. Blackwood, L., Bone marrow—aspiration and biopsy, Waltham Focus, 7, 16, 1997. 206. Morrel, J. M., Kuderling, I., and Hodges, J. K., Influences of semen collection method on ejaculate characteristics in the common marmoset, J. Androl., 17, 164, 1996. 207. Yeomans, R. R., Ricker, R. B., Williams, L. E., Sonksen, J., and Abee, C. R., Vibratory stimulation of ejaculation yields increased motile spermatozoa, compared with electroejaculation, in squirrel monkeys (Saimiri boliviensis), Contemporary Topics, 36, 62, 1997. 208. Gould, K. G., Warner, H., and Martin, D. E., Rectal probe electroejaculation of primates, J. Med. Primatol., 7, 213, 1978. 209. Bornman, M. S., vanVuuren, M., Meltzer, D. G. A., van der Merwe, C. A., and Rensburg, S. J., Quality of semen obtained by electroejaculation from chacma baboons, J. Med. Primatol., 17, 57, 1988. 210. Gould, K. G., Techniques and significance of gamete collection and storage in the great apes, J. Med. Primatol., 19, 537, 1990. 211. Gould, K. G. and Mann, D. R., Comparison and electrostimulation methods for semen recovery in the rhesus monkey (Macaca mulatta), J. Med. Primatol., 7, 213, 1988. 212. Binor, Z., Rawlins, R. G., Van derVen, H., and Dmowski, W. P., Rhesus monkey sperm penetration into zona-free hamster ova: comparison of preparation and culture conditions, Gamete Res., 19, 91, 1988. 213. Lanzendorf, S. E., Gliessman, P. M., Archibong, A. E., Alexander, M., and Wolf, D. P., Collection and quality of rhesus semen, Mol. Reprod. Dev., 25, 61, 1990. © 2002 CRC Press LLC

214. Sarason, R. L., VanderVoort, C. A., Mader, D. R., and Overstreet, J. W., The use of nonmetal electrodes in electroejaculation of restrained but unanesthetized macaques, J. Med. Primatol., 20, 122, 1991. 215. Tarantal, A. F., Interventional ultrasound in pregnant macaques: embryonic/fetal applications, J. Med. Primatol., 19, 47, 1990. 216. Epstein, M. F. and Chez, R. A., Two operative techniques applied in perinatal research in the rhesus monkey, Lab. Anim. Sci., 26, 456, 1976. 217. Hess, D. L., Matayoshi, K., Baker, C. A., and Hendrickx, A. G., Amniocentesis and antenatal sex determination in the rhesus monkey (Macaca mulatta), J. Med. Primatol., 8, 244, 1979. 218. American Association for Laboratory Animal Science Laboratory Animal Technician Training Manual, Lawson, P. T., Ed., Sheridan Books, Inc., Chelsea, MI, 2000, chap. 3. 219. American Association for Laboratory Animal Science Laboratory Animal Technician Training Manual, Lawson, P. T., Ed., Sheridan Books, Inc., Chelsea, MI, 2000, chap. 13. 220. Conti, P. A., Nolan, T. E., and Gehert, J., Immobilization of a chronic intravenous catheter in the saphenous vein of African green and rhesus monkeys, Lab. Anim. Sci., 234, 1979. 221. Bistner, S. I. and Ford, R. B., Therapeutic procedures and techniques, in Handbook of Veterinary Procedures & Emergency Treatment, 6th edition, Bistner, S. I. and Ford, R. B., Eds., W.B. Saunders, Philadelphia, 1995, 536. 222. Crow, S. E. and Walshaw, S. O., Injection techniques, in Manual of Clinical Procedures in the Dog, Cat and Rabbit, 2nd edition, Crow, S. E. and Walshaw, S. O., Eds., Lippincott-Raven, Philadelphia, 1997, chap. 3. 223. Abee, C. R., Medical care and management of the squirrel monkey, in Handbook of Squirrel Monkey Research, Rosenblum, L. A. and Coe, C. L., Eds., Plenum Press, New York, 1985, 447. 224. Dalton, M. J., The vascular port—a subcutaneously implanted drug delivery depot, Lab Animal, 14, 21, 1985. 225. Wojnicki, F. H. E., Bacher, J. D., and Glowa, J. R., Use of subcutaneous vascular access ports in rhesus monkeys, Lab. Anim. Sci., 44, 491, 1994.

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226. Kinsora, J. J., Christoffensen, C. L., Swalec, J. M., and Juneau, P. L., The novel use of vascular access ports for intravenous selfadministration and blood withdrawal studies in squirrel monkeys, J. Neurosci. Meth., 75, 59, 1997. 227. Fitzgerald, A. L., Dillon, L. M., Altroggee, D. M., Bleavins, M. R., and Breider, M. A., Use of subcutaneous vascular access ports in common marmosets (Callithrix jacchus), Contemporary Topics, 35, 57, 1996. 228. Tartarini, K. A., Sils, I. V., and Szlyk-Modrow, P. C., Comparison of blood pressure measurements using vascular access ports and conventional catheters, Contemporary Topics, 35, 57, 1996. 229. Access Technologies, Vascular Access Ports, in Technical Bulletin #PW011199, Access Technologies, Skokie, IL, 1999, 1. 230. Mendenhall, H. V., Piechowiak, M., Wadanoli, M., and Raikowski, D., A long term study on the use of vascular access ports in multiple species, presented at Access Technologies Long-Term Access Roundtable, San Antonio, September 4–6, 1997, 20. 231. Jacobson, A. D., Do not push the V-A-P septum when not accessing, in Vascular Access Catheter Tips, Access Technologies, Skokie, IL, 1996, 2. 232. Stouffer, R. L., Dahl, K. D., Hess, D. L., Woodruff, T. K., Mather, J. P., and Molskness, T. A., Systemic and intraluteal infusion of inhibin A or activin A in rhesus monkeys during the luteal phase of the menstrual cycle, Bio Reprod., 50, 888, 1994. 233. Gordon, K., Williams, R. F., Greer J., Bush, E. N., Haviv, F., Herrin, M., and Hodgen, G. D., A-75998: a fourth generation GnRH antagonist: II preclinical studies in female primates, Endocrine, 2, 1141, 1994. 234. Mufson, E. J., Kroin, J. S., Liu, Y. T., Sobreviela T., Penn, R. D., Miller, J. A., and Kordower, J. H., Intrastriatal and intraventricular infusion of brain-derived neurotrophic factor in the cynomolgus monkey: distribution, retrograde transport and colocalization with substantia nigra dopamine-containing neurons, Neuroscience, 71, 179, 1996. 235. MacDonald, R. L., Weir, B. K. A., Runzer, T. D., Grace, M. G. A., and Pozrasky, M. J., Effect of intrathecal superoxide dismutase and catalase on oxyhemoglobin-induced vasospasm in monkeys, Neurosurgery, 30, 529, 1992. © 2002 CRC Press LLC

236. Fazleabas, A. T., Donnelly, K. M., Sudha, S., Fortman, J. D., and Miller, J. B., Modulation of the baboon (Papio anubis) uterine endometrium by chorionic gonadotropin during the period of uterine receptivity, Proc. Natl. Acad. Sci., 96, 2543, 1999. 237. Halliday, L. C., Fortman, J. D., and Bennett, B. T., A mouth speculum for orogastric administration of compounds to nonhuman primates, Contemporary Topics, 37, 76, 1998. 238. Hilloowala, R. A. and Miller, R. L., Extraction of canine teeth from the rhesus monkey, J. Am. Vet. Med. Assoc., 151, 830, 1967. 239. Gibson, W. E. and Hall, A. S., Surgical removal of the maxillary canine tooth in the rhesus monkey (Macaca mulatta), J. Am. Vet. Med. Assoc., 157, 717, 1970. 240. Smith, A. W., Extraction of baboon canine teeth: a simple efficient technique, Lab. Anim. Sci., 21, 604, 1971. 241. Schofield, J. C., Alves, M. E. A. F., Hughes, K. W., and Bennett, B. T., Disarming canine teeth of nonhuman primates using the submucosal vital root retention technique, Lab. Anim. Sci., 41, 128, 1991. 242. Tomson, R. N., Schulte, J. M. and Bertsch, M. L., Root canal procedure for disarming nonhuman primates, Lab. Anim. Sci., 29, 382, 1979. 243. Coman, J. L., Fortman, J. D., Alves, M. E. A. F., Bunte, R. M., and Bennett, B. T., Assessment of a canine crown reduction technique in nonhuman primates, Lab. Anim. Sci., 37, 67, 1998. 244. Holmstrom, S. E., Frost, P., and Gammon, R. L., Eds., Endodontics, in Veterinary Dental Techniques, W.B. Saunders, Philadelphia, 1992, chap. 7. 245. Friel, J. P., Ed., Dorlands Illustrated Medical Dictionary, 26th edition, W.B. Saunders, Philadelphia, 1981, 142. 246. King, J. M., Dodd, D. C., Roth, L., and Newson, M. E., The Necropsy Book, Charles Louis Davis, D.V.M. Foundation, Gurnee, IL, 1999.

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selected readings Taxonomy, Functional Morphology, Behavior, Anatomic/Physiologic Features 1. Berger, E., Monkeys and Apes, Arco Publishing Co., New York, 1985. 2. Hershkovitz, P., Living New World Monkeys (Platyrrhini) Vol. 1, University of Chicago Press, Chicago, 1977. 3. Kavanagh, M., Monkeys Apes and Other Primates, The Viking Press, New York, 1984. 4. Napier, J. R. and Napier, P. H., Handbook of Living Primates, Academic Press, London, 1967. 5. Napier, J. R. and Napier, P. H., The Natural History of Primates, MIT Press, Cambridge, 1985. 6. McDonald, D., Clutton-Brock, T. H., Matin, B. D., and Mittermeie, R. A., All The Worlds Animals Primates, Torstar Books, Inc., New York, 1985. 7. Rosen, S. I., Introduction to the Primates: Living and Fossil, Prentice-Hall, Englewood Cliffs, NJ, 1974. 8. Whitney, R. A., Taxonomy in nonhuman primates, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 3. 9. Turnquist, J. E. and Hong, N., Functional morphology, in Nonhuman Primates in Biomedical Research, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 4. 10. Williams, L. E. and Berstein, I. S., Study of primate social behavior, in Nonhuman Primates in Biomedical, Biology and Management, Bennett, B. T., Abee, C. R., and Henrickson, R., Eds., Academic Press, San Diego, 1995, chap. 5. © 2002 CRC Press LLC