The Laboratory Nonhuman Primate

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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 (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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43. National Research Council, The Psychological Well-Being of Nonhuman Primates, National Academy Press, Washington, D.C., 1998, chap. 6. 44. Rasmussen, K. M., Ausman, L. M., and Hayes, K. C., Vital statistics from a laboratory breeding colony of squirrel monkeys (Saimiri sciureus), Lab. Anim. Sci., 30, 99, 1980. 45. Taub, D. M., Adams, M. R., and Auerbach, K. G., Reproductive performance in a breeding colony of Brazilian squirrel monkeys (Saimiri sciureus), Lab. Anim. Sci., 28, 562, 1978. 46. Richter, C. B., Lehner, N. D. M., and Henrickson, R. V., Primates, in Laboratory Animal Medicine, Fox, J. G., Cohen B. J., and Loew, F. M., Eds., Academic Press, Orlando, 1984, chap. 11. 47. Schiml, P. A., Mendoza, S. P., Saltzman, W., Lyons, D. M., and Mason, W. A., Seasonality in squirrel monkeys (Saimiri sciureus): social facilitation by females, Physiol. Behav., 60, 1105, 1996. 48. National Research Council, The Psychological Well-Being of Nonhuman Primates, National Academy Press, Washington, D.C., 1998, chap. 8. 49. Else, J. G., Tarara, R., Suleman, M. A., and Eley, R. M., Enclosure design and reproductive success of baboons used for reproductive research in Kenya, Lab. Anim. Sci., 36, 168, 1986. 50. Werner, R. M., Montry, R. D., Roberts, C. R., Tsoy, A. C., and Huxsoll, D. L., Establishment of a cynomolgus monkey (Macaca fascicularis) breeding colony in Malaysia: a feasibility study, Lab. Anim. Sci., 30, 571, 1980. 51. Gardin, J. F., Jerome, C. P., Jayo, M. J., and Weaver, D. S., Maternal factors affecting reproduction in a breeding colony of cynomolgus macaques (Macaca fascicularis), Lab. Anim. Sci., 39, 205, 1989. 52. Riddle, K. E., Keeling, M. E., Alford, P. L., and Beck, T. F., Chimpanzee holding, rehabilitation and breeding: facilities design and colony management, Lab. Anim. Sci., 32, 525, 1982. 53. Herring, J. M., Fortman, J. D., Anderson, R. J., and Bennett, B. T., Ultrasonic determination of fetal parameters in baboons (Papio anubis), Lab. Anim. Sci., 41, 602, 1991. 54. Committee on the Care and Use of Laboratory Animals of the Institute of Laboratory Animal Resources, Guide for the Care and Use of Laboratory Animals, National Academy Press, Washington, D.C., 1996. © 2002 CRC Press LLC

55. Committee on Well-Being of Nonhuman Primates of the Institute of Laboratory Animal Research, The Psychological Well-Being of Nonhuman Primates, National Academy Press, Washington, D.C., 1998. 56. Bielitzki, J., Susor, T. G., Elias, K., and Bowden, D. M., Improved cage design for single housing of social nonhuman primates, Lab. Anim. Sci., 40, 428, 1990. 57. Halliday, L. C., Fortman, J. D., Nelson, M. C., Bartholomew, A. M., Hoffman, R., and Bennett, B. T., A modified cage to minimize catheter contamination in the chronically catheterized baboon, Contemporary Topics, 38(4), 16, 1999. 58. Kelley, S. T. and Hall, A. S., Housing, 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. 10. 59. Reinhardt, V. and Roberts, A., Effective feeding enrichment for nonhuman primates: a brief review, Animal Welfare, 6, 265, 1997. 60. Reinhardt, V., Enticing nonhuman primates to forage for their standard biscuit ration, Zoo Biology, 12, 307, 1993. 61. Byrne, G. D. and Suomi, S. J., Effects of woodchips and buried food on behavior patterns and psychological well-being of captive rhesus monkeys, J. Med. Primatol., 17, 257, 1991. 62. Reinhardt, V., Caged rhesus macaques voluntarily work for ordinary food, Primates, 35, 95, 1994. 63. Eppel, G., Belcher, A., Küderling, I., Zeller, U., Scolnick, L., Greenfield, K., and Smith, A. B. III, Making sense out of scents: species differences in scent-glands, scent-marking behaviour, and scent-mark composition in the Callitrichidae, in Systematics, Behaviour, and Ecology, Rylands, A. B., Ed., Oxford University Press, Oxford, 1993, 123. 64. Animal Welfare Information Center, Environmental Enrichment for Nonhuman Primates Resource Guide, January 1992–February 1999, Kreger, M.D., Ed., Beltsville, MD, 1999. 65. Reinhardt, V. and Reinhardt, A., Environmental enrichment for primates, in Annotated Database on Environmental Enrichment and Refinement of Husbandry for Nonhuman Primates, Animal Welfare Institute, Washington, D.C., 2000.

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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