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6 Veterinary Care Veterinary care in laboratory animal facilities includes monitoring of animal care and welfare, as well as the prevention, diagnosis, treatment, and control of diseases. It entails providing guidance to investigators on han- dling animals and preventing or reducing pain and distress. To perform those and related functions, attending veterinarians must be trained or have experience in the care and management of the species under their care. The responsibilities of an attending veterinarian are specified by the Animal Welfare Regulations (AWRs; 9 CFR 2.33 for research facilities and 9 CFR 2.40 for dealers and exhibitors), the Public Health Service Policy on Hu- mane Care and Use of Laboratory Animals, or PHS Policy (PHS, 1996), and the Guide for the Care and Use of Laboratory Animals, known as the Guide (NRC, 1996 et seq.~. PREVENTIVE MEDICINE Procurement Rodents (excluding mice of the genus Mus and rats of the genus Rattus) that are acquired from outside a research facility's breeding program must be obtained from dealers licensed by the U.S. Department of Agriculture (USDA) or sources that are exempted from licensing (9 CFR 2.11. A1- though laboratory mice and rats are excluded from direct USDA oversight, it is recommended that they be acquired from dealers whose facilities and 85

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86 RODENTS: LABORATORY ANIMAL MANAGEMENT programs conform to the Guide (NRC, 1996 et seq.~. Documentation of animal health status, site visits by users, history of client satisfaction, USDA licensing for production of other rodent species in the same facilities, and accreditation by the American Association for Accreditation of Laboratory Animal Care can be used to assess dealers. Sources Rapid advances in animal-production technology and disease-control methods during the past 20 years have made it easier to obtain laboratory rodents of known health status and genetic definition. Commercial animal producers often maintain colonies of hysterectomy-derived mice, rats, and guinea pigs in barrier facilities designed and operated to prevent the intro- duction of microbial agents. Those producers regularly monitor their colo- nies for evidence of infection and infestation and publish the test results in health reports, which they make available to their clients. There is an increasing trend toward maintaining other rodents (e.g., hamsters and ger- bils) under similar conditions, although usually not produced from hysterec- tomy-derived stock. It is recommended that animals be acquired from such sources whenever it is possible and appropriate for the study. When ani- mals that are not barrier-reared are acquired, precautions should be taken to isolate them until health evaluations are conducted and decisions are made regarding their care and use. Transportation The protection of the health status of specific-pathogen-free (SPF) ro- dents during transportation to the user has improved greatly in recent years. USDA supervision of animal carriers has resulted in important changes, including the requirements that rodents covered by the AWRs not be ware- housed for long periods before and after shipment, that adequate space be provided in shipping enclosures, and that acceptable temperatures and ven- tilation be maintained during all phases of transportation (9 CFR 3.35- 3.411. The International Airline Transport Association (IATA) has devel- oped guidelines for shipping all animal species, including recommendations for shipping rodents (IATA, 1995 et seq.~. Another major improvement has been in the commercial development of disposable shipping containers with filter-protected ventilation openings. In addition, sterile food and moisture sources have become available for use in such containers. Despite the many changes for the better, problems remain. For ex- ample, the potential still exists for contamination of container surfaces dur- ing shipment. It is recommended that the surfaces of shipping containers be decontaminated before the containers are moved into clean areas of animal

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VETERINARY CARE 87 facilities. Several types of disinfectants-including quaternary ammonium solutions, iodinated alcohols, sodium hypochlorite solutions, and chlorine dioxide-containing solutions can be applied with a small hand sprayer. Chlorine-containing solutions are considered to be very effective against stable agents, such as parvoviruses and spore-forming bacteria (Ganaway, 1980; Orcutt and Bhatt, 1986~. The handling of imported rodents on arrival in U.S. airports can also constitute a problem. Laboratory rodents and rodent tissues that are not inoculated with infectious agents do not require a USDA permit; however, U.S. customs inspectors do not always acknowledge this. Unclear lines of authority often cause unnecessary delays in customs clearance, and such delays can have disastrous effects on the health of the animals. To lessen the probability of delays, as much information as possible should be ob- tained from the involved authorities (USDA, U.S. Customs, and U.S. De- partment of the Interior) well in advance of ordering rodents from any foreign source. A permit must also be obtained from the Division of Quar- antine, Centers for Disease Control and Prevention, before rodents that can carry zoonotic agents are imported (42 CFR 1, 71.54~. Sources of informa- tion are listed in the appendix. All necessary documentation should also be obtained before one attempts to export rodents. Specific instructions are usually obtained from the embassy of the country of destination and from the person or institution receiving the animals. Quarantine and Stabilization Ideally, rodents being introduced into an animal facility are isolated until their health status can be determined. The period of quarantine also provides time for physiologic and behavioral stabilization after shipment. The users, in cooperation with the veterinarian, should make decisions about the method and duration of quarantine for different kinds of facilities, stud- ies, and types of animals. Unless it is inconsistent with the goals of the study, animals should be allowed to stabilize before the experiment begins. One of the most common methods of quarantine is to place each group of incoming animals in the same room in which they will eventually be studied. No animals other than those being quarantined should be housed in the quarantine area. For this system to work, each room requires a separate air supply and effective sanitization between studies. Daily animal-care and support activities for quarantine rooms should be conducted after all neces- sary tasks in the nonquarantine rooms have been performed. Another approach is to have a single quarantine room for all incoming shipments of animals. This approach has regained favor since the develop- ment of isolation-type caging systems, which permit true isolation of many small groups of animals in a single room. Filter-top cages, for example, can

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88 RODENTS: LABORATORY ANIMAL MANAGEMENT be used as miniature rooms within a room. This system works well if animals are moved from dirty to clean cages, one cage at a time in a lami- nar-flow hood; soiled cages are then closed and autoclaved before they are emptied outside the hood; and appropriate protocols for handling the cages and animals are followed strictly. An advantage of this system is that investigators trained to use it can enter a room and complete short-term studies while the animals are in quarantine. Other variations of quarantine systems have been described elsewhere (NRC, l991aJ. The extent of testing (e.g., serology and parasitology) that is needed during quarantine depends on professional judgment; however, any rodent that dies or becomes ill during quarantine should be subjected to careful diagnostic evaluation. SPF rodents purchased from an established commer- cial supplier and received in clean, disposable transport cages with filter- protected ventilation openings might not require testing. If the animals are to be used in short-term studies where other short-term studies are per- formed and relatively few animals are at risk, clinical observations and reliance on the supplier's health program might be adequate. Periodic con- firmation of an animal supplier's health report by an independent laboratory provides added safety. If the animals are to be used in a facility where long-term studies might be jeopardized or large numbers of animals are at risk, testing for selected agents of concern is advisable. Maximal protection against the entry of pathogens into a facility is provided by introducing only animals that are delivered by hysterectomy and reared in protective isola- tion until they are old enough to be tested for the presence of undesirable agents (including agents that can inhabit the female reproductive tract), such as Mycoplasma pulmonis, Corynebacterium kutscheri, and Pasteurella pneumotropica. This course of action is usually followed only in long- standing, ordinarily "closed" breeding colonies. Animals of undocumented microbiologic status received from any out- side source should be serologically tested for a comprehensive list of infec- tious agents. Animals from such sources might harbor clinically inapparent infectious diseases of major concern. For example, mousepox can be diffi- cult to detect clinically in resistant strains of mice or in mice from colonies with long-standing infections. When introduced into a disease-free colony, mousepox usually becomes evident as an epizootic that can substantially interfere with research (New, 19811. Laboratory rodents and some wild rodents can be subclinically infected with zoonotic agents- e.g., hantaviruses, lymphocytic choriomeningitis (LCM) virus, Lassa fever virus, Machupo vi- rus, and Junin virus that pose a serious or even deadly health threat to personnel (CDC, 1993; LeDuc et al., 1986; Oldstone, 1987; Skinner and Knight, 1979; Smith et al., 1984~. The time of quarantine should be long enough for reasonable expectation that incubating infections will become evident, either clinically or by appropriate testing procedures. As many as

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VETERINARY CARE 89 30 percent of the animals should be tested if the microbiologic status of the source colony is completely unknown. In this situation, it is preferable to obtain extra animals for testing so that not only serology, but bacterial cultures, examinations for parasites, and histopathologic evaluations can be performed if needed. Some pathogens pose special problems for quarantine programs. For example, the chronic form of LCM viral infection in mice, which is con- tracted in utero or immediately after birth, might not be detectable with antibody tests commonly used in commercial testing laboratories. Mice infected at that time develop persistently high titers of virus that is complexed with humoral antibody, rendering the antibody undetectable by comple- ment-fixation or neutralization tests (Bishop, 1990; Oldstone and Dixon, 1967, 19694. The more-sensitive immunofluorescence assay (IFA) and en- zyme-linked immunosorbent assay (ELISA) give weak reactions and cannot be depended on to detect circulating antibody in persistently infected mice (Parker, 1986; Shek, 1994~. That is an important problem because the primary route of transmission in the mouse is vertical, and the infected offspring become lifelong, relatively asymptomatic shedders of virus (Rawls et al., 1981~. An alternative method for detecting LCM virus in asymptomatic virus shedders is to use virus-free sentinels over the age of weaning (Smith et al., 1984~. Once beyond neonatal age, exposed mice develop a short- lived infection and have readily detectable antibodies to LCM virus (Rawls, 1981~. Intracranial inoculation of blood or tissue homogenates into the sentinels is a faster screening method. If virus is present, necrologic dis- ease and death will ensue in 6-9 days (Parker, 1986~. Additional laboratory procedures would have to be performed to confirm the presence of LCM virus in the dead mice. In testing laboratories that maintain cell lines, such as Vero or BHK-21, the quickest method is to inoculate cell-line cultures with blood from the suspect mice and use the IFA 4-5 days later to test for LCM-virus antigen in the cells. The mouse antibody-production (MAP) test can also be used to detect LCM virus. Antibody to LCM virus in rodents other than persistently infected mice is readily detected with the ELISA or IFA procedures. Viable rodent tissues including blood, ascitic fluid, tissue cultures, transplantable tumors, and hybridomas can harbor undesirable agents, and tissues of undocumented microbiologic status should not be introduced into rodent colonies until they are shown to be free of undesirable agents by diagnostic testing (e.g., MAP testing). Separation by Species, Source, and Health Status Pressures to maintain different rodent species in separate rooms have less- ened with advances in knowledge of rodent infections. For example, the

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9o RODENTS: LABORATORYANIMAL MANAGEMENT AWRs do not require species separation, and the Guide (NRC, 1996 et seq.) allows considerable latitude on this issue. It has become recognized that more infectious agents are transmissible among animals of the same species than among those of different species. A more important concern is the microbio- logic status of rodents from different sources (or from different locations at the same source), regardless of species. Common sense dictates that if it is neces- sary to place rodents from different sources in the same room because of space constraints or for other practical reasons, it should be done only with animals of comparable microbiologic status. Such decisions should be made with input from people knowledgeable in rodent-disease pathogenesis and with ad- equate health-status information about the source colonies. Interspecies anxiety does not appear to be a problem if different rodent species or rodents and rabbits are housed in the same room, although sys- tematic studies are needed to support the validity of this premise. However, it is unacceptable to house rodents with species that are their natural preda- tors, that produce intimidating noises and odors, or that can harbor infec- tious agents of known or unknown consequences in rodents (e.g., cats, dogs, and monkeys). SURVEILLANCE, DIAGNOSIS, TREATMENT, AND CONTROL OF DISEASE Daily Observations of Animals One important way to track the health status of rodent colonies is to observe the appearance and behavior of the animals daily. A wide range of abnormal signs can be detected in this manner, including weight loss, ruffled hair coat, dry skin, lacerations, abnormal gait or posture, head tilt, lethargy, swellings, diarrhea, seizures, discharge from orifices, and dyspnea. Under- lying causes for those signs include such things as malfunctioning watering systems, fighting, infectious diseases, and experimentally induced changes. Observations are usually made by animal-care staff and technicians, who should be trained to look for spontaneous and experimentally induced ab- normalities and report them to the supervisory staff, the attending veterinar- ian, and study directors. Veterinary oversight of this process and training given by the attending veterinarian are important. Veterinary programs for overseeing the health of laboratory rodents should have readily available, up-to-date references on the biology and diseases of rodents. Control of Infectious Diseases First and foremost, control of infectious diseases in rodent colonies means preventing their introduction. That is accomplished by using good

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VETERINARY CARE TABLE 6.1 Typical "Core" Agents Monitored in Research Facilitiesa 91 Agent Mice Rats Guinea Pigs Hamsters Kilham rat virus Minute virus of mice Mouse hepatitis virus Mycoplasma pulmonis Pneumonia virus of mice Rotavirus Sendai virus Sialodacryoadenitis virus (rat coronavirus) Simian virus 5 Theiler's murine encephalomyelitis virus + + + . + ++ + + ++ + + b ~ b b +b a"Core" agents for each species are indicated by plus signs. bInfection with related parainfluenza viruses can cause false-positive results of tests for Sendai virus and simian virus 5 (Parker et al., 1987). management practices, such as purchasing pathogen-free animals; using well- planned quarantine systems for incoming animals and animal-derived speci- mens; training animal-care staff to make accurate clinical observations; us- ing protective clothing; vermin-proofing the facility; using filter-protected cages, filtered-air ventilation systems, or both; and controlling the move- ment of personnel and visitors within the facility. In addition, animal-care staff should be encouraged not to maintain pet rodents, because of the possibility of transferring infectious agents into the animal quarters. Even with good management, infections occasionally gain entrance into colonies. Routine monitoring systems should be in place to detect them as quickly as possible, thereby permitting the start of specific measures to eliminate them or prevent their spread. The key elements of an effective monitoring program are daily observation of the animals to detect clinical diseases and regular microbiologic monitoring to detect subclinical infec- tions. Daily observations are extremely important because they quickly reveal signs of spontaneous disease. To achieve full effectiveness, monitor- ing activities require diagnostic capability to investigate disease outbreaks. Microbiologic monitoring can include many kinds of tests, depending on the needs of the facility. Animal suppliers often test for all infectious agents of rodents for which there are commercially available tests so that fully characterized animals can be offered for research use. In research facilities, the staff might choose to test initially or annually for all known pathogenic agents and test more frequently for a smaller number of "core" agents of special concern. Table 6.1 lists typical "core" agents. The re- search requirements or special interests of the staff will dictate what other agents should be added to the list.

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92 RODENTS: LABORATORY ANIMAL MANAGEMENT Several newly recognized viruses that are not listed as core agents de- serve mention because of their apparent high prevalence. These are the so- called orphan parvoviruses of mice and rats that appear to be widespread in laboratory colonies but are of unknown character and pathogenicity. A1- though field strains of the viruses are yet to be isolated, the mouse orphan parvovirus (MOPV) has been demonstrated in tissues by in situ hybridiza- tion (Smith et al., 1993), and a closely related laboratory strain has been isolated (McKisic et al., 1993~. In routine testing, the viruses of both mice and rats have been detected indirectly by IFA demonstration of antibody against nonstructural proteins of the rodent parvovirus group followed by negative results with hemagglutination inhibition (HAI) tests that are spe- cific for recognized parvoviruses (i.e., MVM, KRV, and Toolan H-1 virus). An HAI test specific for MOPV has been developed by using the laboratory strain (Fitch isolate) but is not yet in general use. It is debatable whether Sendai virus and simian virus 5 (SV5) should continue to be listed as core agents for guinea pigs and hamsters. Although serologic positivity is often found, it is believed by some to be caused by infection with antigenically related parainfluenza viruses, possibly from hu- man sources. Isolation of Sendai virus from guinea pigs has been attempted rarely and described only anecdotallY (Parkers reported bY Van Hoosier and Robinette, 1976~. Failure of transmission of Sendai virus from serologi- cally positive guinea pigs to mice also has been found (W. White, Charles River Laboratories, Wilmington, Massachusetts' unpublished). Isolation of r - , Sendai virus from hamsters has been reported rarely (Parker et al., 19871. Serologic positivity for Sendai and SV5 viruses might be caused by cross reactions with human parainfluenza viruses, but isolation of the human agents from these animals has not been documented. Monitoring can be performed for many combinations of agents and with various frequencies. Emphasis is often on serologic testing because many of the agents of concern cause subclinical infections and are detect- able quickly and inexpensively with this method. Table 6.2 lists infectious agents of commonly used laboratory rodents for which serologic (antibodyJ tests are available. Bacteriologic testing usually entails culturing for primary and opportu- nistic pathogens from the upper respiratory tract and intestines. Table 6.3 , ~ ~ lists the primary pathogens culturable from these sites. Monitoring for ectoparasites is done usually by examining the skin and potage over the head and back with a dissection microscope. For parasites that invade the skin, skin scrapings in immersion oil or 5 percent potassium hy- droxide are examined microscopically. Monitoring for endoparasites is per- formed by using fecal flotation and sedimentation procedures to search for eggs and oocysts, using the Cellophane-tape method to look for Syphacia eggs, examining the cecocolic contents for helminths, and examining the blad

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VETERINARY CARE TABLE 6.2 Infectious Agents of Rodents for Which Serologic Tests Are Available Serologic Test Availablea Agent Clostridium piliforme (formerly called + Bacillus piliformis) Cilia-associated respiratory (CAR) bacillus Ectromelia virus Encephalitozoon cuniculi Hantavirus K virus Kilham rat virus Lymphocytic choriomeningitis virus + Minute virus of mice Mouse adenovirus (MAd-FL, MAd-K87) Mouse cytomegalovirus Mouse hepatitis virus Mouse "orphan" parvovirus Mouse rotavirus Mouse thymic virus Mycoplasma arthritidis Mycoplasma pulmonis Pneumonia virus of mice Polyoma virus Rat coronavirus and sialodacryoadenitis virus + Rat cytomegalovirus Rat "orphan" parvovirus Reovirus 3 Sendai virus Simian virus 5 Theiler's murine encephalomyelitis virus Toolen's H- 1 virus 93 Mice Rats Guinea Pigs Hamsters + + + + + + + + + + + + + + + + + + + + 1 + + + + + + + + + + + + + aAgents for which serologic tests are available are indicated by plus signs. der mucosa for Trichosomoides crassicauda (in rats) and fecal wet smears for protozoa. Descriptions of ectoparasites and endoparasites and their effects on rodents have been published (Farrar et al., 1986; Flynn, 1973; Hsu, 1979, 1982; Ronald and Wagner, 1976; Vetterling, 1976; Wagner, 1987; Wagner et al., 1986; Weisbroth, 1982; Wescott, 1976, 19824. Pathologic monitoring can be used to detect diseases that produce characteristic lesions that are observ- able at necropsy or detectable by histopathologic evaluation. Infectious dis- eases for which this approach is useful include Tyzzer's disease (Clostridium piliforme [formerly called Bacillis piliformis] infection), pneumocystosis (Pneumocystis carinii infection) in some immunodeficient animals, and CAR

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94 RODENTS: LABORATORY ANIMAL MANAGEMENT TABLE 6.3 Important Rodent Bacterial Pathogens Culturable from Upper Respiratory Tract and Intestinesa Agent Mice Rats Guinea Pigs Hamsters Gerbils Bordetella bronchiseptica Campylobacter jejuni Citrobacter freundii (biotype 4280) Corynebacterium hutscheri Helicobacter spp. Mycoplasma pulmonis Salmonella spp. Streptobacillus moniliformis Streptococcus equ is (zoo ep idem icus) Yersinia pseudotuberculosis + + + + + + + + + + + + + + + aCulturable pathogens are indicated by plus signs. Many commonly occulting bacteria can be present as pathogenic strains (e.g., Escherichia cold and Streptococcus pneumonias) or as opportunistic pathogens (e.g., Klebsiella spp., Pasteurella pneumotropica, and Pseudomonas aeruginosa) in stressed or immunoc OCR for page 85
VETERINARY CARE 95 Microbiologic monitoring for evidence of subclinical infections is ac- complished by testing regularly a randomly selected sample of the popula- tion of animals at risk. How to determine the appropriate sample size is a much debated subject. A formula has been used to predict the number of randomly selected animals in a population of 100 or more that must be tested to detect a single case of disease within 95 percent confidence limits, assuming a known prevalence rate (NRC, 19761: log 0.05 No. to be sampled = log N In that formula, N is the percentage of animals expected to be normal. The percentage is derived by subtracting the expected prevalence rate of the disease from 100 percent. The formula is useful for helping to understand -the considerations involved in sampling to detect a single disease. In prac- tice, however, its use is limited by several factors. One factor is that sampling of a rodent population is usually aimed at detecting more than one disease, each with a different expected prevalence. Another problem is that infectious-disease prevalences are affected by population density, caging methods, ventilation systems, and a host of other variables that affect the rate of spread of infections; a disease prevalence expected to be 30 percent in open cages might be only 1 percent in filter-top cages. Still another consideration is that much of the monitoring is done by testing for antibody. If an infection with an expected prevalence of 30 percent has been in a colony for several months, the number of surviving animals with antibody can approach 100 percent. Because of those variables, the formula serves only as a rough estimate. If it is used, one prevalence is selected for all diseases and conditions, even though screening is usually for multiple or- ganisms. For example, a prevalence of 30 percent might be assumed for more contagious infections, and a sample size of 8-10 would be used. This sample size would, of course, be unlikely to detect infections that are less contagious (NRC, l 991 a). Similar calculations can be made for populations of fewer than 100 with other formulas. More complex calculations can be used once the monitoring program is in place and sufficient data have been accrued on the incidence of positive findings and frequency of disease outbreaks. Those calculations can be used to adjust the sample size and frequency of sam- pling to achieve the desired confidence levels for disease detection (Selwyn and Shek, 19941. In summary, there is no easy way to determine sample sizes and fre- quencies for monitoring. Although a mathematical approach can be taken, the inability to conform to the assumptions on which the formulas are based or the lack of precise knowledge of prevalence rates or disease outbreaks

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VETERINARY CARE 103 ket, hot-water bottles, or an incandescent lamp placed 12-14 inches from the animal can be used to supply supplemental heat during the surgical procedure and recovery. Positioning the animal on an insulating surface, such as cloth or paper, will also help to decrease heat loss. The animal should be positioned to provide adequate fixation and expo- sure of the operative site. Tape, positional ties, or similar mechanical means should be used to ensure that the animal's position will not be changed by pressure exerted by the surgeon. Care should be taken so that the selected method of restraint does not impede circulation or cause injury to the ani- mal. Depending on the complexity of the surgical procedure, it might be necessary to place a sterile drape over the animal to prevent contamination of the operative site. Various commercially available cloth, paper, and plastic materials are suitable for use as surgical drapes. In preparation for the procedure, the surgeon should scrub his or her hands and forearms with a povidone iodine scrub, alcohol foam product, or other equally effective disinfectant-detergent. At a minimum, surgical per- sonnel must wear sterile gloves while performing surgery (9 CFR 2.31; NRC, 1996 et seq.~. For rodents other than mice of the genus Mus and rats of the genus Rattus, masks are also required by the AWRs (9 CFR 2.319. Although caps and gowns are not required for rodent surgery, their use can decrease the risk of contaminating the surgical site and sterile supplies. Sterilization of Instruments The AWRs (9 CFR 2.31) and the Guide (NRC, 1996 et seq.) require that all instruments used in survival surgery be sterilized. As many sets of sterilized instruments as possible should be available when a surgical proce- dure will be performed on multiple animals during the same operative pe- riod. If it is necessary to use the same instruments on several animals, instruments that were sterile at the beginning of the procedure should, at a minimum, be disinfected by chemical or other means (e.g., heated glass beads) before they are used on another animal. Various methods and materials are available for sterilization of instru ments and surgical supplies, including heat, steam under pressure, ethylene oxide gas, gamma irradiation, electron-beam sterilization, and such chemical agents as phenols and glutaraldehyde. The method selected should be periodi- cally monitored (e.g., with spore strips in autoclaves) to ensure that steriliza- tion is achieved. When ethylene oxide gas or a liquid chemical agent is used, care should be taken to ensure that all toxic residues are eliminated before the instruments and supplies are used for surgical procedures. Instruments and supplies that are to be sterilized with methods other than contact with liquid agents should be wrapped in paper, cloth, plastic, or similar materials in such a way as to prevent contamination after steril

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104 RODENTS: LABORATORY ANIMAL MANAGEMENT ization. The choice of material should be appropriate for the method of sterilization. Each package should bear some indication that it has under- gone sterilization. The package should also be marked with the date of sterilization. The shelf-life of sterilized items will depend on the type of material used to wrap them and on how they are stored (Berg and Blass, 1985; Gurevich, 1991; Knecht et al., 1981~. Items that are sterilized with liquid agents are generally prepared near the operating room or area and used immediately after they are removed from the liquid and rinsed with sterile water or sterile irrigation solution. Monitoring During Surgery Surgical procedures should not be initiated until the animal has reached a surgical plane of anesthesia. In most rodents, loss of toe-pinch and pedal reflexes indicates that the plane of anesthesia is adequate for surgery. Guinea pigs, however, can maintain a pedal reflex under anesthesia; for them, the pinna reflex is more appropriate for assessing the plane of anesthesia (C. J. Green, 1982~. The animals should be closely monitored throughout the proce- dure. An animal's status can be determined by monitoring respiration, eyes, and mucous membranes. Slow, labored respiration, loss of reflected eye color in albino animals, and pale or cyanotic mucous membranes are all indicators of compromised cardiovascular and respiratory functions. If resuscitation is necessary, a modified bulb syringe can be fitted over the animal's muzzle and gently pumped to force air into its lungs. A gentle, rhythmic pressure can be applied over the apical area of the thorax to induce cardiac contractions. Doxapram can be used to stimulate respiration (Flecknell, 1987~. The attending veteri- narian can instruct investigators about those and other resuscitative techniques most appropriate for the species and procedures used. Postoperative Care A rodent recovering from surgery should be observed regularly until it is conscious and has regained its righting reflex. It should be housed singly in a cage on absorbent material that minimizes heat loss until it is con- scious. Recovery is facilitated by providing supplemental heat as previ- ously described. Care should be taken to prevent thermal injuries if water bottles, electric heating pads, or heating lamps are used. If necessary, body fluid lost during the surgical procedure should be replaced with subcutaneously or intraperitoneally administered fluids. A decision to administer fluids should be based on the nature and length of the surgical procedure and an estimation of fluid loss. Sterile saline, lactated Ringer's and 5 percent glucose solutions are often used. Guidelines on

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VETERINARY CARE 105 fluid-replacement therapy are available (Cunliffe-Beamer and Les, 1987; Lumb and Jones, 1984~. If recovery takes longer than 30 minutes, the animal's position should be rotated to prevent congestion in dependent organs. If there is concern that its toes will become entangled in sutures or that it will harm the inci- sion or damage the bandage or other protective devices, its toenails should be clipped during the postoperative recovery period. Analgesics should be administered as needed during the postoperative recovery period. Possible side effects and drug interactions should be taken into consideration when specific agents are selected for use (Harkness and Wagner, 1989~. Surgical wounds should be examined daily for dehiscence, drainage, and signs of infection. Appropriate nursing care should be given to prevent drainage from the incision from irritating the surrounding skin. If nonab- sorbable sutures or medical staples are used to close the skin, they should be removed when the incision is adequately healed. EUTHANASIA Euthanasia is the act of producing a painless death. It entails disrupting the transmission of signals from peripheral pain receptors to the central ner- vous system (CNS) and rendering the cerebral cortex, thalamus, and subcorti- cal structures of the CNS nonfunctional. The "endpoint" (the point at which euthanasia will be performed) should be specified in any protocol for a termi- nal study or for a study in which the animals are likely to experience pain and distress that cannot be adequately controlled or prevented with pharmacologic agents, including studies associated with infectious diseases or tumor growth. Each investigator should consult with the attending veterinarian to decide on a humane endpoint that will allow collection of the required data without caus- ing undue pain and distress (Amyx, 1987; Montgomery, 1987~. The technique selected for performing euthanasia on laboratory rodents should be based on a number of factors, including the following: species; animal age and condition; objectives of the study; histologic artifacts and biochemical changes induced by the agent or method selected; number of animals to be euthanatized; available personnel; cost and availability of supplies and equipment; controlled-substance use; and skills of assigned personnel.

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106 RODENTS: LABORATORY ANIMAL MANAGEMENT To avoid causing stress in the animals that will be euthanatized, the following principles should be adhered to: Animals should not be euthanatized in the same room in which other animals are being held. The visual, acoustic, and olfactory stimulants that can be present at euthanasia can cause distress in other animals. Animals should be handled gently and humanely during transport from the holding room and during the actual euthanasia process. If a euthanasia chamber is used, overcrowding should be avoided. Euthanasia should be performed only by people trained in the method selected. It is important that the training received include basic information on how the technique works to produce a quick and painless death and on the advantages of using a specific method in a specific protocol. . Counseling should be available for those performing euthanasia to help them understand feelings and reactions that might develop as a result of performing this task. . Death should be verified at the end of the procedure. Possible methods might include exsanguination, decapitation, creation of a pneumothorax by performing a bilateral thoracotomy or incising the diaphragm, and a physical examination to verify the absence of vital signs. PHS Policy (PHS, 1996) requires that methods of euthanasia be consistent with the recommendations of the American Veterinary Medical Association (AVMA) Panel on Euthanasia (AVMA, 1993 et seq.J. AVMA-recommended methods cause death by direct or indirect hypoxia, direct depression of CNS neurons, or physical damage to brain tissues. The approved pharmacologic agents and physical methods include barbiturates, inhalant anesthetics, carbon dioxide, carbon monoxide, nitrogen, argon, and microwave irradiation. Two additional techniques, cervical dislocation and decapitation, can be used if scientifically justified and approved by the IACUC (AVMA, 1993~. Of these agents and methods, four are commonly used for rodents: carbon dioxide, sodium pentobarbital, cervical dislocation, and decapitation. Carbon dioxide is a very safe and inexpensive agent for euthanatizing laboratory rodents. In all but neonates, it causes rapid, painless death by a combination of CNS depression, which is produced by a fall in the pH of the cerebrospinal fluid, and hypoxia. Other methods of euthanasia can be used in newborn animals, which are more resistant to acute respiratory acidosis and hypoxia than older animals. Commercially available cylinders of compressed carbon dioxide or blocks of dry ice can used as the source of carbon dioxide. Compressed gas is preferable because inflow to the cham- ber can be regulated precisely (AVMA, 1993J. If dry ice is used, it should be placed in the bottom of the chamber and separated from the rodent by a barrier to prevent direct contact that could cause chilling or freezing and associated stress.

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VETERINARY CARE 107 Sodium pentobarbital is the barbiturate drug most commonly used for euthanatizing animals and can be administered to rodents either intraperito- neally or intravenously. When administered intravenously to rodents at a dose of 150-200 mg/kg of body weight (NRC, 1992), it causes rapid death by CNS depression and hypoxia. Intracardiac and intrapulmonary routes of administration can cause pain and distress because of the required methods of restraint and other procedural difficulties. Therefore, those routes of administration should not be used unless the animal is anesthetized. Cervical dislocation is an acceptable method for euthanatizing rodents, provided that it is performed by appropriately trained personnel. Death is instantaneous and is caused by physical damage that occurs as the brain and spinal cord are manually separated by anteriorly directed pressure applied to the base of the skull. This technique might be more difficult to perform in~hamsters, rats, and guinea pigs than in other rodents because of the strong muscles and loose skin of the neck region. If the method is selected, it should be remembered that it can produce pulmonary artifacts- blood in the alveoli and vascular congestion (Feldman and Gupta, 19761. For decapitation, only a sharp, clean guillotine or large shears should be used to ensure a clean cut on the first attempt. It is also essential that the cut be made between the atlanto-occipital joint to ensure that all afferent nerves are severed (NRC' 1992~. Decapitation is more difficult in hamsters, rats, and guinea pigs than in other rodents because of the strong muscles and loose skin of the neck region. There has been considerable controversy about how rapidly unconsciousness occurs when this method is used and whether animals should be anesthetized before they are decapitated. There is evidence that unconsciousness occurs very rapidly (in less than 2.7 sec- onds) after decapitation (Alfred and Berntson, 1986-; Derr, 1991~. Recent studies have shown that anesthesia can cause substantial alterations in arachi- donic acid metabolism; lymphocyte assays; and plasma concentrations of glucose, triglycerides, and insulin (Bhathena, 1992; Butler et al., 1990; Howard et al., 19901. It can be concluded that in some cases anesthesia can inter- fere with the interpretation of data obtained from postmortem tissue samples and that appropriately trained personnel can perform decapitation humanely in rodents without anesthesia. REFERENCES ACLAD (American Committee on Laboratory Animal Disease). 1991. Detection methods for the identification of rodent viral and mycoplasmal infections. Special topic issue, G. Lussier, ed. Lab. Anim. Sci. 41:199-225. Aguila, H. N., S. P. Pakes, W. C. Lai, and Y. S. Lu. 1988. The effects of transportation stress on splenic natural killer cell activity in C57BL/6J mice. Lab. Anim. Sci. 38(2):148-151. Allred, J. B., and G. G. Berntson. 1986. Is euthanasia of rats by decapitation inhumane? J. Nutr. 116:1859-1861.

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