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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
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
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
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
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
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
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.
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
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
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<?mpromised animals, or as agents of importance when transmitted from a carrier to a susceptible animal host (e.g., Bordetella bronchiseptica). bacillus infections. Special stains are required to demonstrate those causative agents (e.g., methenamine silver for P. carinii and Warthin Starry silver for C. piliforme and CAR bacillus). Pathologic monitoring can also be used to detect noninfectious conditions, such as nutritional deficiencies, heritable metabolic diseases, and neoplasms. The necropsy is usually the first step in the diagnos- tic workup of clinical diseases, often providing the impetus for using other measures, such as virus isolation, bacterial cultures, or histopathology. Com- plete descriptions of these procedures and the manifestation of infections in rodents are beyond the scope of this report, but such information is available in a number of books, manuals, and review articles (ACLAD, 1991; Baker et al., 1979; Bhatt et al., 1986; Flynn, 1973; Foster et al., 1982; Hamm, 1986; NRC, 1991a; Van Hoosier and McPherson, 1987; Waggle et al., 1994; Wagner and Manning, 19761. Sample Size for Monitoring All animals should be monitored for clinical disease by daily observa- tions. This type of monitoring, combined with a diagnostic workup of animals with unexplained abnormalities, is particularly important for early detection of clinical disease outbreaks. It is complementary to microbiologic monitoring in that diseases that spread slowly and smolder for a considerable time in a few cages in a room (Bhatt and Jacoby, 1987; Wallace et al., 1981) might be missed in the statistical sampling used in microbiologic monitoring. Daily observations should quickly reveal these kinds of diseases.
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
96 RODENTS: LABORATORY ANIMAL MANAGEMENT makes such an approach difficult to apply. For that reason, it is still com- mon to choose sample size and frequency of monitoring in an arbitrary manner, which is often influenced by economic constraints. An alternative method of monitoring uses known pathogen-free sentinel animals to detect infections. Typically, they are randomly dispersed in multiple locations in the facility, and various means are used to promote contagion of any infections that might be present from the animals being monitored by the sentinels. The most effective method is to place the sentinels in the cages with the study animals and move them to cages of different study animals every 1-2 weeks. If such a procedure is not practi- cal, the sentinels should at least be caged on the same rack with the study animals, preferably on a lower shelf, and soiled bedding from the cages of the study animals should be transferred regularly to the cages of the sentinel animals (Thigpen et al., 1989~. Because natural transmission of some pathogens might not occur quickly, the time allowed for seroconversion or production of disease should be about 6-8 weeks. Those pathogens include Myco- plasma pulmonis (Cassell et al., 1986; Ganaway et al., 1973), ectromelia virus (Wallace et al., 1981), and cilia-associated respiratory (CAR) bacillus (Matsushita et al., 1989~; a preferable alternative is to test the animals being introduced into the colony rather than the sentinels. Treatment and Control Health-monitoring data should be reviewed regularly, and a plan of action should be in place for dealing with positive test results. Such plans usually include the names and telephone numbers of research and veterinary staff to be notified, a system for confirming the test results, and appropriate measures for controlling or eliminating infection. Decisions about ways to prevent spread to contiguous areas should be made quickly. They usually involve placing the room under strict quarantine and developing strategies for control- ling access and for handling potentially contaminated items, such as cages and bedding, that will be removed from the room periodically. Investigations are usually initiated immediately to identify the sources of causative agents. Ap- proaches to control depend on the characteristics of the agents, the value of the infected animals, and the type and design of the facility. Bacterial diseases of rodents can be treated with antibiotics. However, when large numbers of animals are involved, this is often considered practi- cal only for temporary control. Failure to eliminate the agent from every animal, as well as from contaminated surfaces, might result in re-emergence of the disease when antibiotics are discontinued. In some instances, antibi- otics can adversely affect rodents, especially guinea pigs and hamsters, by causing an imbalance of the intestinal microflora and overgrowth of del eterious bacteria (Fekety et al., 1979; Small, 1968; Wagner, 1976J. Other
VETERINARY CARE 97 problems include the lack of information on proper dosages, the difficulty of accurately administering antibiotics in food and water, and confounding influences of drug residues and interactions on research results. Parasitic diseases can also be treated; however, even with highly effective antiparasitic drugs, it is very difficult to eliminate from large colonies such parasites as pinworms and mites. It might be possible in small colonies if the treatment schedule is adjusted to overlap the time of the parasite life cycle and if sanitation procedures are stringently performed simultaneously (e.g., fre- quent washing of floors, walls, and cages) (Findon and Miller, 1987; Flynn et al., 1989; Silverman et al., 1983; Taylor, 1992; West et al., 1992~. Viral, bacterial, and parasitic infections are usually eliminated by eu- thanatizing and repopulating the colony with disease-free animals after the room, cages, and other equipment have been decontaminated or, in the case of particular viruses, by allowing the infection to run its course in a closed population to produce noninfected, immune survivors. The latter procedure has been used successfully with such viruses as Sendai virus and mouse hepatitis virus, which are highly contagious, usually remain in the animals for a short time, and are relatively unstable in the animal-room environment (Barthold. 1986: Fuiiwara and Wanner, 19861. For it to be successful, , ,, ~ . . ~ . r . · · · 1 _ 1 _ ~1 ~ ample opportunity for contagion IS required, and new animals, even new- borns, must not be introduced for a period long enough for all animals to become infected, recover, and stop shedding the virus. Contagion can be promoted by transferring infected bedding to numerous cages, placing cage racks near each other, and removing filter tops. Sentinels can be introduced and tested 6-8 weeks later to determine the success of the procedure. No sentinels should be introduced into the room, and no naive animals of any type should be allowed to be introduced or maintained in the room until 6-8 weeks after breeding has been stopped. Necropsies When an animal is unexpectedly found dead or moribund, it is good practice to determine the cause by necropsy. Necropsy, coupled with daily observations by the animal technicians, usually provides the first indication of important clinical infectious and noninfectious diseases. Lesions will often be characteristic enough to permit presumptive diagnoses or point to appropriate additional diagnostic procedures. Routine histopathologic tests are performed in some facilities. EMERGENCY, WEEKEND, AND HOLIDAY CARE The need for adequate animal care does not diminish during holidays and weekends. As stated in the Guide, laboratory animals should be cared
98 RODENTS: LABORATORY ANIMAL MANAGEMENT for daily (NRC, 1996 et seq.) Security personnel should be able to contact responsible people in the event of emergencies. Therefore, a list of names and phone numbers should be posted prominently in the facility and main- tained in the security office. Provisions for emergency veterinary care should be made as well (9 CFR 2.33b2; NRC, 1996 et sequin MINIMIZATION OF PAIN AND DISTRESS Many internal and external environmental factors can induce physi- ologic or behavioral changes in laboratory animals. These factors are called stressors, and their effect is called stress (NRC, 1992~. The intensity of the stress experienced by an animal is influenced by other factors, including age, sex, genetics, previous exposure, health status, nutrition, and medica- tion (Blass and Fitzgerald, 1988; NRC, 19921. If an animal is unable to adapt to stressors, it will develop abnormal physiologic or behavioral re- sponses; when this occurs, the animal is in distress (NRC, 19921. Some times, the effect induced by the stressor is pain. Pain can be described as a physical discomfort perceived by an organism as the result of injury, sur- gery, or disease. Once pain is perceived by an animal, it can itself become a secondary stressor and elicit other responses, such as fear, anxiety, and avoidance. To prevent or alleviate pain and distress in laboratory rodents, the re- search team should anticipate procedures or situations that will elicit these conditions. According to the U.S. Government Principles for the Utiliza- tion and Care of Vertebrate Animals Used in Testing, Research, and Train- ing, "unless the contrary is established, investigators should consider that procedures that cause pain or distress in human beings may cause pain and distress in other animals" (published in NRC, 1996, p. 821. Classifications of the magnitude of pain or distress estimated to be associated with differ- ent types of experimental procedures are available in the literature (NRC, 1992; OTA, 19861. It is the responsibility of the institutional animal care and use committee (IACUC) to evaluate each animal procedure for the potential to cause pain or distress and to ensure that anesthetics, analgesics, and tranquilizers are used, when appropriate, to prevent or alleviate pain and distress in the animals. Anesthetics or analgesics should be given before the painful insult, because it is easier to prevent pain, by blocking nociceptive neurons, than to alleviate it. The exposure of nociceptive neu- rons to painful stimuli produces chemical changes that cause the neurons to be hypersensitive to additional pain stimuli for a long period (Hardie, 1991; Kehlet, 19891. In addition, a cascade of physiologic changes occur that can have substantial effect on the recovery of an animal from surgery or on the information that is obtained in the procedure in which the animal is used. Depending on whether the pain is acute or chronic, responses might include
VETERINARY CARE 99 protein catabolism, sodium retention, immunosuppression, decreases in pul- monary and cardiovascular function, and increases in plasma concentrations of catecholamines and corticosteroids (Engquist et al., 1977; Flecknell, 1987; S. A. Green, 1991; Yeager, 1989~. Recognition of Pain and Distress Every person involved in the procurement, care, and use of laboratory rodents plays a major role in contributing to the total well-being of these animals. It is important to understand and consider species-specific behav- ior and husbandry needs when standard operating procedures and research protocols are developed to minimize exposure of the animals to situations that have a high probability of inducing pain and distress (Amyx, 1987; Montgomery, 1987~. Clinical signs and abnormal behavior displayed by rodents in response to pain and distress can include decreases in food and water consumption, accumulation of reddish-brown exudate around the eyes and nostrils (chromodacryorrhea), weight loss, decrease in activity, hunched posture, piloerection, poor grooming habits, labored respiration, vocalization, in- crease or decrease in aggressiveness, and self-mutilation (Flecknell, 1987; Flecknell and Liles, 1992; Harvey and Walberg, 1987; Heavner, 1992; NRC, 1992; Sanford, 1992~. The degree to which clinical signs are displayed varies within a species and between species. For behavior to be a useful indication of pain or distress, members of the research team, from animal caretakers to principal investigators, should be knowledgeable about the normal behavior of the animals with which they are working. Regular communication among all members of the research team, including the vet- erinary staff, is critical to ensuring timely evaluation and treatment of ani- mals in pain or distress. Alleviation of Pain The Guide recommends the use of appropriate anesthetics, analgesics, and tranquilizers for the prevention and control of pain and distress. How- ever, if for justifiable scientific reasons these agents cannot be administered when a painful procedure is to be conducted, the Guide states that the procedure must be approved by the committee fIACUC] and conducted by persons with adequate training and experience in the procedure used (NRC, 1996,p.101. The drugs routinely used to prevent or control pain in laboratory ro- dents are generally classified as either opioids or nonsteroidal anti-inflam- matory agents. Drugs reported to be effective analgesics in rodents are published elsewhere (Blum, 1988; CCAC, 1980; Clifford, 1984; Flecknell,
100 RODENTS: LABORATORY ANIMAL MANAGEMENT 1984, 1987; C. J. Green, 1982; Hughes, 1981; Hughes et al., 1975; Jenkins, 1987; Kruckenburg, 1979; Lumb and Jones, 1984; Soma, 1983; Vanderlip and Gilroy, 1981; White and Field, 1987~. In some cases, the doses quoted are extrapolations from doses for other species, with little or no scientific evidence to support the recommended use. Because some of these drugs might have systemic side effects that could interfere with a research proto- col, it is important to select and use them carefully. Additional factors that should be considered in selecting an analgesic include species, strain, age, sex, health status, nutritional status, period for which pain prevention or control will be required, recommended route of administration, volume of drug required for effect, compatibility with other pharmacologic agents that the animal will be receiving, cost, and availability (C. J. Green, 1982; Kanarek et al., 1991; Pick et al., 1991~. Principal investigators should get assistance from the attending veterinarian in selecting the most appropriate agent. Alleviation of Stress and Distress The use of tranquilizers can be considered when a laboratory rodent is restrained for long periods or used in a procedure that might cause fear, anxiety, or severe distress. Dosages of tranquilizing agents for rodents have been reported elsewhere (Blum, 1988; CCAC, 1980; Flecknell, 1987; C. J. Green, 1982; Harkness and Wagner, 1989; NRC, 1992; Vanderlip and Gilroy, 1981; White and Field, 1987~. It should be noted, however, that tranquiliz- ers have not been well studied in rodents. The drugs might interfere with experimental results, and suggested dosages might not produce the desired effects. Gradual conditioning to restraint before initiation of a study should also be considered as a means of decreasing associated anxiety or distress. SURVIVAL SURGERY AND POSTSURGICAL CARE Surgical procedures on rodents must be performed only by appropri- ately trained personnel or under the direct supervision of trained personnel (9 CFR 2.32; NRC, 1996 et seq., l991b). It is essential that personnel given the responsibility to perform surgery be knowledgeable about the principles of aseptic technique and the correct methods for handling tissues and using surgical instruments (McCurin and Jones, 19851. It is the responsibility of the IACUC to ensure that people approved to perform surgery on rodents have the required training or experience (9 CFR 2.321. Standards and guidelines for conducting survival surgery have been established by the Guide (NRC, 1996 et seq.) and for rodents other than mice and rats by the AWRs (9 CFR 2.311. Aseptic technique is required whenever a major survival surgical procedure is performed. Aseptic tech- nique is used to reduce microbial contamination to the lowest practical level
VETERINARY CARE 101 (Cunliffe-Beamer, 1993) and includes preparation of the animal, prepara- tion of the surgeon, sterilization of instruments and supplies, and the use of operative procedures that reduce the likelihood of infection. A major surgi- cal procedure has been defined as any surgical intervention that penetrates a body cavity or produces permanent impairment of physical or physiologic function (9 CFR 1.1; NRC, 1996 et seq.~. Other surgical procedures, classi- fied as minor, include catheterization of peripheral vessels and wound su- turing. Less stringent conditions are permitted for minor surgical proce- dures (NRC, 1996, p. 62), but sterile instruments should be used and precautions should be taken to reduce the likelihood of infection. Deviations from those guidelines and standards should not be undertaken unless reviewed and approved by the IACUC. The susceptibility of rodents to surgical infection has been debated; however, available data suggest that subclinical infections can cause ad- verse physiologic and behavioral responses (Beamer, 1972-1973; Bradfield et al., 1992; Cunliffe-Beamer, 1990; Waynforth, 1980, 1987), which can affect both surgical success and research results. Characteristics of surgery on rodents that can justify modifications in standard aseptic technique in- clude smaller incision sites, multiple operations at one time, shorter proce- dures, and complications caused by the use of antibiotics (Brown, 1994; Cunliffe-Beamer, 1993; Small, 1987; Wagner, 1976~. Strategies have been published that provide useful suggestions for dealing with some of the unique challenges of rodent surgery (Cunliffe-Beamer, 1983, 1993~. The area used for surgery, whether or not it is dedicated for that use, must be easily sanitized, must not be used for any other purpose during the time of sur- gery, and should be large enough to enable the surgeon to conduct the procedure without breaking aseptic technique. It might be necessary to perform experimental surgery on animals whose health has been compromised by naturally occurring or experimentally in- duced disease, but generally only healthy rodents should be used in experi- mental surgical procedures. Before being used in experimental surgery, rodents should be allowed sufficient time to acclimate to a new environ- ment and overcome the stress of transportation. Results of several studies have shown that mice experience increased corticosterone concentrations and depressed immune function after transport; these functions return to baseline values within 4-8 hours. The length of time might vary with the species and the mode and duration of transportation (Aguila et al., 1988; Dymsza et al., 1963; Landi et al., 1982; Selye, 19461. During the acclima- tion period, the animals should be examined to ensure that they are not exhibiting clinical signs of disease. To reduce or prevent stress preoperatively, researchers should be trained to handle and restrain animals and give them injections properly (NRC, l991b). The animals should be conditioned to being picked up and handled
102 RODENTS: LABORATORY ANIMAL MANAGEMENT by the people that will be doing the preoperative procedures. Fasting for periods of 12 hours or more is neither recommended nor generally required. However, it is often desirable to remove food at least 4 hours before anes- thesia to promote consistent absorption of intraperitoneal anesthetics (White and Field, 1987~. Access to water should be allowed up to the time that preoperative procedures are to begin (C. J. Green, 1982~. Anesthetics and Tranquilizers Administration of tranquilizers, sedatives, or anesthetics might prevent or alleviate stress in the animals, as well as making it easier for surgical personnel to prepare them for surgery. Dosages of tranquilizers and anes- thetics that can be used in rodents have been reported elsewhere (Blum, 1988; Flecknell, 1987; C. J. Green, 1982; Harkness and Wagner, 1989; Hughes, 1981; Kruckenburg, 1979; Soma, 1983; Stickrod, 1979; White and Field, 1987~. In addition to injectable and inhalational anesthetics, hypo- thermia has been recommended as a means of anesthesia in neonatal ani- mals (C. J. Green, 1982; NRC, 1992; Phifer and Terry, I986~. Criteria for selecting tranquilizers and anesthetics and their dosages should include spe- cies, strain, age, sex, health status, temperament, environmental conditions of the animal holding rooms, drug availability, drug side effects, recom- mended route of administration, equipment required, length of time that drug effect is desired, and skills and experience of the anesthetist. Doses quoted are often extrapolations from doses for other species with little or no scientific evidence to support them. It is important to select and use these drugs carefully to avoid interference with research protocols. Preparation for Survival Surgery Once the animal is tranquilized, sedated, or anesthetized, the operative site should be prepared. The extent of this preparation will depend on the species and maturity of the animal and on the complexity of the surgical procedure to be performed. The preparation might include removing body hair along the surgical site and surrounding areas with clippers, razors, or depilatory agents or by manual plucking. Care should be taken to avoid physical or chemical damage to the skin. Loose hairs should be thoroughly cleared from the surgical site. Various commercially available agents are appropriate for disinfecting the skin, including povidone iodine, alcohol, and chlorohexidine. Because the blink reflex is often lost under general anesthesia, consideration should be given to applying a sterile ophthalmic lubricant before surgery to prevent drying of the corneas (Powers, 19851. Heat loss can affect the course and success of anesthesia in rodents. Rodents lose body heat rapidly to surfaces such as operating tables, bench tops, and instruments. To preserve body heat, a circulating hot-water blan
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
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
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.
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.
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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