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OCR for page 85
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
OCR for page 86
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
OCR for page 87
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
OCR for page 88
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
OCR for page 89
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
OCR for page 90
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
OCR for page 91
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.
OCR for page 92
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
OCR for page 93
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
OCR for page 94
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 95
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
OCR for page 103
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
OCR for page 104
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.
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Representative terms from entire chapter:
laboratory animal
