TRAPPED IN A GUILT CAGE
Arnold Arluke
The author is a professor in the Department of Sociology and
Anthropology at Northeastern University in Boston, Massachusetts.
(Animal Welfare Information Center Newsletter 4(2):1-2,7-8.
April-June 1993)
This article was originally published in the April 4, 1992,
edition of New Scientist (Vol 134 #1815). It is reprinted here
with the kind permission of New Scientist magazine.
I began entering laboratories in 1985 as an anthropologist
might study villages in other cultures. I would hang out and
become almost a native for weeks and sometimes months at a time,
so I could describe the culture of biomedical research. I
watched how researchers behaved with animals and with each other,
and asked questions about their work in everyday conversations
and more formal interviews. In some cases, I was even permitted
to do some of the work of technicians and caretakers, from
cleaning cages to carrying out experiments. So far, I have
studied 15 laboratories and research centers with around 400
principal investigators, veterinarians, postdoctoral and graduate
students, research technicians, and animal caretakers.
What prompted me to conduct this fieldwork was the
controversy over the propriety of animal research. While this
rapidly intensifying debate has led to greater regulation of the
use of animals in the laboratory, it struck me that little if any
attention has been paid to the impact of experiments on the
people who carry them out. It would be naive to think that
researchers might not experience some conflict over using animals
in experiments.
While most people I studied seemed to have come to terms
with their use of animals, many had not. Few people had frequent
signs of depression or anxiety, such as nightmares, sleep loss,
and increased alcohol consumption, that they attributed to
working with animals. However, more moderate and episodic
feelings of discomfort were common and were expressed as
background uneasiness and guilt. About 20 percent of the
interviewees, for instance, compared animal experimentation,
however tentatively, to the Holocaust. Uneasiness was
particularly noticeable among newcomers; with seasoned workers,
it was most common among animal caretakers. It existed among
technicians, and was relatively rare among veterinarians and
scientists. How did researchers live with whatever uneasiness
they felt?
Troublesome Emotions Denied
Open discussion of these feelings was taboo. Scientists,
veterinarians, and administrators tended to deny that laboratory
workers could be troubled by their use of animals. Uneasiness
was not seen as an issue, and was not allowed to intrude on the
normal course of work. This attitude was made apparent to me in a
firsthand way. Invited to speak at a conference of animal
researchers, I chose to call my talk, "The Experimenter's Guilt."
I was told that my choice was too controversial and that "Stress
Among Researchers" would be more palatable. A popular journal
about laboratory research invited me to publish this talk, but
insisted that the term "stress" was too extreme and inaccurate.
They preferred the term "uneasiness," which I used.
Soon after its publication, I was asked to speak on this
subject to the staff at the research center of a major
pharmaceutical company. I was told, however, that I could not
use "uneasiness" in the title because it would inflame research
directors. They suggested "How Researchers Deal with Their
Feelings". To make matters easier, I have decided simply to call
future talks "Untitled".
New workers believed they were not supposed to talk about
their feelings to anyone. Feelings remained private, extraneous
to the "real work" of the laboratory. Individuals believed that
their colleagues were better able to handle their feelings, only
vaguely aware that many others dealt with the problem in similar
ways. Yet within the laboratory culture were unspoken rules and
resources for dealing with unwanted emotions and thoughts,
despite the silence surrounding this topic.
People most commonly coped by seeing laboratory animals as
different from pets, zoo animals, or wild animals. Once the
creature was defined as a laboratory animal, certain emotions
would not be tapped, making it easier to carry out experiments.
Many social forces in the laboratory culture helped to make this
definition. Animals became "models" chosen to suit particular
experiments. Their cost was listed under "supplies" in grant
proposals, and they could be ordered through catalogues of
animals specially bred for laboratory use.
Turning Animals Into Objects
As interchangeable and anonymous objects, each animal or an
entire cage was identified by a code that might include the date
of delivery, the researcher's name, the experiment's number, and
the animal's number. These codes were clearly displayed on all
cages, and the identification numbers of some animals were marked
on their bodies: the ears of mice were hole-punched and the
bellies of dogs and primates were tattooed. Unstated rules
dictated how people interacted with laboratory animals. Social
norms stipulated that they were objects and not pets, and
sanctions supported this definition. For example, the chief
technician in one laboratory had to tell a worker to stop naming
sheep because that made it harder for others to perform the
experiments.
Making this definition was easier for researchers than it
was for technicians and caretakers. Having taken laboratory
courses that used animals in college or medical school, many
researchers learned not to make laboratory animals into pets long
before starting their first full-time research positions.
Instead, animals were transformed into data or silent research
collaborators. Lack of direct contact with the animals
reinforced the transformation. Researchers, typically, did not
routinely conduct experiments and handle animals; they stopped by
their laboratories for a brief visit during the day or
occasionally performed delicate surgery on animals after they
were fully anesthetized. Also, most applied biomedical
researchers were primarily interested in answering particular
scientific questions, and animal models would be selected on that
basis.
Technicians and caretakers found it harder to treat animals
as objects because they commonly lacked prior research experience
and had frequent and direct contact with the animals. They would
learn not to treat them as pets after being shocked by the death
of a special animal that they regarded as a friend or partner.
While people tried to detach themselves from the animals, they
rarely succeeded completely. Some described themselves as "a
little desensitized". In the words of one technician: "You have
to put up some walls. Sometimes you have to create a distance
between yourself and the animal you are working with. But I try
occasionally to do some checking to see how big that distance is.
I don't want it to become so big that I lose the sense that I'm
working with animals."
While most people accepted this detachment as necessary for
self-protection, not everyone found it comfortable. One
technician, for instance, told me that "it didn't feel right" to
stop playing with the primates in her laboratory. But those who
did bond closely to laboratory animals were often reminded and
even teased about the dangers. At one facility, for example, a
technician was called a "problem child" by her peers for this
reason. At another facility, in an effort to curtail bonding, a
scientist told his technicians to remove the names of animals
from cage identification cards because it "looked
unprofessional".
Workers still found ways to treat animals as pets and
express their affection for them. Technicians and caretakers
would single out an animal for a laboratory pet. Often a mouse,
rat, or guinea pig, these animals were not experimented upon or
at least not sacrificed. In addition to being named, caged
singly, fed special foods, and given much attention, they would
also sometimes be taught tricks and allowed occasionally to run
free in the laboratory. They were safe animals with whom workers
could become attached without fear of loss. Affection for animals
also resulted in "rescues" where they were taken home by workers
who were strongly attached to them. For instance, in all seven
dog laboratories studied, staff members had quietly taken home at
least one animal in the previous year. And photographs, cartoons,
dolls, and other images of animals hung on the walls of
laboratories, as constant reminders to workers that they cared
about animals and found them interesting.
It was also important for people to learn to cope with the
death of animals. Novices were usually eased and coached into
killing their first animals. Sometimes long before they did their
own killing, they observed others doing it matter-of-factly.
More experienced people almost never cajoled or pushed newcomers
to kill and waited until they seemed ready to do it. Still,
certain types of sacrifice were contrary to the novice's
"instincts", such as slamming rodents against the bench or
cutting off their heads, and this required special teaching.
Newcomers were reassured that, if done correctly, the death was
quick and painless, regardless of the particular method they
used. For example, after breaking the necks of mice, new workers
were often troubled by animal movements that looked like
suffering. Someone more senior would usually explain to the
novices that these movements were only "muscle spasms."
Rituals Help Workers To Cope
For some people, it was important not to see death as just
another task in the day because it would quickly become
mechanical, especially in laboratories that conducted experiments
like factory assembly lines where the individuality of animals
was lost. As one researcher said, "It doesn't mean that we're
callous about killing them, but there's not really a second
thought for that animal as an individual." Death could become
merely the final step in the protocol, signifying noxious tasks
such as disposing of corpses and more pleasant associations such
as going home for the day. In a few laboratories, workers
followed certain rituals when killing animals, giving death
special meaning. In one case, the scientist asked her graduate
students and technicians to observe a minute of silence before
sacrificing animals. In another laboratory, a technician
privately recited a prayer each time she killed an animal, asking
that its death be forgiven. Some laboratories made memorials to
commemorate "favorite" animals that died.
Yet for most people, using the term "sacrifice" was the
primary device for giving meaning to death. Journals and grant
agencies prohibit use of this term, and some individuals
described it as an inappropriate euphemism, but it did indeed
mean something special to many research workers. "`Killing'
connotes no purpose, while `sacrifice' connotes there is a
reason," noted one technician. Similarly, an investigator
explained to me that "sacrifice" was different from "wanton
murder" described in detective novels; the former had a larger,
worthwhile aim while the last was pointless. Besides
"sacrifice," there were other terms with less meaning that
shielded people from the harshness of death. Animals were
"dispatched," "terminated," "cervically dislocated,"
"exsanguinated," "decapitated," or "put down," while whole rooms
were "depopulated" or simply "cleaned."
People also acquired a vocabulary that aggressively framed
their actions toward animals, reinforcing the image of animals as
objects. People injecting animals were shooters and their
injections were "sticks." "Guns" were syringes attached to
devices like pool cues that reached into cages, and "torture
chambers" were devices to restrain mice. Animals were labeled
according to their experimental purpose: there were "controls," "
recipients," "donors," "carriers," "bleeders," "breeders,"
"junk," or simply "X-animals." Even the very term "experiment"
was infrequently used; people more often referred to a
"preparation" or "project." And the subjective term "suffering"
was deliberately avoided in favor of the more neutral "distress."
The Scientist As `Hunter'
Rationalizing the use of animals in science was also a
mainstay in the coping skills of researchers. People in
laboratories saw little difference between animals used in
experiments and those killed for food and clothing. A few
compared it to hunting, which they saw as acceptable if animals
were eaten rather than killed merely for recreation. As one
researcher said of his hunting: "I do it strictly for the meat
from the rabbits, to pheasants, to ducks, to geese. I've had
opportunities to shoot bear, but I haven't because bear meat
isn't good to eat and I can't see killing something that I can't
use personally." I was frequently reminded that most laboratory
animals were bred for research, so they knew of no other
existence. And when former pets and strays were obtained from
shelters where they would have been killed "wastefully," their
use in experiments was seen as giving the animal's life and death
added purpose.
For the most part, though, people did not have elaborate
moral justifications for their use of animals. Instead, many of
them appeared ethically inarticulate. Predictably, scientists
and research technicians saw scientific and medical goals as
moral imperatives to do their work. Caretakers justified their
work with animals by ensuring that they could not be better
treated, giving the animals enough love and attention in their
last days so they could experience what it was like to be loved
as pets. For some workers this was almost an addiction. People
spoke about being unable to quit because they were afraid that no
one else could be hired that would be as dedicated as they were
to the welfare of laboratory animals.
I also observed a different way of coping among those who
felt "animal activists" seriously threatened biomedical research.
Some scientists have started a countermovement to educate the
public about the need to use animals in science. Part of this
campaign has been to denounce activists as dangerous and evil
because medical advances would halt if they succeeded in
preventing animal experiments. By demonizing those strongly
opposed to animal research, the charge of immorality levelled at
researchers was reversed.
Finally, researchers had to learn to manage the occasional
sarcastic remark, heated argument, or blunt criticism encountered
when discussing their work with lay people. New workers were
often disturbed to be called "mouse murderers" and discovered
that conversations about animal experimentation quickly
degenerated into a "ping pong" of polarized opinions. Scientists,
though, were less likely than technicians or caretakers to be put
in this position because as physicians or academics they could
talk about their work without mentioning animal experimentation.
Also, their social networks usually included many people
sympathetic to biomedical research. Those not in this position
would sometimes, out of frustration, carefully avoid mentioning
animal experimentation by telling people that they "did cancer
research" or "worked at Boston General Hospital." Others would
assess whether conversations were likely to become "shouting
matches," gradually releasing more information about their use of
animals as long as the unfolding talk seemed safe to them. Some
also told people that they owned pets themselves, perhaps to
suggest that they were hardly insensitive and heartless
scientists.
While these coping devices certainly made it easier for many
people to conduct experiments on animals, it is not clear whether
these adjustments should be encouraged. There are two lines of
thinking. Some people argue that by coping in this manner, there
will be an ethical blunting or a coarsening of the moral
sensitivities of researchers. Others are more struck by the
significance of the conflicts that prompt defensive behavior. The
surfacing of these conflicts among researchers may be due to the
diffusion into the laboratory of society's heightened awareness
of how animals should be viewed and treated. Coping devices will
be called out when humanity's standards clash with traditional
scientific practice. This is cheering to some who see this as a
willingness to pay more attention to humanitarian ideals in
animal experimentation.
Applying Principles of Aseptic Surgery to Rodents
Terrie L. Cunliffe-Beamer, DVM, MS
The author is Head, Clinical Laboratory Animal Medicine, The
Jackson Laboratory, Bar Harbor, Maine
(Animal Welfare Information Center Newsletter, 4(2):3-6.
April-June 1993)
The 1985 revision of the Public Health Service Guide for the
Care and Use of Laboratory Animals (`PHS Guide') (Committee,
1985) and 1985 amendments to the Federal Animal Welfare Act (9
CFR, 1992) both contain provisions requiring aseptic technique
for rodent survival surgery. The `PHS Guide' applies to all live
vertebrate animals used in research and, thus, includes
laboratory rats and mice. Regulations of the Animal Welfare Act
apply to hamsters, guinea pigs, and unusual laboratory rodents,
but currently exclude rats of the genus Rattus and mice of the
genus Mus.
Rodents are widely used in biomedical research, as evidenced
by 55,074 citations for 1990 and 46,519 citations for 1991 under
the Medline (on-line database of the National Library of
Medicine) heading "Rodentia". However, only approximately 1.2
percent of the Rodentia citations (741 citations in 1990 and 548
citations in 1991) reported surgical procedures. When Rodentia
citations with surgical procedures were subdivided by species of
rodent, rats were first with the most listings, mice were second,
and guinea pigs were third. Hamsters, gerbils, and other rodents
were a distant fourth.
Occasionally, the argument is still made that aseptic
technique is not necessary for rodent surgery because mice or
rats often survive surgical procedures performed using less than
aseptic technique. However, survival alone is not a valid
criterion for judgment of the acceptability of a rodent surgical
technique. The criterion for acceptability should be the absence
of untoward, unplanned alteration of physiological functions or
behavior due to perioperative infection. Post-surgical adhesions
and subclinical infection can complicate analysis or observation
of tissues. Failure to utilize aseptic surgical technique
increases the potential for introducing bacteria and activating
immune responses in reaction to the bacteria. Recently,
responses of rats subjected to aseptic or septic surgical
procedure were compared. Although there were no obvious clinical
signs in either group of rats, differences were observed in open
field behavior, "freezing" behavior, plasma fibrinogen, serum
glucose, total white cell count, and wound histology scores
(Bradfield, Schachtman et al. 1992). Activation of macrophages
in response to intraperitoneal inoculation of bacteria (Bancroft,
Schreiber et al. 1989), stimulation of cytokines and activation
of B cells by bacterial endotoxins (lipopolysaccharides) (Abbas,
Lichtman et al. 1991), and alterations of other physiological
processes by subclinical viral, mycoplasmal, bacterial or
parasitological infections (Committee on Infectious Diseases of
Laboratory Rats and Mice 1992), are well documented in the
literature. It has been documented that use of aseptic surgical
technique has increased the success of ovarian transplants in
mice and speeded the return to normal following other surgical
procedures in mice (Cunliffe-Beamer 1972-73; Cunliffe-Beamer
1990).
A further argument for aseptic surgical technique in rodents
is the fact that hamsters and guinea pigs are intolerant to many
antibiotics. In these species, antibiotics can selectively
destroy gram positive intestinal flora resulting in overgrowth of
gram negative organisms and endotoxemia (Wagner 1976; Small
1987). Administration of antibiotics to "protect" against the
consequences of poor aseptic technique could increase morbidity
and mortality in hamsters and guinea pigs.
Development of protocols for aseptic rodent surgery can
challenge the attending veterinarian, principal investigator, and
Institutional Animal Care and Use Committee. The challenges
arise from several sources. First, the same person often serves
as surgeon, anesthetist, surgical technician, and scrub nurse
when surgical procedures are performed on rodents. Careful
planning is required to assure that all supplies and equipment
required to complete the surgical procedure are not only ready
for use, but are also placed exactly where they are needed before
surgery begins. Second, experimental design frequently requires
repetitive surgery, that is, performing the same surgical
procedure on individual members of a group of rodents during a
single sitting. In repetitive rodent surgery, it may not be
feasible to have a new sterile pack of instruments for each
rodent. Procedures to decontaminate instruments between each
rodent must be developed. Third, the small body size of many
laboratory rodents mandates dissecting microscopes and delicate
microsurgical or ophthalmic instruments for many otherwise
routine surgical procedures.
The `PHS Guide' defines major survival surgery as "any
surgical intervention that penetrates a body cavity or has the
potential for producing a permanent handicap in an animal that is
expected to recover." The standards of the Animal Welfare Act
in part 1.1 similarly define a major operative procedure as "any
surgical intervention that penetrates and exposes a body cavity
or any procedure which produces permanent impairment of physical
or physiological functions." Minor surgeries, by default, are
all surgical procedures that do not penetrate a body cavity or
produce a permanent impairment of function. However, one should
remember that a relatively minor surgical procedure, such as
vascular catheterization, can have life-threatening complications
if bacteria are introduced into the blood stream.
The `PHS Guide' states that "survival surgery on rodents...
should be performed using sterile instruments, surgical gloves,
and aseptic procedures to prevent clinical infections." The
standards of the Animal Welfare Act in part 2, state "...survival
surgery will be performed using aseptic procedures including
surgical gloves, masks, sterile instruments, and aseptic
technique." However, neither document further defines aseptic
surgical technique in detail. The primary objective of aseptic
surgical technique is to reduce microbial contamination of the
incision and exposed tissues to the lowest possible practical
level. Items to address during development of aseptic technique
for repetitive rodent surgery include (1) selection and
sanitation of surgical table and associated equipment, e.g.,
microscopes, (2) preparation and sterilization of surgical
instruments, (3) maintenance of sterility between rodents, (4)
decontamination of skin surrounding the incision site, (5) use of
surgical drapes, and (6) preparation of the surgeon.
When major survival surgical procedures are performed on
non-rodents, `PHS Guide' and standards of the Animal Welfare Act
require a dedicated surgical facility. In this facility, the
`PHS Guide' requires separate areas for performing the surgery,
storing supplies and preparing surgical instruments, preparing
the animal for surgery, preparing the personnel, and providing
intensive care and supportive treatment of post-operative
animals. A dedicated surgical facility is not required for major
survival rodent surgery by either the `PHS Guide' or the Animal
Welfare Act. A rodent surgical area can be a room or part of a
room that is easily sanitized and not used for other activities
when rodent surgery is in progress. The area should be
subdivided so that there are specific places for cages of rodents
awaiting or recovering from surgery, preparing rodents for
surgery, and performing the surgery. This approach reduces the
potential for contamination of the surgical field by fur, feces
and bedding. Before beginning rodent surgery, the laboratory
bench or table where the surgery will be performed should be
cleaned and disinfected. Quaternary ammonium disinfectants or
70% alcohol are good choices for disinfecting laboratory benches
prior to rodent surgery. Laboratory benches in front of open
windows, next to doors, or similar locations where air currents
and dust are difficult to control should be avoided as rodent
surgery tables. Likewise, rodent surgery should not be performed
in or in front of an exhaust hood because air and particulates
from throughout the laboratory are drawn over the surgical field.
A high efficiency particulate absorbent (HEPA) filtered hood can
be used as a rodent surgical area if the air flow within the hood
does not desiccate exposed tissues. A glove box or plastic
bubble can be used to create an isolated "rodent surgical suite"
within a laboratory or animal treatment room.
Surgical instruments used in rodent surgery usually have
delicate tips that are easily damaged. Autoclavable tip guards
are commercially available and should be used to protect tips of
instruments. Special instrument trays with rows of soft plastic
fingers can be used instead of flat trays to store delicate
instruments. The plastic fingers prevent instruments from
sliding into each other if the tray is tilted. After use,
instruments should be soaked in lukewarm water to remove blood
and tissue, washed with a free rinsing neutral pH detergent,
rinsed thoroughly, and air dried. A toothbrush can be used to
scrub delicate surgical instruments. Before delicate instruments
are returned to storage, the tips should be examined, preferably
under a microscope, to be certain that the ends meet properly,
and grooves should be examined to verify that no blood or tissue
remains in grooves. The cutting edge of microdissecting scissors
should be examined under a microscope and be tested by cutting a
single thread in a gauze sponge or piece of fine suture.
Instruments with damaged tips or dull blades should not be used
because their use can increase the amount of trauma associated
with the surgical procedure.
Methods to sterilize surgical instruments include steam, dry
heat, ethylene oxide, chemical sterilants, and radiation (Block
1991). By definition, sterilization means the absence of
microbial life, including viable bacterial spores. Steam or dry
heat are preferred methods to sterilize surgical instruments.
Sterilization should be verified through periodic use of
biological indicators manufactured for this purpose. Glass bead
sterilizers are a fast way to sterilize unwrapped surgical
instruments (Callahan, Fiorillo et al. 1992). However,
instruments must be allowed to cool on a sterile surface before
use to avoid thermal injury (burning tissues). Instrument packs
sterilized by ethylene oxide must be aerated to remove residual
gas. Some chemical sterilants, e.g., chlorine dioxide, are
corrosive to metals as well as irritating to tissues. Even
noncorrosive chemical sterilants can be irritating to tissues.
If chemical sterilants are used on surgical instruments,
sufficient time must be allowed to achieve sterilization and
instruments must be rinsed with sterile water or sterile saline
before use. Contact time varies with the chemical sterilant and
manufacturer's instructions should be consulted for contact time
required to achieve sterilization. Rinse solutions should be
changed frequently to prevent contamination by the sterilant.
Quaternary ammonium, iodophor and phenolic disinfectants
used to sanitize animal facilities should not be used on surgical
instruments. These disinfectants are not sterilants. Alcohol,
contrary to popular belief, is neither a sterilant nor a
high-level disinfectant (Block 1991; Rutala 1990).
Recommendations for selection of disinfectant based on the
physical make-up of the instrument and its use have been
published (Rutala 1990).
Maintaining sterile instruments when performing repetitive
rodent surgery is a challenge. Contamination can be reduced by
segregating surgical instruments according to function. Surgical
instruments used to incise the skin are placed at one end of the
tray. Instruments used in subcutaneous tissues are placed next
to the skin instruments. Instruments used within internal
cavities are placed next to instruments used in subcutaneous
tissues and so on. The tips of the instruments are placed toward
the top of the tray. This arrangement places instruments used in
deep body tissues "off to the side" and minimizes reaching over
them to reach other instruments (Cunliffe-Beamer 1983;
Cunliffe-Beamer 1990).
Contamination of instruments by aerobic bacterial skin
contaminants in repetitive rodent surgery can be reduced by
wiping tips of instruments with 70% alcohol and a sterile swab
between rodents. Alternatively, a glass bead dry heat sterilizer
could be used after the tips of instruments are wiped with
sterile saline or water to remove blood or tissue residue. Use
of a sterile instrument holder with pockets also reduces
potential for contamination because tips of instruments can be
tucked in the pocket and covered while the next rodent is
prepared for surgery. Even with alcohol wipe between rodents and
holder with pockets, a new sterile instrument pack should be used
after 4 or 5 individual rodents.
A surgical drape is a sterile cover that is draped over all
or part of the rodent. The drape protects against accidental
contamination of surgical instruments by providing a sterile
"buffer zone" and provides a sterile surface on which to lay
exteriorized organs. Surgical drapes for rodents can be made
from a variety of materials. Lightweight, clear plastic drapes
manufactured for larger animals can be cut in small pieces and
steam sterilized between two paper towels. This type of drape
conforms to the rodent's body and makes it easy to observe
respiration. Opaque disposable paper or cloth drapes make it
difficult to monitor respiratory rate of small rodents. In some
circumstances, a sterile non-woven surgical sponge can be used to
"drape" a small rodent.
Preparation of the incision site is an important part of
aseptic technique. If fur is not removed over the incision site
and skin is not decontaminated, hair and associated skin bacteria
can be carried into deeper tissues. Alternatives for removing fur
from rodents include plucking, clipping, shaving, or in selected
instances, depilatories. Plucking the fur from an anesthetized
mouse or similar-size rodent has many advantages. It is fast and
easy and does not leave a stubble. Hair follicles in adult mice
are usually in the telogen (resting) phase, and the hair can be
removed manually with minimal injury (Sundberg 1993). If fur is
removed with clippers, pressing a piece of adhesive tape over the
clipped area picks up loose hair that would otherwise migrate
into the incision. Use of depilatories should be reserved for
situations where complete removal of fur from a very large area
of skin is required. If the depilatory remains in contact with
the skin for too long, a chemical burn could result. After the
fur is removed from the area where the incision will be made, the
skin needs to be cleansed and disinfected. In large rodents,
e.g., rats or guinea pigs, skin can be washed with soap, rinsed
with water, and disinfected with 70% alcohol or a surgical
iodine. In small rodents, three applications of 70% alcohol, or
two applications of 70% alcohol and one application of surgical
iodine are often used to disinfect rodent skin. Sterile gauze
sponges or sterile cotton swabs, depending on the size of the
rodent can be used to disinfect the skin. Begin at the incision
site and work outward in circles of increasing diameter (Bennett,
Brown et al. 1990).
It is difficult to generalize about rodent surgery because
the "patient" can vary in body weight, from a 1.5 or 2.0 gram
new-born mouse to a 500-700 gram rat or guinea pig. The
magnitude of this difference on a percent-body-weight basis is
equivalent to comparing a 2 or 3 kg cat and a 765 kg horse. Even
among rodents, surgical instruments must be matched to the size
of the patient. Surgical procedures in small rodents, e.g.,
young mice, require delicate instruments such as those designed
for micro or ophthalmic surgery in order to minimize surgical
trauma. Several books contain detailed descriptions of rodent
surgical procedures (Waynforth 1980; Cunliffe-Beamer 1983).
Water is not usually withheld from small rodents prior to
surgery. The inability of mice and rats to vomit prevents
regurgitation of stomach content. The nibbling nocturnal feeding
behavior of most small rodents and rapid intestinal transit times
combine to eliminate distended digestive tracts as a problem for
most laboratory rodent surgery. Thus, withholding food is not
common practice prior to many rodents surgical procedures,
although guinea pigs are often fasted prior to surgery (Harkness
and Wagner 1989).
Hypothermia from anesthesia, wetting a significant portion
of the body during preparation for surgery, or cooling of exposed
body cavities is a potential problem during any rodent surgery.
Decontamination of the skin should be accomplished without
soaking the body of the rodent. The degree of hypothermia is
influenced by the type and duration of anesthesia (Gardner, Davis
et al. 1992) and environmental factors. Heat transfer should be
considered when selecting the surgical table. Stainless steel is
easy to sanitize, but it conducts heat away from the body. A
temperature-controlled small water `blanket' should be placed
under the rodent during prolonged surgical procedures. A cork
board, a plastic tray, or a few paper towels can be placed under
the rodent to minimize heat transfer during short procedures.
Post-operative care should include an external heat source while
the rodents recover from anesthesia. The heat source should be
positioned so that the rodents can move away from it as they
recover from anesthesia. An electric light (50-75 W bulb)
suspended over one end of the cage is a very simple heat source
for rodents recovering from anesthesia.
In summary, when aseptic surgical technique is not
practiced, infection can be expected. These infections are often
subclinical in rodents; nevertheless, adverse physiological
effects have been demonstrated. Preventing post-surgical
infection by using aseptic technique improves the quality of life
for the rodent and eliminates a source of uncontrolled variation
in research data.
References:
9 Code of Federal Regulations. Chapter 1, Subchapter A-Animal
Welfare.
Abbas, A., A. Lichtman, et al. (1991). Cellular and Molecular
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U.S. Food and Drug Administration
Animal Use in Testing FDA-Regulated Products
Position Paper
(Animal Welfare Information Center Newsletter 4(2):14. April-June
1993)
Current laws administered by FDA including the Federal
Food, Drug and Cosmetic (FD&C) Act are intended to ensure product
safety and effectiveness, thereby protecting consumers' health.
These laws place responsibility on FDA to ensure that human and
animal drugs, biologics, and medical devices are safe and
effective and that food products are safe and wholesome.
Animal testing by manufacturers seeking to market new
products is often necessary to establish product safety. FDA
supports and adheres to the provisions of applicable laws,
regulations, and policies governing animal testing, including the
Animal Welfare Act and the Public Health Service Policy on Humane
Care and Use of Laboratory Animals. Moreover, in all cases where
animal testing is used, FDA advocates that research and testing
derive the maximum amount of useful scientific information from
the minimum number of animals and employ the most humane methods
available within the limits of scientific capability.
FDA advocates the use of validated non-whole animal
techniques, which may include such screens and adjuncts as in
vitro (e.g., tissue culture) methodologies and biochemical
assays. As an example, FDA announced in the Federal Register of
Feb. 19, 1988, the availability of guidelines for the Limulus
Amebocyte Lysate (LAL) test as an end-product endotoxin test for
human injectable drugs (including biological products), animal
injectable drugs, and medical devices. The guidelines inform
manufacturers of acceptable methods of validating the LAL test so
that it can be used as an alternative to the rabbit pyrogen
test. At present, many other procedures intended to refine,
reduce, or replace animal testing are still in the relatively
early stages of development.
With respect to cosmetic products, the FD&C Act does not
specifically require that cosmetic manufacturers test their
products for safety in the context of premarket approval by the
agency. However, FDA strongly urges cosmetic manufacturers to
conduct toxicological or other tests necessary to substantiate
the safety of a particular cosmetic product. If the safety of a
cosmetic product is not adequately substantiated, the product is
considered misbranded and may be subject to regulatory action
unless the principal display panel bears the statement "Warning
the safety of this product has not been determined."
Much of the attention given to animal testing has focused on
the LD50 test and the Draize eye and skin irritancy tests. FDA
does not require LD50 test data to establish levels of toxicity,
and in 1988, published a policy statement in the Federal Register
to clarify this position.
The Draize eye and skin irritancy tests continue to be
considered among the most reliable methods currently available
for evaluating the safety of a substance introduced into or
around the eye or placed on the skin. Non-animal tests, such as
in vitro tests, may be useful as screening tools to indicate the
relative toxicity or safety of a substance that comes into
contact with the eye or skin. However, the responses and results
of in vitro tests alone do not necessarily demonstrate the safety
of a substance. The effects of a substance on a biochemical
reaction or on a specific cell or tissue in culture may differ
from its effect on a specific organ system in the whole animal.
The precise nature of testing needed to determine the safety
or effectiveness of a specific product regulated by FDA depends
upon the characteristics and intended use of the product. More
specific guidance may be obtained through consultation with FDA
scientists on a case-by-case basis.
October 1992
Go to:
The Animal Welfare Information Center
U.S. Department of Agriculture
Agricultural Research Service
National Agricultural Library
10301 Baltimore Ave.
Beltsville, MD 20705-2351
Phone: (301) 504-6212
FAX: (301) 504-7125
Contact us: http://www.nal.usda.gov/awic/contact.php
