Non-response on Protocols Pending Approval
If a there are questions on a submitted protocol and the PI fails to respond within 30 days
of a written letter, the protocol will be administratively withdrawn.

Post-approval Monitoring
The Montana State University post-approval monitoring program is in place to ensure the
well-being of animals involved in research activities, to make certain that activities are
conducted in compliance with state and federal regulations and standards, and to ensure
that activities are consistent and conducted in compliance with IACUC approved protocols,
as well as other institutional policies.

Since the animal care and use program at MSU is fairly small with only seven employees,
there is no one person or institutional component responsible for performing post-approval
monitoring. A multitude of individuals contribute to the post-approval monitoring program,
including the IACUC, the research staff, the ARC staff, and the Attending Veterinarian. A
variety of procedures are in place to assure compliance. These include:

Protocol Modifications
Any proposed modifications of animal use methods must be communicated to the IACUC.
Major modifications will be reviewed and approved by the full committee
prior to implementation. Examples of major modifications include the following changes:

The objectives of the study

Non-survival to survival surgery

Increasing the invasiveness of a procedures or pain/distress to the animal

Change or addition of species

Increasing the number of animals used

Anesthetic or analgesic agents

Method of euthanasia

Status of biosafety, radiation safety, or environmental safety usage

Change in the Primary Investigator

Reporting Animal Concerns (Whistle-Blower Policy)
The use of animals in biomedical research is necessary and appropriate. Montana State
University is committed to the humane treatment of animals and strives to ensure that
institutional facilities and procedures adhere in all respects to NIH guidelines for care and
use of laboratory animals.

The IACUC wants to be informed of any concerns that individuals of the campus
community may have with respect to the care and use of laboratory animals and
feels it is important that these concerns be addressed on an individual basis. Any
individual who witnesses what he/she perceives to be inappropriate treatment of
research animals can contact any member of the IACUC, or can file a concern
anonymously by filling out an “Animal Concern Form” (posted at all locations
where animals are housed) and submitting it to the IACUC. The identity of the
person raising the concern will be held in confidence. Retaliation against a person who
raises a good-faith concern
will not be tolerated.

The IACUC will follow-up on all complaints and gather additional information as
necessary and appropriate. If indicated, corrective action will be initiated and the
results of the investigation will be communicated to the individual who raised the complaint.

Research Projects using Insects
Protocols utilizing insects are exempt from IACUC review.

Research Projects using Chick Embryos
Protocols utilizing chicken embryos up to day 15 of incubation are exempt from IACUC review.

Tours, Videotapes and Photographs
Tours of the ARC must be approved by the ARC Director or Manager. Requests will be
reviewed on an individual basis, and if questionable, will be referred to the IACUC for
approval. Any request for videotaping or photographing animals and animal procedures
need to be approved by the IACUC. In addition, once approved, the IACUC can select to
review the finished product before its release or publication. Cell Phones are prohibited from
use in the ARC as the majority of cell phones have photography capabilities.

Vivarium Access
Both the Animal Resources Center and the Jutila Research Laboratory required keycard
access. For the ARC, individuals must attend the orientation session and complete the
AALAS Learning Library on-line course titled “Working with the IACUC” prior to gaining
access to the facility. There are three levels of access that can be grants, and the access
given is dependent on the request of the PI in whose lab the person will be working. Access
options include business hours only, ABSL1 area only, and ABSL2 access.
The Investigators are responsible for notifying the ARC Manager when
someone is no longer working in the laboratory. At that time, the keycard is inactivated.

For the Jutila Research Laboratory, individuals must complete select agent requirements
and access must be granted through the MSU Biosafety Officer and the Laboratory Director.

Single Housing of Rodents

The Guide for the Care and Use of Laboratory Animals (p. 37) states, “Consideration should be given to an animal’s social needs. The social environment usually involves physical contact and communication among members of the same species…..When it is appropriate and compatible with the protocol, social animals should be housed in physical contact with conspecifics.”

At Montana State University, every attempt will be made to socially house rodents. The IACUC recognizes that social housing is not always possible for experimental, health, and behavioral reasons. Some strains of rodents engage in barbering behavior and/or aggression which results in wounding of cage mates.

If individual housing is a required part of a research project, it should be discussed and justified within the animal care and use protocol. If single housing is required for health, behavioral, or breeding colony reasons it may be performed by the Animal Resources Staff using the appropriate professional judgment. Individual IACUC approval is not necessary for these cases.

Sanitation Frequency of Primary Enclosures for Rodents

The Guide for the Care and Use of Laboratory Animals (p. 43) states, “In general, enclosures and accessories, such as tops, should be sanitized at least every 2 weeks. Solid-bottom caging, bottles, and sipper tubes usually require sanitation at least once a week. Some types of cages and racking might require less-frequent cleaning or disinfection; these might include large cages with very low animal density and frequent bedding changes, cages that house animals in gnotobiotic conditions with frequent bedding changes, individually ventilated cages, and cages used for special circumstances.

The standard husbandry practice at the MSU Animal Resources Center is to sanitize rodent solid-bottom caging on a weekly basis. Under certain circumstances, sanitation may be performed every two weeks. Examples include rodents housed at a low density in individually ventilated cages, dams with new litters, and rodents housed in a high biocontainment environment.

The MSU IACUC understands that sanitation of primary enclosures for rodents is typically performed on a weekly basis, but under circumstances such as those described above, may be extended to every two weeks. Animal Resources Staff, utilizing professional judgment, will be allowed to determine the frequency of rodent cage sanitation not to exceed once every two weeks.

Animal Procedure Policies

Access to Nonhuman Primate Areas
Personnel who require access to the nonhuman primate areas are required to attend a
nonhuman primate orientation session administered by the Director of the ARC. They
must also complete the AALAS Learning Library course on nonhuman primates.
Personnel who enter nonhuman primate areas are required to wear a dedicated
long-sleeved lab coat, gloves, disposable bonnet, surgical mask, shoe covers,
and eye protection. Long pants are required and open-toed shoes are not allowed.

Escaped Rodents
If a rodent escapes during the process of cage cleaning or handling and can be easily
identified, it is placed back in the appropriate cage. If a rodent is found loose in an animal
holding room and cannot be positively identified, it is placed in a separate cage and taken
to the necropsy room for euthanasia. If a rodent is found loose within the facility outside
an animal room, such as in the feed and bedding storage area, cage wash room, etc. the
animal is captured, placed in a separate cage and taken to the necropsy room for euthanasia.
Prior to euthanasia, the ARC staff will perform a health analysis on the rodent similar to
that done for sentinel animals. Blood will be collected for serology and the animal will be
examined for external and internal parasites.

Euthanasia
When required for scientific data collection or clinical or operational requirements,
animals will be humanely killed, as detailed in the approved protocol. Where applicable,
a secondary method of euthanasia will be performed. Unintended recovery of animals
after apparent death constitutes serious noncompliance with PHS Policy and serious
deviation from the provisions of the Guide for the Care and Use of Laboratory
Animals. MSU is obligated to report to the NIH any instances of animals awaking
after intended death.

Public Health Policy (PHS) requires IACUC’s to determine that methods of
euthanasia proposed in protocols meet criteria outlined in the 2007 Report of the
AVMA Panel on Euthanasia. Primary Investigators (PI) are expected to follow the
methods of euthanasia as outlined and approved in their protocols. All individuals
who may perform or be expected to perform euthanasia should be familiar with
the details in the protocol and the information outlined in this policy.
Training in proper methods of euthanasia is part of the initial orientation
and is provided by the ARC Director and Staff.

Acceptable methods for adult animals include: Carbon Dioxide (rodents only),
Isoflurane, barbiturates, physical methods performed under anesthesia
(cervical dislocation, exsanguination, decapitation), and MS222
(tricaine methane sulfonate) for aquatic species.  Physical methods of
euthanasia without anesthesia are conditionally acceptable and must be
scientifically justified and reviewed and approved by the IACUC. Personnel
responsible for performing these techniques must be properly trained and
the procedure will be reviewed by the ARC Director.

Carbon Dioxide Euthanasia
The NIH Office of Laboratory Animal Welfare (OLAW) issued clarification regarding the use
of carbon dioxide for euthanasia of small laboratory animals (July 17, 2002). The Animal
Care Unit of the Animal and Plant Health Inspection Service, U.S. Department of Agriculture,
has reviewed and has issued concurrence with the guidance provided by OLAW as it relates
to USDA covered species.

MSU Guidelines for using CO2 include:

All personnel using CO2 for rodent euthanasia must have received appropriate
training in its use. Training in the use of equipment and appropriate methods is
provided by the ARC.

The only acceptable source of CO2 is compressed gas cylinders. It is NOT acceptable to use
“dry ice” as a CO2 source.

Acute exposure of animals to high concentrations of CO2 is distressful and pre-filling the
chamber with CO2 gas is therefore not recommended. The euthanasia chamber should contain
only room air when placing the animals inside. The CO2 gas supply should then be turned
on so that CO2 concentrations gradually rise.

Chambers must not be overcrowded. In this regard, it is important to also consider that
mixing of incompatible animals in the same chamber may be distressful.

Death must be verified after euthanasia and prior to disposal. Unintended
recovery must be avoided by the use of appropriate CO2 concentrations
and exposure times or by other means. Animals subjected to CO2 euthanasia
should be observed for cessation of respiratory motions and apparent death
before removal for the chamber. It is best to administer CO2 for at
least 3-5 minutes and allow the animals to remain in the chamber for
an additional 3-5 minutes after the gas is turned off to determine that
respirations have not resumed. Death may also be verified by cessation
of a palpable heartbeat or ensured by performing cervical dislocation or
t horacotomy.

Fetal and Neonatal euthanasia (Rodents)
The AVMA Guidelines on Euthanasia (2007) do not provide specific
recommendations for the euthanasia of prenatal and neonatal animals. These
guidelines are based on NIH recommendations; Exceptions to these guidelines
will be considered by the IACUC on a case-by-case basis.

Rodent Fetuses:

Fetuses up to 14 days in gestation: Neural development at this stage is
minimal and pain perception is considered unlikely. Euthanasia of the mother
or removal of the fetus should ensure rapid death of the fetus due to non-viability
of fetuses at this stage of development.

Fetuses 15 days in gestation to birth: The literature on the development
of pain pathways suggests the possibility of pain perception at this time. Whereas
fetuses at this age are less sensitive to inhalant anesthetics, euthanasia may be
induced by the skillful injection of chemical anesthetics. Decapitation with surgical
scissors or cervical dislocation are acceptable physical methods of euthanasia.
When chemical fixation or rapid freezing (immersion in liquid nitrogen)
of the whole fetus is required, fetuses should be anesthetized prior to immersion
in or perfusion with fixative solutions. Anesthesia may be induced by hypothermia
of the fetus, by injection of the fetus with a chemical anesthetic, or by deep anesthesia
of the mother with a chemical agent that crosses the placenta, e.g., pentobarbital. The
veterinarian should be consulted for considerations of fetal sensitivity to specific
anesthetic agents. When fetuses are not required for study, the method chosen
for euthanasia of a pregnant mother must ensure rapid death of the fetus.

Rodent Neonates:

Up to 14 days of age: Acceptable methods for the euthanasia of neonatal
rodents include: injection of chemical anesthetics (e.g., pentobarbital),
decapitation, or cervical dislocation. Additionally, inhalant anesthetics
(e.g., isoflurane used with appropriate safety considerations), may be used.
However, neonates have a high tolerance for hypoxia, so exposure must be
prolonged (>20 minutes) and death confirmed by a secondary means. Pups
should be anesthetized prior to freezing with liquid nitrogen. Similarly, anesthesia
should precede immersion or perfusion with chemical fixatives. Anesthesia may
be induced by inhalant or injectable anesthetics; the veterinarian should be
consulted for appropriate agents and dosages.

Older than 14 days: Follow guidelines for adults.

Enrichment Policy
Various terms are used to describe the welfare requirements of research
animals, such as psychological well being, environmental enrichment, behavioral
needs, etc. The 1985 amendments to the Animal Welfare Act require facilities
to provide exercise for dogs and programs to promote the psychological
well being of nonhuman primates. The Guide for the Care and Use of
Laboratory Animals encourages “enriching the environment as
appropriate to the species”.

The ARC is responsible for the Enrichment Program. The mission is to promote
psychological well-being of all laboratory animals housed within the MSU
research program through appropriate housing, enrichment, health care
and activity, thereby allowing the animals to engage in species-typical
behaviors.  Methods of enrichment include, but are not limited to: social
housing; housing devices; manipulative devices; visual, auditory, and
olfactory devices; food items. The goals of the program are to address the
psychological well being of all species housed in the facility, provide interaction
and self-initiated behaviors by allowing a degree of control over their
environment, and to increase species-typical behaviors while decreasing
pathological maladaptive behaviors.

The complete and detailed program is reviewed and approved by the IACUC and is
available from the ARC.

Food and Water Restriction

Background:     Behavioral research often requires that an animal perform a task for which
it receives food or fluid reward. This situation is not unlike conditions in the wild, in which
animals must forage, travel distances, solve problems, or otherwise work to obtain food
and water. Performing a task for rewards may also be behaviorally enriching for laboratory
animals, especially nonhuman primates. However, a fundamental concern with studies
that may involve food or water restriction is that animals are maintained in a healthy state
and that they do not experience pain or distress. This document provides an overview
of the issues and explains the IACUC’s current policy regarding food or water restriction.

The IACUC referenced the following documents in setting this policy:

NRC (1996). Guide for the Care and Use of Laboratory Animals. Washington DC:
National Academy Press.

NRC (2003). Guidelines for the Care and Use of Mammals in Neurosciences and
Behavioral Research. Washington DC: National Academy Press.

The federal Animal Welfare Regulations (Code of Federal Regulations, Title 9).

Toth, LA and Gardiner TW. Food and Water Restriction Protocols:
Physiological and Behavioral Considerations. Contemporary Topics, 39(6). pg 9-17; 2000.

Definitions:

Standard for Food Intake: “Animals should be fed palatable,
non-contaminated, and nutritionally adequate food daily or according
to their particular requirements…..” (p. 38, “Guide for the Care and Use of
Laboratory Animals,” National Research Counsel, 1996.)

Standard for Water Intake: “Ordinarily, animals should have access to potable,
uncontaminated drinking water according to their particular requirements.”
(p. 40, “Guide for the Care and Use of Laboratory Animals,” National Research
Counsel, 1996.)

Restriction: Any deviation from the standards for food and water intake.

Fasting for surgical procedures: usually a period of less than 12 hours. This is
not considered restriction and the following guidelines do not apply.

Policy:

Individual animals may vary in their physiologic response to food or fluid restriction.
Therefore it is imperative that animals be closely monitored on a daily basis to ensure
that they are healthy, adapting normally and consume sufficient food and water to maintain
their health status. Monitoring and if necessary, early intervention are the most important
objectives of this policy.

It is the obligation of the investigator to demonstrate to the IACUC that food
or water restriction is the only effective way to accomplish the scientific goals of
the study. It is the investigator’s ongoing obligation to continue to attempt
methods of positive reinforcement that do not involve food or water restriction
and to use restriction only when other methods fail. The least restriction that will
achieve scientific objective should be used.

The use of restriction to motivate behavior must be specifically discussed, adequately
justified, and approved in each protocol in which it is used.

Criteria for monitoring animal health must be defined in the protocol (i.e. body weight,
urine and fecal output, urine specific gravity, etc.). Criteria must be defined for
temporary or permanent removal of an animal from the experimental protocol.

Investigators must maintain water consumption and food intake records in a form
acceptable to the IACUC and readily available for inspection.

At the discretion of the IACUC, Investigators may be required to file updates or reports
regarding the status of the animals on the restriction protocol.

At the discretion of the veterinary staff, a particular animal’s food or water ration must
be increased if:

The animal becomes ill or requires medical treatment.

The animal exhibits weight loss of greater than 10-15% of its initial body weight.

The animal shows evidence of clinical dehydration.

A young animal fails to show reasonable weight increase during a time when it
should be growing.

The animal undergoes any procedure requiring anesthesia.

Under any other circumstance deemed necessary by the veterinarian.

Humane Experimental Endpoints
The MSU IACUC is charged with evaluating and ensuring that research projects
are conducted in accordance with the Animal Welfare Act and the Public Health
Service (PHS) Policy on Humane Care and Use of Laboratory Animals. These
regulations and policies require that any research, testing and teaching that
uses animals must be performed in such a way as to minimize discomfort,
distress and pain consistent with sound research design.

The IACUC recognizes that as a sequela of research or testing, laboratory
animals may experience pain and/or distress. For certain studies the procedures
may lead to the moribund condition or death. Therefore, the IACUC has developed
guidelines for establishing humane experimental endpoints to assist investigators
with the process of completing the UIC Protocol for Animal Use and to insure
that animal welfare issues are appropriately addressed. Since the diversity
of research and testing protocols preclude the establishment of a rigid set of
guidelines, the ACC reserves the right to evaluate each protocol on an
individual basis.

Definitions

Humane Endpoint - The humane endpoint is the point at which pain and/or distress
is terminated, minimized or reduced by taking actions such as euthanatizing the animal;
terminating a painful procedure; or giving treatment to relieve pain and/or distress.

Moribund Condition - The moribund condition is a severely debilitated state that precedes
imminent death.

Investigators should identify humane experimental endpoints that minimize pain,
distress, or discomfort by choosing the earliest endpoint that is compatible with the
scientific objectives of the research project.
Whenever possible, the moribund condition should be used as an experimental
endpoint instead of death, because it is assumed that euthanatizing a moribund animal will
avoid or reduce terminal distress.
The following specific sequela should also be considered:

The procedure should not interfere with the ability of the animal to ambulate, eat, drink,
urinate and defecate.

The procedure should not result in a net weight loss of more than 20% of the
body weight.

The procedure should be ended if the investigator has conclusive evidence that
untreatable organ failure has occurred, and the animal exhibits signs associated
with the failure of the organ system.

Animals should be euthanatized if unrelated health conditions develop
that make their experimental use of no value to the investigator.

Obvious signs of illness should serve as alternatives to death as an experimental
endpoint.

The use of death as an endpoint requires strong scientific justification
in the Protocol and full committee review and approval by the IACUC.

Maximum Dosing Volumes

Non-terminal Blood Withdrawal

These stated volumes are the maximum blood to be withdrawn in a single bleed. At maximum,
it can be repeated at two-week intervals. If more frequent bleeding is needed, the volume
should be proportionally reduced. If more blood than this maximum volume is needed,
the investigator must consult with the Veterinarian and receive IACUC approval.

Polyclonal Antibody Production
The production of antibodies to specific and nonspecific antigens is a tool utilized in
nearly all fields of biomedical research. Today antibodies are routinely made to a
vast array of proteins, carbohydrates, fats, and nucleic acids. In addition modern
biochemical, biosynthetic, and recombinant techniques have created increasingly pure
antigens, both natural and synthetic, in an effort to induce very specific immune responses.
Many of these newer antigens are small and generally weak immunogens and it is
necessary to augment their immunogenicity with adjuvants. One of these antigens
is Freund’s Complete Antigen (FCA).

The NIH has published recommendations for the use of FCA. In these
recommendations NIH stated that FCA is an "important biologic resource for
investigators which should be used responsibly and with care to avoid or minimize
the adverse effects of excessive inflammation." Recommendations are included on
dosages and locations with specific mention of using caution
with footpad and peritoneal inoculation. In addition, they recommend
that the use of noninflammatory or less inflammatory adjuvants be considered
when possible. Studies subsequent to the publication of the NIH Recommendations
for the use of FCA indicate that FCA, when administered appropriately, may be
pursued humanely in the production of polyclonal antibodies.

The MSU IACUC recognizes that FCA and other commercial adjuvants used in the
production of polyclonal antibodies can cause granulomatous/inflammatory reactions.
The IACUC also recognizes that FCA and other commercial adjuvants can be
pursued humanely to produce polyclonal antibodies when administered in an
appropriate manner by trained individuals familiar with the procedures. The
following guidelines for the production of polyclonal antibodies in rabbits were
developed to ensure consistency of technique and to minimize
pain and distress.

All procedures related to production of polyclonal antibodies in rabbits using
FCA will be performed by the ARC staff.

Commercial adjuvants should be administered in a manner consistent with
the manufacturer’s recommendations.

It is recommended that FCA only be administered via the subcutaneous route using
a 25 gauge needle. It should be administered over multiple sites with a maximum of
0.1ml dose at each site.

Gloves and lab coat should be worn when working with FCA. Individuals should be vigilant
to prevent self-inoculation, which has been associated with granulamatous reactions in
individuals that result in tuberculin skin test positive reactions.

Booster injections should be done with Freund’s Incomplete Antigen (FIA). Animals should
not undergo more than four booster immunizations during the initial phase of antibody
production.

Booster immunizations with FIA should be administered in the same manner as FCA. It is
important that subsequent immunizations be administered in a different location than the
previous immunizations.

Since some antigens are powerful immunomodulators, which can augment the reactive nature
of FCA and commercial adjuvants, all animals administered adjuvants for production of
polyclonal antibodies should be observed on a regular basis.

The administration of adjuvants for the production of polyclonal antibodies should only be
performed by experienced individuals.

The Use of Complete Freund's Adjuvant in rabbits

Complete Freund’s adjuvant (CFA) is widely used and considered to be the most
effective adjuvants available for consistently producing high titer antibodies
to diverse antigens. It is irreplaceable and vital for immunology research
and antibody production at the present time. The Institutional Animal Care and Use
Committee (IACUC) has developed the following recommendations to permit the
continued use of CFA while maximizing antibody titer and minimizing the adverse
effects of excessive inflammation, arthritis and other
toxic effects.
CFA is a water-in-oil emulsion containing mycobacterial cell wall components that
potentiate the humoral antibody response to injected immunogens. Adjuvant activity
results from sustained release of antigen from the oily deposit and stimulation of a local
immune response. The improper or unnecessary use of CFA may cause inflammation,
induration, and/or necrosis in laboratory animals.
The intention of the following guidelines is to minimize potential animal discomfort
associated with the use of adjuvants in research. Unless specifically approved
by the IACUC, antibody production using CFA at Montana State University must be done using
these guidelines. Requests for deviations must be scientifically justified and will be
considered on a case-by-case basis by the IACUC at the time of protocol review or in
response to a request for protocol modification.

Alternatives which reduce the number of animals (tissue cultures, chicken eggs, etc.) or
utilize non-inflammatory adjuvants (Ribi, liposomes, incomplete Freund’s) must be considered.

CFA should be used only when necessary, such as small amounts of available antigen and
weak antigens. The USDA has declared that the use of CFA may cause more than
momentary pain or distress and an alternative must be considered. A written narrative
description of the methods and sources used to search for alternatives must be provided
in the animal care and use protocol. Key words should include CFA and alternatives.

CFA may be used only for the first (priming) dose. Subsequent immunizations should
be with incomplete Freund’s or another adjuvant.

The inoculums must be free of extraneous microbial or other particulate contamination.
When possible, sterilization by filtration through a low binding 0.22 micron filter should
be performed.

Young adult rabbits (2.5-3.5 kg) should be used for primary immunization because of the
vigorous antibody response. Immune function peaks at puberty and primary responses
to new antigens decline with age.

Multiple injection sites must be separated from each other widely enough to insure
continued blood supply. Anatomic sites which are weight bearing or used in restraint
should be avoided (neck area in rabbits). It is strongly recommended to use sites close
to the draining lymph nodes (ventral inguinal and axillary regions).

Fur must be clipped and the site aseptically prepared with the appropriate antiseptics.

Injections should be given subcutaneously. Any other route of administration requires
additional scientific justification for IACUC approval.

Injections must be divided into fractions so that no more than 0.1 ml is injected per site,
with no more than a total volume of 1 ml (10 sites).

Injection sites must be observed at least three times a week for four weeks after each
injection. This should be properly documented in the animal record. If a lesion develops
at any injection site, the Veterinarian must be notified so the animal can receive the
appropriate treatment.

Stabilization Period after Animal Transport
All new animal arrivals, including mice, rats and other rodents, must undergo a minimum
48-hour stabilization period before experimental use. No component of the study may be
initiated during this stabilization period. The stress associated with transportation has long
been recognized to have profound physiological consequences. It is important to provide
an adequate period before experimentation to allow animals to recover to a physiologically
stable condition. This stabilization enhances both the validity of the experimental results
and animal welfare.

Tail Snips
Rodents that are over 21 days of age must be anesthetized for a tail snip (amputation)
procedure. Hemostasis must be ensured before the animal is returned to its cage.

Weight Loss
If weight loss reaches 15% of original body weight, the Veterinarian should be
consulted to determine a course of action regarding the animal. In most cases the
active use of the animal on study is suspended so the animal’s condition can be improved.

Surgery and Anesthesia Policies

Anesthetic Cocktail Expiration
Anesthetic Cocktail mixtures should be dated at the time of mixing and
will be considered to expire 2 weeks later.

Multiple Major Survival Surgeries
Multiple major survival surgical procedures can be justified if they are
related components of a research project, if they will conserve scarce animal
resources, or if they are needed for clinical reasons. It is unacceptable to subject
an animal to multiple unrelated survival
surgeries simply to save money or because the animals are in short supply.

Pain Management
Investigators are required to administer appropriate analgesics to all animals
associated with a procedure that would normally require pain medication in humans. 
Questions concerning appropriate medication and dosage for a given species should
be directed to Veterinarian.  Exemption from this regulation is possible only on
a protocol–by-protocol basis after Full IACUC review and approval.

Rodent Surgical Guidelines

Pre-Operative

Surgery should be conducted in a disinfected, uncluttered area, which promotes
asepsis during surgery.

Prepare the animal by removing hair from the surgical site. Perform this procedure
in an area separate from where the surgery is to be conducted.

 Prepare the surgical site(s) with an appropriate skin disinfectant.

Surgeons should wash and dry their hands before aseptically donning sterile
surgical gloves.

Operative

The animal must be maintained in a surgical plane of anesthesia throughout
the procedure.

Begin surgery with sterile instruments (autoclave, chemical sterilant, ethylene oxide)
for each animal and handle them aseptically. Glass bead sterilizers may be used to
sterilize the tips of instruments between animals or procedures.

Instruments and gloves may be used for a series of similar surgeries provided they are
maintained clean and disinfected between animals.

Monitor and/or maintain the animal's vital signs.

Close surgical wounds using appropriate techniques and materials.

Post-Operative

Move the animal to a warm, dry area and monitor it during recovery. Return the animal
to its routine housing only after it has fully recovered from anesthesia.

Provide analgesics as appropriate.

Generally, remove skin closures 10 to 14 days post-operatively.

Maintain a surgical record (e.g., annotate cage card with procedure and date).

Survival Surgery
Regulations pertaining to survival surgery are very specific and include the
following definitions:

Survival Surgery includes all surgical procedures (even very minor ones
such as vessel cut-downs) carried out on animals that are expected to recover
from the anesthetic.

Major Survival Surgery is defined as survival surgery in which a major
body cavity (abdomen, thorax, cranial cavity) is entered or which has the
potential for causing substantial impairment of physical or physiologic function.

All survival surgery must be carried out following aseptic technique,
including the use of sterile gloves and properly sterilized instruments.
  This applies to all species including rats and mice. Major survival surgery on
mammals other than rodents must be carried out in dedicated survival surgery
facilities (ARC Room 111).  Major survival surgery on rodents must be done
according to the Guidelines for Survival Rodent Surgery.