Laboratory Animal Care Service
Special Division of Biology policies and guidelines

Currently, the Kansas State University IACUC has adopted a series of special guidelines that may affect the experimental design of an investigators research. These guidelines include such procedures as types if animal restraint, aseptic surgery techniques, tail clipping, use of ether for anesthesia, multiple surgery procedures, food and water restrictions, euthanasia of neonates, use of Freund's complete adjuvant, and use of death as an endpoint. Investigators must justify, within the body of their IACUC application, any reasons why an experimental procedure should deviate from the approved guidelines.

In addition to IACUC guidelines, the Division of Biology has approved similar and sometimes additional guidelines affecting use of animals by Biology faculty and staff. These are presented below (NOTE: Whenever a Division of Biology guideline conflicts with an IACUC guideline, the more restrictive of the two guidelines applies):

Policy on post-surgical medical care procedures

Animals will be provided postsurgical care according to current professional and regulatory guidelines. These policies have been endorsed by the University's IACUC, and failure to completely and properly monitor animals during the postsurgical recovery period will result in the cancellation of previously approved protocols.

During the postsurgical period, a postsurgical record must be kept in the room where the animal is housed or in an adjacent area. It should include a brief description of the surgical procedure, anesthetic used, time of induction, duration of surgical anesthesia and time returned to cage. It should also include the findings of each physical examination during the recovery period. The postsurgical medical record will be retained as a part of the animal's permanent medical record. It is best to keep experimental notations in a separate location.

For purposes of medical monitoring, it is helpful to stage animals according to extent of recovery from surgery and anesthesia. The animal should be examined and the finding recorded according to the following schedule:

1. Stage 4 - Animal unconscious or semiconscious and unable to sit or maintain sternal recumbency.
   1. Examine and record findings no less frequently than every 2 hours. More frequent examination is recommended. Examples of notations include: body temperature, heart rate, respiratory rate capillary refill time (record in seconds), jaw tone (record resistance or no resistance), response to toe pinch (record withdrawal or no withdrawal, and time of extubation.
   2. The animal should be turned from side to side frequently to prevent dependent pulmonary congestion and edema.
   3. Ambient temperature should be adjusted (heat lamp or warming board) to bring body temperature to normal. Care must be taken not to burn or overheat animal at this stage of recovery. The animal should be kept dry.
   4. The state of hydration should be assessed and fluids should be provided as necessary.
2. Stage 3 - Animal conscious and can maintain sternal recumbency to sit, but cannot stand.
   1. Examine and record findings no less frequently than every 6 to 10 hours depending on the nature of the surgery and the status of the animal. Examples of notations include: body temperature, until it becomes normal within 2 F; capillary refill time, and condition of the operative site.
   2. Examine closely for other abnormalities.
   3. Keep the animal dry and adjust the ambient temperature to bring the body temperature to normal.
   4. Consider use of analgesic medication
   5. Professional judgement should be exercised in cases in which there is difficulty in making examinations every 6-10 hours. We recommend caution in prolonging examinations of animals in stage 3.
3. Stage 2 - Animal can stand and move about but is not eating and drinking normally
   1. Examine daily and record findings. Examples of notations include: body temperature, hydration, attitude (alert or depressed), activity (active or inactive), food consumption, water consumption, and condition of operative site.
   2. Examine closely for other abnormalities.
   3. Consider use of analgesic medication
4. Stage 1 - Animal active, alert, eating and drinking normally; skin sutures are in place
   1. Examine daily and keep a postsurgical record or surgical site care until sutures are removed.
   2. Sutures to be removed within 10-14 days of surgery.
5. Stage 0 - Animal normal and skin sutures removed (specific postsurgical care and record no longer required)

If progress from Stage 4 to Stage 0 is not as expected or if medical complications occur, the veterinary staff is available (2-5640, 2-5648). The veterinarian on call during weekends and holidays can be reached by beeper (532-5000, 031). The Department of Diagnostic Medicine/Pathobiology can provide clinical laboratory support.

This staging system is designed for investigators utilizing rabbits, carnivores, and hoofed stock. Although rodents are exempt from this organized scheme, it is imperative that these species also receive adequate postsurgical care. Rodents are poor thermo-regulators, so special care must be taken to ensure that their body temperature is maintained during the postsurgical period. Antibiotic ointments should be applied to all wound sites to assist in preventing postsurgical infections.

Policy on decapitation and cervical dislocation as a form of euthanasia

Appropriate methods of euthanasia are an important component of an effective Animal Care and Use Committee program. In most cases we defer to the policies described in the 1993 report of the AVMA panel on euthanasia. However, we believe that the panel's findings are inappropriate in dealing with the use of decapitation as a method of euthanasia. Decapitation without anesthesia shall be considered an appropriate form of euthanasia for small mammals (including mice, rats and guinea pigs) and small rabbits in those situations in which the investigator has stated that the use of anesthetics would obscure the dependent measures of interest. Cervical dislocation is appropriate for euthanasia of avian species, mice and immature rats and rabbits.

To conform to our policy, decapitation and cervical dislocation should be scientifically justified and carried out by trained personnel. Decapitation does require some manipulative skills and experience to be with assurance that the investigators are trained in using this procedure. When a Principal Investigator proposes to euthanize research animals by cervical dislocation or decapitation, a rationale is needed for using these procedures over other, more preferred procedures (e.g., overdose with appropriate anesthetic).

Procedures should be executed as rapidly and painlessly as possible. Every attempt should be made to perform either of the methods of euthanasia on animals so that reactions among other living animals are minimized (i.e., performed in a room separate from the housing area). Decapitation and cervical dislocation should be performed after the animal has been lightly anesthetized, unless the anesthetic would obscure the dependent measures of interest.

A guillotine is recommended for decapitation of avian species, rodents and small rabbits. A sharp scissors is permissible to use for decapitation of small animals such as neonatal rodents and small mice. The instrument used in a decapitation procedure must be properly designed to insure decapitation in a uniformly instantaneous manner, and it should be cleaned free of blood and tissue debris between each successive use.

Appropriate members of the Animal Care and Use Committee of Kansas State University are available to discuss specific experimental requirements that do not strictly conform to these recommendations.

Policy concerning care and use of atypical species

As stated in the NIH Guide for the Care and Use of Laboratory Animals, the guidelines apply to ...any warm-blooded vertebrate animal used in research, testing, and education. Although marine mammals and cold-blooded animals are not discussed specifically, the humane principles stated are applicable to their care and use.

Thus, there seems to be no escaping the conclusion that, though atypical or wild-caught animals are not dealt with specifically, the guidelines apply to them as well. However, the guidelines are very general in nature, and not always rigid. As stated in italics in the Guide Professional judgment is essential to the application of these guidelines. It is our recommendation, therefore, that each protocol and program that uses atypical species be dealt with individually. It may be necessary to seek ad hoc advice. Since the Animal Welfare Act is law, all regulations and standards must be met.

Policy on photography in animal facilities, research and teaching laboratories

Kansas State University maintains an open policy regarding reasonable requests to visit its animal facilities, research and teaching laboratories; however, to protect the confidentiality of faculty research, provide a minimally disruptive atmosphere for animal residents and guard against the misinterpretation of appropriate and humane policies and procedures, photography is not allowed except for official purposes that are approved as such by the IACUC.

Policy on videotapes of animal research

The IACUC asks that all faculty who have or will have made videotapes of their research animals take the following precautions immediately:

1. Store tapes in a secure, locked area with very limited access.
2. Do not authorize copies of tapes unless supervised by the Principal Investigator personally.
3. Erase immediately all tapes no longer in use or of value to the research.
4. Keep a checkout list including dates and circumstances of persons borrowing tapes. Even under these circumstances, access should be very limited and closely supervised.

Policy on use of Freund's (complete) adjuvant in laboratory animals

The IACUC and the LACS have developed the following guidelines for the use of Freund's Adjuvant in an attempt to eliminate, or reduce to a minimum, animal discomfort associated with the use of this agent. Departure from these guidelines requires written scientific justification to the IACUC.

Freund's Complete and Incomplete Adjuvant causes local inflammation at the site of injection, often resulting in painful lesions. Before using these or other inflammatory adjuvants, please consider the use of non-inflammatory alternatives such as ethylene-vinyl acetate copolymer, ribi adjuvant system, muramyl dipeptide and liposomes. In many cases, non-inflammatory adjuvants have been shown to produce antibody titers comparable to those induced by inflammatory adjuvants.

General guidelines

1. Freund's Complete Adjuvant (FCA) should only be used for the first (primary) dose. Subsequent doses of antigen-adjuvant complex should be made using Freund's Incomplete Adjuvant (FIA).
2. The ratio of antigen to adjuvant should be 1:1. If FCA is used, the concentration of mycobacteria should be 0.1 mg of dry bacterial mass/ml or less. The use of formulations containing higher concentrations of mycobacterium has been associated with proportionally increased amounts of inflammation and necrosis. Commercial preparations containing more than the acceptable number of mycobacteria per ml can be diluted to the approved concentration with FIA.
3. Acceptable routes of administration include subcutaneous and intramuscular injection in all species and intraperitoneal administration in mice. Any other route of administration requires written scientific justification to the IACUC. In all species, intradermal injection of FCA or FIA has been associated with a high incidence of necrotizing dermatitis and tissue sloughing. Intradermal injection of Freund's Adjuvant must be justified to the IACUC. The use of FCA or FIA in the footpad or toepad of rabbits and rodents has been associated with a high degree of pain and distress. Footpad or toepad injection of Freund's Adjuvant must be justified to the ACUC. FCA should never be given intravenously. Acceptable routes of administration are listed for each species (see below).
4. The volume of antigen-adjuvant complex injected is a major determinant of the degree of inflammation and pain that results when FCA and FIA are used in animals. The appropriate volume varies with the species used and the route of administration.
5. Whenever possible, the sites of injection should be widely scattered to prevent individual lesions from coalescing. This often results in disruption of local blood supply and contributes to cell death and tissue sloughing.
6. The interval between first, second and subsequent doses of antigen-adjuvant complex should be determined by the antibody response of the animal but should be maximized whenever possible. If there is pain and discomfort following injections, the interval between subsequent injections should be long enough to allow a recovery period.
7. The injection site must be observed by the investigator or his/her designate a minimum of three times/week, for four weeks after each injection. If a lesion develops at the injection site, it must be reported through established channels (e.g., LACS personnel) and receive appropriate veterinary care. Such lesions must be inspected at least three times per week by the investigator or his/her designate, until all lesions are healed.

Species specific guidelines

1. Mice and other small rodents: Approved routes and volumes (maximum volume/site): Subcutaneous, in neck region, 0.10 ml/site; Intramuscular, in biceps femoris muscle, 0.05 ml/site; Intraperitoneal, 0.10 ml/injection
2. Rats: Approved routes and volumes (maximum volume/site): Subcutaneous, in neck region, 0.10 ml/site; Intramuscular, in biceps femoris muscle, 0.05 ml/site
3. Guinea pigs: Approved routes and volumes (maximum volume/site): Subcutaneous, in neck region, 0.40 ml/site; Intramuscular, in biceps femoris muscle, 0.10 ml/site

Policy on use of hypothermia as a method of anesthesia

Hypothermia is an acceptable form of anesthesia for the preweanling animal (< 2 weeks of age) listed below:

1. Rodents
2. Rabbits
3. Ferrets
4. Marsupials (including opossums)

Three alternative procedures are acceptable for inducing anesthesia:

1. The preweanling animal is placed directly on a bed of crushed ice (NOTE: this may be complicated by skin freezing at the point of contact with ice that is below 0 C; care must be taken).
2. The animal is placed inside a glass test tube that is immersed in ice (NOTE: the time of onset of lethargy may be considerably slower than in the other methods).
3. The animal is immersed in ice water, with care being taken to avoid water aspiration (NOTE: there is a risk of drowning; care must be taken).

Postoperatively, the animals are warmed to normothermia by placing them in an incubator at 33 C. Placing the animal on a heating pad or under a heat lamp is strongly discouraged, as it may lead to tissue damage. The animal should be mobile and able to return to its cage in 20-30 minutes.

REFERENCE
Phifer CV, Terry LM. Use of hypothermia for general anesthesia in preweanling rodents. Physiological Behavior. 1986;38:887-890.

Policy on open-drop anesthesia of rodents

The open-drop (anesthetic chamber) method for inducing anesthesia in laboratory rodents has been approved by the Institutional Animal Care and Use Committee under the following conditions:

1. The closed anesthetic chamber must contain a raised floor to protect the animal from direct contact with the anesthetic agent, which is usually soaked in gauze.
2. The chamber must be used in a chemical fume hood to exhaust the anesthetic vapors.
3. The use of ether is discouraged because of its explosive nature, but if chosen as an anesthetic, it must be used and stored in a chemical fume hood.

Policy on use of live vertebrate animals in the classroom

Statement of policy.

The Division of biology at Kansas State University recognizes the importance of using live animals in some classroom demonstrations and teaching laboratories, particularly classes containing high numbers of biology, microbiology, and/or wildlife majors. The use of live animals in teaching is a common and acceptable educational practice, and is considered vital in some courses for students to understand certain biological processes. Specific examples include, but are not restricted to, physiology, behavior, immunology, parasitology, virology, bacteriology, developmental biology and embryology, evolutionary biology, and ecology. In the majority of cases, the instructor for a course will be responsible for deciding whether the use of live animals in teaching is warranted. The Division of Biology Laboratory Animal Care Service (LACS), or Animal Resources Facility (ARF), will be responsible for providing any husbandry required for animals used in the classroom.

Applicability of Federal Guidelines.

It is University policy that procurement, housing, care, use, and euthanasia of animals used in teaching conform to the NIH Guide for the Care and Use of Laboratory Animals, to all standards set forth by the US Department of Agriculture, to the Animal Welfare Act (7 USC, 2131-2156), and to the recommendations of the 1993 report of the American Veterinary Medical Association panel on euthanasia.

Division of Biology Guidelines.

The principle investigator is the faculty member in charge of all animal use associated with the teaching project. The faculty member is responsible for assuring the following:

1. That the use of live animals in the classroom, or direct manipulation of live animals during laboratory field trips, has prior approval of the Instutional Animal Care and Use Committee. The appropriate class handouts, manuals, etc. that directly relate to use of the animals in the classroom will accompany the IACUC application.
2. Acquiring any and all federal and/or state collecting permits that might be required prior to use of animals in the classroom.
3. Acquiring any and all federal Drug Enforcement Agency (DEA) permits, or State of Kansas licenses, necessary for the administration of any regulated drugs.
4. Insuring that all students are properly educated, equipped, and trained to reduce exposure to animal bites and potential zoonosis during classroom, laboratory, and field exercises, and that students receive adequate instruction in emergency procedures in case of animal bites or direct exposure to known zoonoses.
5. Assuring that appropriate safeguards are in place so that students, staff, faculty, and the general public are not directly exposed to any venomous, infective, or otherwise dangerous animals in the classroom or laboratory, and that such live animals are properly housed and/or restrained during the classroom session.
6. Assuring that all animals are treated humanely in the classroom or laboratory, and that the fewest numbers of animals are used to achieve the desired class objectives.
7. Assuring that videotapes or simulations are used in place of live animals whenever surrogates result in achieving the desired level of education. In general, surrogates are commonly used in place of live animals in freshman and sophomore courses that may contain high numbers of non-biology majors.
8. Assuring that a sufficient number of properly educated, equipped, and trained instructors or assistants are present to insure humane use of animals in the classroom.
9. Alerting students early in the course about the use of live vertebrate animals in the classroom or laboratory, and for assuring that procedures are in place to deal with conflicts that arise with students who may disagree about how and why animals are being used. A mechanism should be in place so that individuals in non-major courses are not unjustly penalized or prevented from graduating because of their beliefs.

Kansas State University | Laboratory Animal Care Service