Guidelines for Rodent Surgery

Rodent Surgery Guide

Case Western Reserve University
Institutional Animal Care and Use Committee
Updated 7/20/99

See also the NIH Guidelines for Rodent Surgery on the IACUC web site.

These guidelines are taken verbatim from the IACUC web site at Guidelines for Rodent Surgery

Recommendations for the performance of rodent surgery are based on the 1996 edition of the NIH "Guide for the Care and Use of Laboratory Animals" and 9 CFR, the Animal Welfare Act (AWA). Part 2 of the AWA states that major surgical procedures on rodents "must be performed using aseptic procedures." This would include the use of sterile instruments, sterile surgical gloves, and aseptic preparation of the surgical site in order to prevent postoperative infections. A separate facility for rodent surgery is not necessary. A rodent surgical area can be a room or portion of a room that is easily sanitized and not used for any other purpose during the time of the surgery. While dedicated facilities are not required for rodent survival surgery, the use of sterile instruments, surgical gloves, and aseptic procedures are required. General requirements are given on pages 60-65 of the NIH guide (National Academy Press 1996).

Rodents include hamsters, gerbils and guinea pigs, as well as rats and mice. Guinea pigs and hamsters are USDA covered species, meaning that they are not exempt from USDA regulations and the provisions of the AWA.

Three things that are extremely important to maintaining a sterile procedure are the reduction of time, trauma and trash:

1. Time of surgical procedure is an important factor, as the longer a procedure takes the greater the possibility of contamination and therefore infection.
2. Trauma that is sustained by the tissue as a result of rough handling, drying out upon exposure to room air, excessive dead space, implants or foreign bodies or non-optimal temperatures will contribute to infections.
3. Trash refers to contamination by bacteria or foreign matter.

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 physiologic 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 groups of rats, differences were observed in open field behavior, 'freezing' behavior, plasma fibrinogen, serum glucose, total white cell count, and wound histology scores. Activation of macrophages in response to intraperitoneal inoculation of bacteria, stimulation of cytokines and activation of B cells by bacterial endotoxins (lipopolysaccharides) and alterations of other physiological processes by subclinical viral, mycoplasmal, bacterial or parasitological infections are well documented in the literature.

Rodent surgery can be classified as minor or major in nature.

PART I- MINOR SURGERY

"Minor survival surgery does not expose a body cavity and causes little or no physical impairment" (the "Guide," p 63) and includes injections, vena-puncture, and subcutaneous implants. When conducted with proper care, these techniques present few difficulties. "Minor procedures are often performed under less stringent conditions than major procedures but still require aseptic technique and instruments and appropriate anesthesia." (The"Guide," p 62)

The implanting of a chronic intravenous catheter is intermediate in nature, but is the technique that presents the most severe postsurgical infections. Because one is opening a direct venous access, surgical technique needs to be meticulous, as for major surgery. Postsurgically, use sterile technique when accessing the catheter (s). The most critical requirement is to inject only sterile solutions into the catheter. Solutions should be freshly prepared or stored under refrigeration if prepared in advance. The top of the vial or mouth of the container containing solutions for injection must be kept clean and wiped with alcohol, or flamed, before drawing up the solution. Inoculation of even a few organisms into an intravenous catheter may result in death of the animal due to sepsis.

PART II- MAJOR SURGERY

Major surgery includes invasion of the cranial, abdominal, or thoracic cavities. Any procedure that might leave the rodent with a permanent handicap, whether physical or physiological, would also be considered major surgery. The use of aseptic technique is mandatory in these surgeries to minimize the possibility of postsurgical infection. Consultation with an attending animal veterinarian or veterinary technician is recommended if you have questions regarding techniques appropriate for these situations.

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 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. Before beginning rodent surgery, the laboratory bench or table where the surgery will be performed should be cleaned and disinfected.

Surgical instruments must be sterile. Heat sterilization is ideal. Agents such as chlorine dioxide or gluteraldehydes can be used for cold sterilization. Chlorine dioxide is not documented as being toxic to animal tissue but will corrode stainless steel instruments. Glutaraldehyde must be thoroughly rinsed off of instruments with sterile saline or water before use of delicate items, such as drills and burrs. Catheters and implants can be sterilized using ethylene oxide or ionizing radiation (see attached Table 2).

A pre-surgical evaluation should be performed to insure that your prospective patients are not overtly ill. Is the animal alert with a smooth coat and clear eyes? The withholding of food is not necessary in rodents unless specifically mandated by the protocol or surgical procedure. Water should NOT be withheld unless required by the protocol. Withholding of food for more than six hours should be discussed with a veterinarian.

Preparation of the animal should include clipping, shaving or plucking the hair from the surgical site with enough border to keep hair from contaminating the incision (hair removal should be performed in a location remote from the surgical area). The surgical site should be scrubbed at least twice with a germicidal scrub (see attached Table 3), being careful to scrub from the center of the site toward the periphery. The site can then be rinsed with a 70% alcohol or dilute iodine solution. Note that alcohol will contribute to hypothermia if liberally used. Finally, the area should be draped with sterile drapes which not only helps prevent stray hair from entering the surgical field, but provides a sterile area on which to lay sterile instruments during surgery.

Preparation of the surgeon: The surgeon must thoroughly scrub his or her hands with a bactericidal scrub (see attached Table 3). The use of sterile surgical gloves is necessary if the surgeon's fingers will be touching the surgical site. A surgical mask should be worn for major surgeries. The wearing of a clean lab coat is mandatory. A sterile gown is preferable for major surgeries.

Use of surgical instruments on more than one animal: Surgical instruments must be steam or gas sterilized prior to use. If care is used to maintain asepsis of surgical instruments, they may be used for a maximum of five animals. However, chemical or heat disinfection of instruments between animals is preferred. The purchase of a hot bead sterilizer for this purposed is recommended. Any item used on multiple animals must be carefully cleaned and disinfected between animals (see attached Table 4). Alternating two or more sets of instruments is one way to allow time for instruments to sit in a disinfectant or sterilant solution for more than just a few minutes.

The "No Touch" technique is useful for mouse surgery. It may also be used to advantage with certain rat procedures such as stereotaxic placement of brain electrodes or cannulas. With this method, the hands never touch the incision; only instrument tips enter the surgical field. The use of nonsterile gloves is acceptable and a hot bead sterilizer is used to maintain the sterility of the working end of the surgical instruments. Care must be taken to replace instruments on the instrument tray or drape in such a manner than the asepsis of their tips is maintained. Tissue and wound clips must be used with the "No Touch" technique as the sterility of suture material cannot be reliably maintained.

Evaluation of the animal during surgery is critical. Monitoring of anesthetic depth is usually of first importance. Unfortunately, techniques for monitoring anesthetic depth vary somewhat with the agent used. A quiet animal that does not move when a painful stimulus is applied is the most certain indicator of adequate anesthesia, however, the zone between quiet and too quiet is very narrow in rodents.

Maintaining body temperature is next in importance. A warm water blanket or hot water bottles provides supplementary warmth without being too hot. Bubble wrap may help a small rodent maintain body temperature. During long surgeries, warmed sterile fluids (saline or lactated Ringers solution) should be provided. These can be administered subcutaneously, intravenously or intraperitoneally. Any tissues exposed for very long during surgery should be kept moist with these same warmed solutions.

Observation during postsurgical recovery is important. The animal, in or out of its cage, must be kept warm. Warm water pads, blankets, or the blue "diaper" pads work well. The use of electric heat pads or heat lamps may overheat the animal; their use is discouraged. If electric heat pads or heat lamps must be used, provision must be made to make frequent observations and turning of a somnolent animal so that the animal will not be overheated. Provision must also be made so that an awake animal can escape the heat source when it becomes too warm. Warmed fluids can be administered subcutaneously, intravenously, or intraperitoneally if there is any suspicion the animal may be dehydrated. (over hydration is not generally a problem in animals with normal kidney function). This may be done by giving 1 to 2 ml of warm fluids (0.95% NaCl or equivalent) per 100 gm of body weight by subcutaneous injection. If blood loss occurred during the surgical procedure, or if the animal is slow to recover from anesthesia, provide additional fluids.

A recovering animal should be watched very closely until securely in sternal recumbency, and able to move around without plugging its nostrils with bedding. Some rodents left overnight on pads or paper bedding will eat that bedding.

Suture and staple removal: Post-surgical animals should be seen every day by a member of the investigator's staff or other individual to whom post-operative care has been delegated. Postsurgical observations include a minimum daily observation of the condition of the animal and the surgical site. Animals should be observed daily until all wounds have been healed and sutures or wound clips are removed. Sutures and/or staples must be removed by two weeks following surgery, if the rodent has not already done so. Any foreign substance left in the incision for long period of time serves as a nidus of irritation and infection. Incisions that do not appear to be healing should be examined by a veterinarian.

A postoperative record should be completed and affixed to the animal's cage card. Having the record in the room accomplishes several functions. 1) It explains the condition of the animals to animal care staff (a sedated animal may otherwise be thought to be ill), 2) It assures animal care staff and USDA Animal Welfare inspectors that the animal is being cared for, and 3) It informs animal care staff how recently the investigator has seen the animal; this knowledge helps them decide whether or not there is a need to contact the investigator to inform him or her of the present condition of the animal. Although individual records are desirable, USDA allows a composite post-operative record to be used for a group of rodents.

Techniques which are important and often difficult to perfect are the following:

• Touch only "prepped" areas with sterile instruments and sterile gloved hands.
• Keep operating fields draped.
• Do not let catheters or implants become contaminated.
• Use sterile solutions.
• Disinfect the tops of containers of solutions.
• Use sterile technique to access implanted catheters.

Not only are the above recommendations more humane, but following these recommendations will improve one's research by providing a less stressed animal and thereby decreasing the number of variables in a research protocol. The rat, especially, has always been considered "hardy" and not subject to postsurgical infections. Published research has documented that postsurgical infections in rats are subtle. The rat appears to eat and act normally, but will not respond appropriately to research stimuli. As with all new and improved techniques, patience and practice are required to harvest full benefits from the use of aseptic surgical techniques in rodents.

There is ample literature available supporting the recommendations presented in this document. The attending and /or clinical laboratory animal veterinarian or veterinary technician is available for assistance or to provide referrals to other researchers with applicable knowledge or skills.

Table 1. Recommended Hard Surface Disinfectants

(e.g., table tops, equipment) Always follow manufacturer's instructions.

A disinfectant is a germicidal chemical substance that kills microorganisms on inanimate objects, such as instruments and other equipment, that cannot be exposed to heat. Disinfectants are essential for good housekeeping in hospitals and clinics, where they are used on floors, cages, tables, and counter tops.

^* The use of common brand names as examples does not indicate a product endorsement.

Table 2. Recommended Instrument Sterilants

Always follow manufacturer's instructions.

Sterilization is the complete elimination of microbial viability, including both the vegetative and spore forms of bacteria.

^* The use of common brand names as examples does not indicate a product endorsement.

¹ Instruments must be rinsed thoroughly with sterile water or saline to remove chemical sterilants before being used.

Table 3. Skin Antiseptics

An antiseptic is a chemical agent that either kills pathogenic microorganisms or inhibits their growth as long as the agent and microbe remain in contact. By custom as well as by federal law, the term antiseptic is reserved for agents applied to the body. The antiseptic may actually be a disinfectant used in dilute solutions to avoid damage to tissues.

^* The use of common brand names as examples does not indicate a product endorsement.

Table 4. Recommended Instrument Disinfectants

Always follow manufacturer's instructions.

^* The use of common brand names as examples does not indicate a product endorsement.

¹ Instruments must be rinsed thoroughly with sterile water or saline to remove chemical sterilants before being used.

Table 5. Suture Selection

^* The use of common brand names as examples does not indicate a product endorsement.

Suture gauge selection: Use the smallest gauge suture material that will perform adequately.

Cutting and reverse cutting needles: Provide edges that will cut through dense, difficult to penetrate tissue, such as skin.

Non-cutting, taper point or round needles: Have no edges to cut through tissue; used primarily for suturing easily torn tissues such as peritoneum or intestine.

PART III- GUIDELINES FOR RECOGNIZING POST-SURGICAL ANIMAL PAIN

Post-surgical animal pain poses special problems for researchers, faculty, employees and students. Animal surgery is to be done under complete anesthesia. The following comments relate to the post-operative period and may be helpful as we try to minimize or prevent as much pain as we can (before, during and after surgery).

The following statements are general statements held by most veterinarians and veterinary surgeons. Exceptions will exist to all these statements. Professional judgments are certainly called for if there are questions or disagreements.

1. Extensive abdominal surgery is less painful in animals than humans. Because animals don't need their abdominal muscles for postural support, movement puts less tension on their incision lines. Unlike humans, many animals become ambulatory soon after abdominal operations.
2. Lumbar and thoracic spine surgery in animals is also less painful than equivalent procedures in humans, because the human upright posture requires greater use of the lumbar and abdominal muscles and structures.
3. Chest surgery involving the sternum is more painful for animals than humans but if a lateral operative approach is used, the animal will probably feel less pain and be more likely to move with minimum distress after surgery.
4. Surgery on the eye, ear or surrounding structures seems to distress most animals. Look for typical signs of eye and ear pain which includes: head tilt, head shaking, pawing at the ears, or rubbing on an object. In addition to providing the animal with pain relief, make sure to protect the affected areas form the animal's rubbing and pawing which can create more damage.
5. Surgery on the femur and humerus is painful because of large muscle mass trauma and direct bone manipulation.
6. Trauma, disease or operative procedures involving the cervical skeletal structures are uncomfortable, but not necessarily painful. The animal's reluctance to move or a head-down stance signals pain.
7. As with operations around the eyes and ears, perirectal procedures seem to produce discomfort and distress.

IACUC POLICY ON USE OF ANALGESIA IN RODENTS:

To comply with PHS policies on minimizing pain and distress in laboratory animals, analgesics are required for all vertebrates, including rodents, that are subjected to a painful procedure. A painful procedure is considered to be any procedure that in the absence of evidence to the contrary, would cause pain in a human.

Pain in rodents may be manifested as decreased activity, grooming, or food and water consumption, guarding behavior (e.g. limping or a hunched posture) or increased aggression. However, the natural tendency of rodents is to hide signs of pain. Therefore, failure to observe post operative signs of pain in rodents is not a valid justification for withholding analgesia.

Any exceptions require that scientific justification is given to the IACUC, in writing, as to why the use of such analgesics would interfere with the goals of the study.

It is the responsibility of the investigator to provide and administer analgesics. The ARC can dispense drugs that may not be otherwise available to investigators and can provide drug administration services for a fee if requested. Contact Veterinary Services at 368-3490 to arrange for either service.

Topical/Local/Regional Anesthetics:

Topical anesthetic jelly, lidocaine (xylocaine) can be used in all species for painful wounds. Apply three times a day. Other uses of local anesthetics include local infusion, nerve blocks, pleural infusions for thoracotomy pain management and spinal anesthesia. Consult with the ARC (36803490) for specific techniques.

Bupivicaine (Marcaine®) is a long-acting local anesthetic. Infiltration of the surgical site will provide local anesthesia for 8 to 12 hours post-operatively.

Purchase 0.25% Marcaine
Dosage is 0.01 ml/25 g mouse, or 0.1ml/250 g rat
Dilution of 1:10 in sterile water, saline or PBS would give a final dose of 0.1 ml/mouse or 1.0 ml/rat

Narcotic Agonist/Antagonists:

Narcotic agonist/antagonists cause minimal cardiovascular or respiratory depression. They are excellent when given prior to recovery from anesthesia or for post-operative pain management. They can also be used with anesthetics to reduce the dose of anesthetic needed to perform surgery.

Don't forget that all controlled substances must be stored in a locked area. Complete usage records must be maintained.

Narcotic Agonists:

Narcotic agonists cause profound respiratory and CNS depression, decreased intestinal motility

Other:

Xylazine is an alpha-2 agonist with sedative and mild analgesic properties. It can be administered to frogs at 10 mg/kg IP or SQ BID provides very effective analgesia of up to 24 hours duration. Contact the ARC for availability of these drugs.

Carprofen is a nonsteroidal antiinflammatory drug with antiinflammatory and analgesic effects and minimal risk for toxicity in animals. The dose for carprofen in rodents is 5-10 mg/kg PO or parenterally. Currently, only oral preparations are available in this country (Rimadyl). A single dose after surgery, or dosing every 3 hours for 4 treatments have been shown to have comparable beneficial effects on post-surgical recovery.

Acetaminophen is a nonsteroidal antiinflammatory drug with minimal antiinflammatory and antiplatelet effects. While this drug provides good analgesia when administered as bolus doses either parenterally or enterally, administration in the drinking water has not provided measurable analgesia and unless otherwise contraindicated, buprenorphine administered as described above is recommended over acetaminophen administered in the drinking water.

Commonly used doses:

• 200 mg/kg IP or PO in rats and mice
• 15-30 ml of pediatric elixir (32 mg/ml) per 500 ml water bottle in drinking water.

Oral Administration of Buprenorphine or Carprofen in Jello

Animals must be fed Jello for several days before procedure to acclimate them to the food. To prepare, mix Jello according to the package directions to make "Jigglers". From the chart below, add an amount of Buprenex or carprofen to the corresponding volume of water, pour into ice cube trays for cooling with the corresponding volume per cube. Give 1/4 cube to each animal.

Post Operative Evaluation Form

A postop evaluation form has been provided for your convenience.

References: