Survival Surgery in Rodents & Species Not Regulated by the USDA
Last updated: March 5, 2012
The Public Health Service (PHS) and the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC) require that survival surgeries done on laboratory rodents and animals not regulated by the United States Department of Agriculture (USDA) be conducted using aseptic technique. Survival surgery is defined as any surgical procedure from which the animal will recover, even if only for short periods of time.
As with any activity involving animals, the individual(s) carrying out the procedure(s) must be adequately qualified and trained before beginning work involving survival surgery. In most cases, it will be necessary for the surgeon(s) to practice basic operative techniques on inanimate models and the specific procedure on cadavers before performing it on a living animal. Even then, the Institutional Animal Care and Use Committee (IACUC) may require additional animals to be used for non-survival training and/or observation and approval of technique by the Attending Veterinarian (AV) or appropriate representative before any animals may be recovered from anesthesia. Please contact Laboratory Animal Resources (LAR) or the AV to arrange for guidance and practice.
In the list below explanatory information is provided in italics for several requirements. Recommendations intended to assist in the implementation of this policy and enhance the well being of the surgical patient follow the requirements.
Institutional Recommendations for Non-USDA-Regulated Survival Surgery
Preparation of the Surgical Area
Unlike the requirements for a USDA surgical suite, the space does not need to be dedicated to surgery all the time. However, the surgery area and associated equipment must be organized, sanitized [chlorinated compound (MB-10 ®, Clidox ®) or quaternary ammonium chloride (MikroQuat ®)] and dedicated to that purpose while surgery is conducted. Contact time of the compound with the surface of the surgical area is the most critical component of effectively killing contaminating microorganisms. The surgery area must be dedicated to that purpose while surgery is conducted. It is recommended to clean the surface, then lightly mist the work surface and allow at least 10 minutes to air dry.
Covering the surface of the surgical work area with a circulating warm water blanket or clean paper (e.g., plastic-backed lab bench paper) or cloth will help prevent hypothermia and absorb fluids. Separate areas (e.g., different tables or portions of counters) should be used for patient preparation, surgery, and recovery.
Volatile anesthetic agents must be suitably scavenged using active house exhaust or passive charcoal canister. Use of injectable controlled substances must be recorded on institution-issued drug logs. Records of drug doses used, anesthetic monitoring data, and postoperative pain medication administered must be recorded and available for inspection upon request.
Preparation of the Surgical Instruments and Supplies
Surgical instruments, suture, and implants must be sterile before beginning the first surgery. Typically, surgical instruments are sterilized by steam autoclaving (minimum 121°C, 15 PSI, for 15 minutes), sutures are purchased in sterile packages, and heat-sensitive implants or instruments are sterilized chemically (e.g., Cidex), with gas (e.g., ethylene oxide, vaporized hydrogen peroxide), or irradiation. Depending upon the nature of the surgical procedure and the degree of instrument contamination during the surgery, a sterile set of instruments may be sued for up to 2-10 rodents, birds, amphibians or reptiles during the same surgery session. Between animals, the instruments must be cleaned of blood and gross debris and decontaminated using chemical disinfection or a point heat source (e.g., glass bead sterilizer). Care should be taken to ensure that instrument tips have cooled before using on animal tissue. Chemical sterilants should be removed from instruments by rinsing with sterile water or saline before contacting animal tissues to avoid chemical tissue damage. Ethanol alone is not sufficient to sterilize instruments. If a break is taken between groups of surgeries, instruments should be covered with a sterile drape to protect instruments from dust, hair, and air-borne environmental contaminants.
Preparation of the Animal
Food and water are not usually withheld from rodents due to their rapid metabolic rates. The surgery site must be clipped free of hair/fur. Preferred methods for removal of hair/fur are manual plucking or shaving with electric clippers. Loose hair can be removed with a vacuum, tape, or wet gauze. Skin should be aseptically prepared with an iodine (Betadine ®) or chlorhexidine- (Novlasan ®) based surgical soap followed by warm sterile water, warm sterile saline, or 70% alcohol (ethyl or isopropyl) rinses. Use of sterile cotton-tipped applicators is recommended for small rodents to minimize skin wetting that can cause hypothermia. Ethanol should be used sparingly to prevent serious, rapid drops in core body temperature. The aseptically prepared area should extend beyond the immediate area of the anticipated incision in order to minimize the risk of cross contamination and to allow for safe extension of the incision, if necessary. It is not recommended to use antiseptics on amphibian skin.
Whenever possible the surgery site should be draped with sterile drapes of cloth, paper, surgical gauze, or clear adhesive vinyl to minimize the risk of contaminating the surgery site. Care should be taken to avoid placing a drape such that the animal cannot be monitored. Drapes can have the added benefit of keeping the animal warm.
Preparation of the Surgeon(s)
Sterile surgical gloves, a surgical mask, and a clean outer garment (e.g., lab coat or scrub top) are required. Surgical gloves and the working surface must be cleaned between each animal.
It is important to minimize the 3 Ts: time, trash and trauma. Time is kept to a minimum by planning ahead, organizing equipment, and mastering the procedure to be performed. Trash is avoided by keeping instruments sterile, using proper aseptic technique, and wearing appropriate protective gear. Trauma is minimized by using gentle tissue handling techniques, using sharp instruments, keeping tissues moist with sterile saline, and placing sutures and staples to achieve tissue apposition while avoiding excessive tension. A subcuticular skin suture pattern will often preclude the chewing and premature removal of sutures by the animal.
Animals should be kept warm using an external heat source, particularly for procedures of any significant length (i.e., longer than 30 minutes). A circulating water blanket is the safest choice. Great care must be taken to prevent overheating or burning the animal when using other modalities. Some heating pads for rodents come with a rectal temperature probe that acts as a feedback thermostat to turn the pad on and off. In all other cases the external heat source should be separated from the animal by a towel or other protective barrier. Also for prolonged procedures, particularly those accompanied by blood loss, warmed fluid therapy should be administered. The recommended amount is equal to 1-2 cc per 100g body weight per hour of anesthesia plus any blood loss. Because of the small size of the patients covered by this policy, the intraperitoneal or subcutaneous routes are usually used. During the surgery, the animal's vital signs such as pulse rate, respiration rate, rectal temperature, tissue color, and response to noxious stimuli should be monitored and recorded serially so that corrective action can be taken promptly if necessary and an anesthetic record is generated.
The surgical work area and instruments should be thoroughly cleaned and sanitized after surgery. All blood and tissue must be removed and instruments re-sterilized. Any animal carcasses should be disposed properly.
Postoperative Recovery and Monitoring
Animals must be monitored until they have recovered satisfactorily from anesthesia (i.e., normal respiration, able to stand and walk). For mammals weighing less than 500 grams, it is advisable to use a heat lamp, warming pad, or external heat source beneath a portion of the housing cage for post-op recovery because they typically lose body heat during anesthesia. They should be able to choose the environment in which they are most comfortable - the warm area of the cage or the cooler, non-heated section. Animals must be monitored post-operatively to assure their well being as described in the IACUC-approved animal use protocol (AUP).
Rodents can often be stimulated to breathe in the case of apnea using gentle chest compression or inflating the lungs with a rubber bulb (from a pipette) applied to their nostrils. An oxygen rich environment also may be beneficial. The drug doxapram can be used to stimulate respiration and heart beat when administered orally at a dose of 0.5-1.0 mg/100g. Yohimbine (approx. dose 0.1-0.2 mg/100g by intraperitoneal injection) can be used to hasten the recovery in animals anesthetized with anesthetic combinations containing xylazine. Reversal of xylazine's anesthetic properties will also reverse its analgesic properties, so allowing it to wear off slowly may be advantageous to managing post-operative pain.
Animals cannot remain in investigator laboratories or other unapproved housing areas for longer than twelve hours without IACUC approval.
The date and name of the surgical procedure should be noted on the animal's cage card.
Depending upon the nature of the procedure and the condition of the animal, post-operative monitoring may range from once daily for one or two days to multiple times per day for extended periods.
Academy of Surgical Research. Guidelines for Training in Surgical Research with Animals. J Invest Surg 2009;22(3):218-225.
Brown MJ, Pearson PT, Tomson FN. Guidelines for animal surgery in research and teaching. Am J Vet Res 1993;54(9):1544-1559.
Fossum TW. Small Animal Surgery. St. Louis, MO: Mosby, Inc.; 1997.