Purpose

This document explains the current requirements of federal laws and federal funding agencies with respect to survival surgery in rodents.

Background

The federal Animal Welfare Act and the ILAR Guide for the Care and Use of Laboratory Animals (Guide) both set standards for biomedical research involving live, vertebrate animals. These documents have been adopted by virtually all-public and private funding sources, and are pertinent to all research projects, regardless of funding source.

Survival surgery in non-rodent mammals must be conducted in dedicated facilities. Investigators cannot perform survival surgery in non-rodent mammals in their own laboratories, but must make use of survival surgery suites built according to the federal standards.

Rodent Survival Surgery: Investigators may conduct rodent survival surgery in their laboratories or in an animal facility procedure room. While there is no requirement for a dedicated surgical facility for rodents, there are requirements about how rodent surgery must be conducted.

While dedicated facilities are not required for rodent survival surgery, aseptic technique is required by Federal Guidelines. General requirements stated on pages 60-65 of the ILAR Guide (NRC Publication 86-23, revised 1996) include the use of sterile instruments, sterile surgical gloves, clean surgical attire, and aseptic technique. The balance of this document will expand and clarify the statements in the Guide.

Procedure

Surgery Facilities: rodent surgical area must be easily sanitized. The immediate surgical area should not be used for other purposes during the time of surgery. Surgery may be conducted on a clean, uncluttered lab bench or table, in a laminar flow HEPA-filtered hood, or in a glove box or other type of isolator. It is strongly recommended that surgeries be performed in a HEPA-filtered laminar flow hood to minimize the amount of contamination during surgery and protect the animals from unwanted infections, such as mouse hepatitis virus, rat coronavirus, mouse parvovirus, etc.

Preparation of Surgery Table Surface: Prior to and between surgeries, clean and disinfect the surface upon which surgery will be performed. Use soap and water, rinse thoroughly, and follow with an appropriate disinfectant. Commonly used disinfectants are quaternary ammonium compounds (such as Roccal); household bleach diluted 1 part to 32 parts water, chlorine dioxide-based sterilant (Clidox), chlorhexidine (Nolvasan), 70% alcohol, or other antimicrobial agent. Disinfectants must be prepared and used according to the manufacturer's recommendations.

Preparation of Surgical Instruments: Surgical instruments must be sterilized for use in survival rodent surgery. Several techniques (steam, dry heat, ethylene oxide, or chemical agents) can be used to sterilize instruments and other materials that will come in contact with the animal's tissues. Steam or dry heat are the preferred methods to sterilize surgical instruments.

Chemicals used to sterilize surgical instruments must be classified as a sterilant not a disinfectant. Chemical sterilants typically require a contact time of 6-24 hours, depending on the chemical used. For example, chlorine dioxide requires a minimum of 6 hours of contact time. Glutaraldehyde and Cetylcide require instruments be soaked a minimum of 10 hours. Chemical sterilants must be prepared and used according to the manufacturer's recommendations. All instruments sterilized by chemicals must be rinsed in sterile water before use in tissues.

Multiple Surgeries: When performing surgeries on multiple animals, at least 2 sets of sterile instruments should be available to allow re-sterilization of instruments between animals. Chemical sterilants typically require hours of contact time, therefore they are seldom practical for re-sterilizing instruments on the same day as surgery.

Using a glass bead sterilizer is the optimal method for re-sterilization of instrument tips on the day of surgery. While the first set of instruments is being re-sterilized, the second set is used. After using a set of instruments, remove all organic material and then immerse the instruments in a glass bead sterilizer. Make sure the tips of the instruments have cooled before using them on tissue. Tips may be cooled by dipping in sterile water. It should be noted that glass bead sterilizers and tips of instruments sterilized in glass bead sterilizers are capable of producing severe burns. Care must be exercised when using a glass bead sterilizer, and all manufacturer instructions and safety precautions must be followed to avoid injury.

A new sterile instrument pack should be used after every 4 or 5 major surgical procedures.

Preparation of the Animals: While under anesthesia and prior to taking the animals to the surgery area, remove all hair for at least a centimeter on either side of the surgical site. Hair can be removed by clipping with a #40 clipper blade (for example Pocket Pro-Trimmer), shaving with a razor, plucking (in anesthetized rodents), or by using a depilatory cream. Then remove loose hair with a dry gauze or careful vacuuming.

Place lubricating ophthalmic ointment (such as Lacrilube or Tearfair) in the anesthetized animal’s eyes to prevent drying of the cornea.

Clean and aseptically prepare the surgical site. Use an effective antiseptic surgical scrub solution (Nolvasan, Betadine, etc.). Carefully scrub the area with a new clean surgical sponge or sterile cotton swab. Scrub in a gradually enlarging circular pattern from the center of the proposed incision to the periphery. The sponge or swab should not be brought back from the contaminated periphery to the clean central area. Repeat with a 70% alcohol (or sterile water) soaked sponge or sterile cotton swab. Repeat this process three times to minimize the presence of micro-organisms.

To prevent hypothermia, do not wet the animal any more than necessary. Care should be taken to prevent contamination of the sterile surgical field during subsequent handling and positioning of the animal.

Place the animal on a clean absorbent surface and maintain body temperature using a circulating water blanket, warm water bottle, bubble wrap, or equivalent external heat source, taking care to not cause thermal burns to the animal's skin.

Preparation of the Surgeon: Surgeons must wash their hands with a surgical scrub (e.g., Betadine Scrub, Nolvasan Scrub).

Wear a mask, sterile gloves, and clean scrub shirt or lab coat. A new pair of sterile surgical gloves should be used for each animal.

During Surgery: The surgical field must be kept sterile throughout the procedure. Sterile instruments must be prevented from contacting non-sterile surfaces. Instruments must be placed on a sterile surface when not in use. Sterile surgeon's gloves are required by the Guide. In most cases, the use of sterile drapes is also required for maintenance of the sterile field.

Monitor the animal carefully during the surgical procedure. Anesthetized animals should not be left unattended. Surgeons should pay close attention to the animal's heart rate, respiratory rate, and body temperature. Evaluating the animal’s response to surgery will also help determine the anesthetic depth.

Postoperative Care: Recovering animals should not be placed onto loose bedding material until they are fully awake, as suffocation can result. A paper towel may be placed between the bedding and the animal until it awakens from anesthesia. Prevent hypothermia by placing the recovering animals in a warm cage. If necessary, the cage may be placed on a bedded or padded surface and supplied with supplemental heat as required (such as a circulating hot water pad). Be cautious with supplement heat sources; hyperthermia can be as detrimental as hypothermia.

Dehydration can be ameliorated by the administration of appropriate fluid therapy. Initially this may be done by giving 1 to 2 ml of warm fluids (0.9% NaCl or equivalent) per 100 grams 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. Consult the veterinary staff for assistance with fluid therapy.

Animals should not be returned to the vivarium until they are sternal and clearly awake. To prevent cannibalism or suffocation, it is best to separate non-ambulatory from ambulatory rodents.

A member of the investigator’s staff or other individual to whom postsurgical care has been delegated must see post-surgical animals at a minimum every day for 7-14 days. Animals must be given analgesics as specified in approved Animal Care and Use Protocols and if needed thereafter, as prescribed by a clinical veterinarian.

In general, intra- or postoperative antibiotics are unnecessary when aseptic technique is maintained. If an inadvertent contamination occurs during surgery, consult with a clinical veterinarian immediately. If routine postoperative antibiotics are thought to be needed, their use needs to be included in the approved Animal Care and Use Protocol.

All daily observations and treatments must be recorded on the animal's postsurgical record. External wound clips, staples, or sutures must be removed when surgical incisions are healed 7-14 days after the surgery, or as outlined in the approved Animal Care and Use Protocol. Consult with a clinical veterinarian if you have questions regarding the optimal staple or suture removal time.

The veterinary staff must be notified if postsurgical complications occur.

Records: Postsurgical records must be kept in the room where the animals are housed per records policy:

http://safetyservices.ucdavis.edu/iacuc/attending-veterinarian/anesthetic-records

Use the following link for an example of a completed record:

http://safetyservices.ucdavis.edu/iacuc/attending-veterinarian/AnesthRecordSimpleSample.pdf

Having the record in the room accomplishes the following.

1. It explains the condition of the animals to animal care staff (e.g., a sedated animal may otherwise be thought to be ill).
2. It assures animal care staff, veterinary staff and USDA Animal Welfare Inspectors that proper care is being given to the animals.
3. It informs animal care staff and veterinary 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 postsurgical record to be used for a group of rodents (see above referenced example). Such a record would have a list of the animal numbers down the side and columns indicating dates. The column entries would include a notation that the animal has been checked, any abnormal observations, and a list of any therapeutics given including drugs, doses, and routes of administration. Records should be kept current during the immediate postoperative period (7-14 days).

To assist animal care and veterinary staff with identification and observation of postoperative rodents, it is recommended that special identification cage cards be placed on the cage when rodents are returned to their cages and removed at the end of the postoperative period or when sutures or staples are removed. Postsurgery identification cage cards are available through animal facility supervisors or Campus Veterinary Services.

After all wounds have healed and all sutures/wound clips have been removed, the postsurgical record requires no further entries, but should continue to be kept in the area where the animals are housed. When the study is completed and the animals are euthanized, the record may either be kept by the investigator or discarded.

Non-Survival Rodent Surgeries: While it is not necessary to follow aseptic technique when performing non-survival surgeries in rodents, at a minimum the surgical site should be clipped, the surgeon should wear gloves, and the instruments should be clean.

Contact Campus Veterinary Services (530-752-0514) for questions regarding animal health, anesthetic support, surgical wound care, postoperative analgesia, or other questions regarding these guidelines.

Procedure: IACUC-22
Date: 8/7/2008
Enabled By: AWA, PHS
Supersedes: 11/29/2007, 06/22/2004, 02/12/2004