Revised on
10/03/16 06:33pm

Survival Surgery Guidelines for Rodents

Purpose:

This document explains the current requirements for performing survival surgery in rodents.

Policy:

General requirements for rodent survival surgery include:

  • Designated surgical area
  • Use of sterile instruments
  • Aseptic technique
  • Anesthesia and analgesics as outlined in approved protocols
  • Monitoring and care of animal’s well-being
  • Post-operative care
  • Records
  • Training of personnel

Investigators may be approved to conduct rodent survival surgery in their laboratories in special circumstances however use of an animal facility procedure room is preferred. While there is no requirement for a dedicated surgical facility for rodents, there are requirements for how rodent surgery must be conducted.

Major survival surgery in non-rodent mammals must be conducted in dedicated facilities. Investigators cannot perform major survival surgery in non-rodent mammals in their own laboratories, but must make use of IACUC approved survival surgery suites which meet federal standards.

 

Procedure:

A. Acclimation Period:  Per the Guide for the Care and Use of Laboratory Animals, newly received animals should be given a period for physiologic, psychological, and nutritional stabilization before their use.

Rodents should have a 3 day acclimation period prior to anesthesia, surgery, or similarly invasive procedures.

It is the responsibility of the principle investigator to ensure the animal has been released for study prior to placing the animal on project. Additional information on acquisition and acclimation can be found here.

B. Designated Surgery Area:  The Guide recommends that rodent survival surgery be performed in a procedure room. The surgical area must be easily sanitized. The immediate surgical area should not be used for other purposes during the time of surgery and traffic in this area should be minimized.

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), or other antimicrobial agent. Disinfectants must be prepared and used according to the manufacturer's recommendations.

 

C. Use of Sterile 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: 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 it is recommended to have at least 2 sets of sterile instruments 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. It is suggested that a new sterile instrument pack be used after every 6 major surgical procedures.

Glass bead sterilizer: The optimal method for re-sterilization of instrument tips on the day of surgery is using a Glass bead sterilizer. 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 for 20-30 seconds (follow manufacturers guidelines). 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.

D. Aseptic Technique:

Preparation of the Animals: While under anesthesia (as approved in a protocol) and prior to taking the animals to the surgery area, remove all hair from at least a centimeter on either side of the surgical site. Hair can be removed by clipping with an appropriate sized clipper, 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 Purilube®) 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 (e.g., Nolvasan, Betadine). 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.

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 must 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. In most cases, the use of sterile drapes is also required for maintenance of the sterile field.

E. Monitoring and care of animal’s well-being: Monitor the animal carefully during the surgical procedure. Anesthetized animals must not be left unattended during the procedure. Surgeons must pay close attention to the animal's heart rate, respiratory rate, and body temperature.  Assess the animal’s depth of anesthesia by toe/tail pinch prior to making an incision.  Evaluating the animal’s response to surgery (increased respiratory rate, movement, vocalization) will also help determine the anesthetic depth. 

Maintain Body Temperature: 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, or equivalent external heat source, taking care to not cause thermal burns to the animal's skin (heat lamp).

F. 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 (heat lamps); hyperthermia can be as detrimental as hypothermia.

Observe Animal: Animals must be in an area where they can be frequently observed until they are sternal and clearly awake. To prevent cannibalism or suffocation, it is best to separate non-ambulatory from ambulatory rodents. Once animals return to standard housing, the cage should be identified with a blue index card reading “Post-Op”. Post-Op cards can be obtained from the facility manager and husbandry staff.

Maintain Hydration: 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. 

Daily Post-op Checks: A member of the investigator’s staff or other individual to whom postsurgical care has been delegated must check postsurgical animals at least daily for a minimum of 7 days, until wounds have healed, and/or sutures/staples have been removed, unless the protocol has been approved for shorter duration for minor procedures. Animals must be given analgesics as specified in approved Animal Care and Use Protocols and if needed thereafter, as prescribed by a clinical veterinarian.  

 

Antibiotics: 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.

G. Records: Postsurgical records must be kept in the room where the animals are housed the duration of post-operative care. An example of an anesthesia/surgery record and the policy for record documentation can be found on the TRACS Veterinary Services website under forms: Standards of Care Program Policies

Documentation: 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. 

H. Training: Principle Investigators (PIs) are required to ensure all staff conducting or assisting with rodent survival surgeries are appropriately trained and that training has been documented.  “Researchers conducting surgical procedures must have appropriate training to ensure that good surgical technique is practiced – that is, asepsis, gentle tissue handling, minimal dissection of tissue, appropriate use of instruments, effective hemostasis, and correct use of suture materials and patterns. Personnel conducting survival surgery must take the online UC Davis Survival Surgery course. Personnel must obtain training on aseptic techniques from other knowledgeable personnel or by attending the Aseptic Techniques instructor led training.  PIs and staff may also receive hands-on training for specific surgical techniques from others proficient in the procedure.

Training Opportunities:

  • An Aseptic Technique course and UC Davis Survival Surgery is offered by the IACUC Office and can be found here.
  • There is also an AALAS Learning Library online module for Aseptic Technique for Rodent Survival Surgery (http://www.aalaslearninglibrary.org/).

Please contact the IACUC office for questions regarding training or to obtain an AALAS Learning Library username/password.

I. Non-survival Rodent Surgeries: "In non-survival surgery, an animal is euthanized before recovery from anesthesia. It may not be necessary to follow all the techniques outlined in this section if non-survival surgery is performed, but at a minimum, the surgical site should be clipped, the surgeon should wear a lab coat and gloves, and the instruments and surrounding area should be clean. For non-survival procedures of extended duration, attention to aseptic technique may be more important in order to ensure stability of the model and a successful outcome. 

You may contact TRACS Veterinary Services (530-752-0514 or lahc@ucdavis.edu) for questions regarding animal health, anesthetic support, surgical wound care, postoperative analgesia, or other questions regarding these guidelines.

References:

  1. Guide for the Care and Use of Laboratory Animals, Eight Edition - pp. 111, 115-120, 144
  2. Animal Welfare Regulations, 2008 - pg. 26

Procedure: IACUC-22
Date: May 07, 2015
Enabled By: AWA, PHS
Supersedes: February 21, 2013