DIRECTIONS: Review the following material. Keep copies of guidelines with applicable protocols. You may find it helpful to post a copy of these guidelines in your laboratory. Questions should be forwarded to the APLAC office, 723-4550.
TRAINING: Training in these techniques and the humane treatment of laboratory animals during the procedures is taught by the Veterinary Service Center (VSC) staff. All new personnel who will be performing these techniques should contact VSC staff for training (725-9901).
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GUIDELINES FOR RODENT SURVIVAL SURGERY
I. Preparation of Surgical Facilities and Instruments
Surgical Facilities
The Guide for the Care and Use of Laboratory Animals states, "For most rodent surgery, a facility may be small and simple, such as a dedicated space in a laboratory appropriately managed to minimize contamination from other activities in the room during surgery." [1] The location should be uncluttered, easily disinfected, and separate from where the animal will be prepared for surgery. Avoid locations under air ducts and high traffic areas. [2] Clean and disinfect the lab bench or table top using soap and water, followed by a hard surface disinfectant* (e.g., Cide Swipes®, Nolvasan®). [3]
Surgical Instruments
Surgical instruments must be sterilized. Several techniques, such as autoclaving, hot bead sterilizers, dry-heat sterilization, or cold sterilants (e.g., Spor-Klenz-RTU®, Clidox-S®)* can be used. Organic material must be removed prior to sterilization. Appropriate contact time (often several hours; see http://oacu.od.nih.gov/ARAC/surguide.pdf; Table 3) must be allowed. You must rinse instruments with sterile water following sterilization with cold sterilants.*
When performing surgeries on multiple animals during a single session, instruments may be disinfected3 or sterilized between animals. First, remove all organic material (e.g., blood, tissues) from the instruments. Instruments can be sterilized using a hot bead sterilizer (tips only), dry-heat sterilizer, autoclave, or cold sterilant, following recommended contact times and remembering to rinse instruments with sterile water after using a cold sterilant, or disinfected with a disinfectant *(see http://oacu.od.nih.gov/ARAC/surguide.pdf; Table 4)
*It is important that disinfectants are prepared according to the manufacturer's instructions. All organic materials must be cleaned from the surface before the disinfectant is used. Some disinfectants contain paraformaldehyde and must be rinsed off with sterile water or saline.
II. Preparation of the Animal for Surgery
Preparation of the animal for surgery is best done at a location distant from where the surgery will be performed.
Protect the eyes. After induction of general anesthesia, place a lubricating ophthalmic ointment (e.g., Lacri-Lube®) in the anesthetized animal's eyes to prevent drying of the cornea. Note: some anesthetics result in the eyes remaining open.
Remove hair from the surgical site. Use an electric clipper, razor, depilatory cream, or, in mice, by plucking the hair. Do not abrade the skin. Depilatory creams can result in burns, so they should be used with caution and only for short periods of time (< 1 min). Removing the hair only with scissors is not sufficient. Collect loose hair and dander with a vacuum or an adhesive-tape lint remover, e.g., ScotchTM Lint Roller or similar tool.
Disinfect the surgical site. Using cotton-tipped swabs or similar, apply a thin layer of Techni-Care® (Care-Tech Laboratories®, Inc.) in a gradually enlarging circular pattern from the future incision site outwards towards the periphery. Repeat once if surgical site has a heavy organic load. After a 2-minute contact time, air dry or blot dry with a sterile gauze. Do not remove with alcohol; alcohol inhibits Techni-Care’s® activity. Alternatively, disinfect the skin by alternating between a surgical scrub (e.g., Nolvasan® or Betadine® Surgical Scrub) and 70% ethanol for 3 cycles. Using swabs prevents excessive wetting of the animal, which can result in hypothermia. Although 70% ethanol is preferred, sterile water may result in less hypothermia.
Transport the animal to the surgical area. Avoid contaminating the surgical site.
Place the animal on a clean absorbent surface, preferably on a heat source (e.g., circulating water blanket, slide warmer, instant heat device) to prevent hypothermia.
Drape the animal with a sterile drape. In addition to keeping the surgical site clean, draping provides a sterile surface on which to place one’s instruments.
III. Preparation of the Surgeon
Disinfect the hands with an appropriate antiseptic soap or Techni-Care®.
Wear a mask, sterile gloves, and a clean scrub shirt or lab coat. A mask is not worn if mouth pipetting of embryos is required. If multiple animals are operated at one sitting, one pair of sterile gloves may be used, provided that the gloves are rinsed with 70% ethanol AND you did not contaminate your gloves, e.g., by handling another rodent. [2]
IV. Intraoperative Monitoring and Rodent Surgery Log Table
Monitoring in rodents is a constant process once general anesthesia is induced. Animals must never be left alone during surgery. All required materials (e.g., viral vectors, drugs, incubators, cells, etc.) should be easily accessible to the surgeon.
Surgery logs must be maintained. Group Health Records are appropriate if the same procedure is being performed on a group of animals. If the surgery is relatively short and the surgeon is working alone, completing a Rodent Surgery Log Table is sufficient. An inexpensive digital voice recorder that is capable of recording for >8 hours, e.g., Olympus VN-4100, can be used to document a day’s surgery and later transcribed into a notebook.
A Rodent Surgery Log Table should include, at minimum, Protocol Number, Surgeon, Date of Surgery, animal or cage ID #, Weight, all Drugs (anesthetics, analgesics, antimicrobials, reversal agents, etc.) including dose, route, and time of administration, Fluids, any Complications (surgical or post-surgical, including deaths), and visual assessment of the following parameters:
- Anesthesia depth: Assess withdrawal reflex prior to making any incision and periodically during the surgery
- Skin color of the ears, paws, muzzle, or tail: Monitors oxygenation; should be pink
- Breathing pattern: Breathing should be regular and within normal range
NOTE: When controlled substances are used (e.g., ketamine, buprenorphine), the animal ID number and dose written in the Rodent Surgery Log must match that which is written in the Controlled Substance Log. APLAC may require more extensive monitoring for procedures using neuromuscular blocking agents or with a potential for perioperative pain.
V. Post-operative Care
If using inhalant anesthetics, e.g., isoflurane, analgesics (opioids, NSAIDs, and/or local anesthetics) are best given before the start of surgery. This pre-emptive analgesia results in overall better anesthesia and blocks “pain wind-up.” If using injectable anesthetics, e.g., ketamine/xylazine, analgesics may be given at closure of the incision site and prior to recovery. Consult with a VSC veterinarian if you have questions about potential adverse effects of your choice of analgesic medication on either the animal or your experiments.
To minimize the effects of dehydration, 1 - 2 ml per 100 gm of body weight of prewarmed fluids (0.9% NaCl or equivalent) may be administered subcutaneously simultaneously with analgesics. Fluids are highly recommended if the surgery lasts longer than 30-45 minutes and will result in quicker recovery. If blood loss occurs during the surgery, or if the animal is slow to recover from anesthesia, provide additional prewarmed fluids.
Do not leave an animal unattended until it fully recovers consciousness. Recovering animals should be monitored every 15 minutes until they are mobile.
To prevent aspiration of bedding and asphyxiation, place anesthetized animals on clean paper towels during recovery; never place them directly on bedding.
To prevent cannibalism, do not place anesthetized rodents in a cage with normal, non-anesthetized animals.
To minimize hypothermia, place the cage in a warm room or partially on a circulating warm water blanket, heating pad, or slide warmer, or under a low wattage light bulb. Periodically turn the animal to prevent pulmonary hypostatic congestion. Monitor the temperature at the level of the animal with a thermometer. Temperature should not exceed 85-95°F (29.4-32.2°C). [3]
After the immediate postoperative period, check the animal daily until the removal of the sutures or wound clips to ensure that there are no complications. Complications must be noted in the Rodent Surgery Log (see above). Sutures or wound clips must be removed in 7-14 days if the incision site is healing normally. If the animal appears ill or if the incision site is moist and appears abnormal, contact the VSC at 723-3876 and ask to speak to the Veterinarian On Call.
VI. "Tips-Only" Technique [2]
The “Tips-Only” technique is appropriate for surgeries involving small incisions, e.g., embryo transfer and ovariectomy. The “Tips-Only” Technique does not require that the surgeon wear sterile gloves. This allows the surgeon to manipulate the animal, lights, anesthesia machine, or focus a microscope throughout a surgery without concern about breaking sterility. However, the “Tips-Only” technique requires that the surgeon never contaminate the instrument tips, suture, suture needle, or any part of the surgical field either directly by touching or by inadvertently placing the surgical instruments, suture, or suture needle on a surface contaminated by the gloves. Surgical instrument tips are sterilized with a hot-bead sterilizer.
Contact the VSC if you have any questions regarding proper “Tips Only” techniques.
VII. Additional Sources of Information for Analgesia/Anesthetic Agents:
http://med.stanford.edu/compmed/animal_care/mice.html
http://med.stanford.edu/compmed/animal_care/rats.html
VIII. Resources on Disinfectants, Sterilants, and Wound Closure Materials:
Med. Records for Animals Used in Research, Teaching, & Testing. ILAR J. 2007. 48:37-41.
NIH Guidelines for Survival Rodent Surgery, revised 10/07 (http://oacu.od.nih.gov/ARAC/surguide.pdf)
National Research Council, 1996, Guide for the Care and Use of Laboratory Animals, p. 78.
Tech. in Aseptic Rodent Surgery. 2008. Curr Protoc Immunol. Chap. 1:Unit 1.12.1-1.12-14.
[1] National Research Council, 1996, Guide for the Care and Use of Laboratory Animals, p. 78.
[2] Tech. in Aseptic Rodent Surgery. 2008. Curr Protoc Immunol. Chap. 1:Unit 1.12.1-1.12-14.
[3] NIH Guidelines for Survival Rodent Surgery, revised 10/07 (http://oacu.od.nih.gov/ARAC/surguide.pdf)
[4] Med. Records for Animals Used in Research, Teaching, & Testing. ILAR J. 2007. 48:37-41.
Contact: A-PLAC Administrator
Last updated: July 2009
