Survival Surgery in Rodents
Purpose and Background
This document provides guidelines for survival surgery in rodents conducted at The Ohio State University. The federal Animal Welfare Act and the NIH Guide for the Care and Use of Laboratory Animals both set standards that are obligatory for biomedical research involving live, vertebrate animals. These guidelines are consistent with the Animal Welfare Act and the NIH Guide for the Care and Use of Laboratory Animals.
Surgery Facilities
A rodent surgical area can be any room or portion of a room that is 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. Prior to surgeries, clean and disinfect the surface upon which surgery will be performed. Commonly used disinfectants include quaternary ammonium compounds (such as Roccal), household bleach diluted 1 part to 32 parts water, chlorine dioxide-based sterilant (Clidox), and Spor-Klenz. Disinfectants must be prepared and used according to manufacturer's recommendations.
Preparation of Surgical Instruments
Surgical instruments must be sterilized for use in survival rodent surgery. Steam or dry heat are the preferred methods to sterilize surgical instruments. Alternatively, instruments may be soaked in a cold sterile solution (such as Cetylcide) according to the manufacturers recommendations. If chemical sterilants are used on surgical instruments, the instruments must be rinsed in sterile water before use. When performing surgeries on multiple animals, the instruments may be used again if all blood/tissue is wiped off the instruments with a gauze soaked in 70% alcohol, and they are then immersed in 70% alcohol until ready to re-use.
Where only the tips of instruments have contacted the animals, instrument tips may be wiped with 70% alcohol and maintained in a sterile environment between animals. In this case, use of a sterile cloth instrument holder with pockets may also reduce the potential for contamination. The instruments are placed in the pockets and covered while the next rodent is prepared for surgery. Even with the use of an alcohol wipe between rodents and use of the sterile cloth instrument holder, a new sterile instrument pack should be used after every 4 or 5 major surgical procedures.
Preparation of the Animals
Remove food (not water) from the animal the evening prior to surgery. Anesthetize the animal as outlined in the approved IACUC protocol. Ensure that analgesics are administered perioperatively if not contraindicated by the study requirements.
Prior to taking the animal to the surgery area, clip the hair from the surgical site. Hair can be removed by clipping with a #40 clipper blade, shaving with a razor, or by using a depilatory cream. Be sure to remove hair from a large enough area to incorporate the entire surgical site. Vacuum or otherwise remove loose hair before transferring the animal to the surgery area. Place lubricating ophthalmic ointment (such as Lacrilube or Tearfair) in the anesthetized animal's eyes to prevent drying.
Clean and aseptically prepare the surgical site using an effective antiseptic surgical scrub solution (Nolvasan surgical scrub, Betadine Scrub, etc.). Carefully scrub the skin with a clean cotton swab or surgical scrub pad soaked in the surgical scrub. Scrub in a gradually enlarging circular pattern from the center of the proposed incision to the periphery. Do not bring the swab/pad back from the periphery to the clean central area. Next, using a cotton swab or surgical pad soaked in 70% alcohol repeat the cleaning pattern. Repeat the surgical scrub/alcohol cycles for a total of 3 times, each time beginning at the center and proceeding to the periphery. Do not wet the animal any more than necessary in order to prevent hypothermia. 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.
Preparation of the Surgeon
Wear a mask, hair bonnet (or secure hair), and clean scrub shirt or lab coat. Surgeons should wash their hands with an appropriate surgical scrub (e.g. Betadine Scrub, Nolvasan Scrub).
A new pair of sterile gloves should be used for each animal. Alternatively, surgeons may wipe their gloves for 30 seconds with sterile gauze pads soaked in 70-90% alcohol between animals. Latex exam gloves may be used if from a freshly opened box or if previously sterilized IF the items coming in contact with the surgical wound remain sterile. In order not to contaminate the whole box or batch, gloves should be removed with sterile forceps.
During Surgery
The surgical field must be kept sterile throughout the procedure. Sterile instruments must not contact nonsterile 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. Clear plastic sterile drapes are available or can be made by gas sterilizing plastic-wrap cut in appropriate sizes and edged with masking tape to keep it flat during use. The drape should be nonpermeable to maintain a true sterile field.
Monitor the animal carefully during the surgical procedure. Surgeons should pay close attention to the animal's level of anesthesia. Parameters such as respiratory rate, muscle tone, heart rate, and toe pinch reflexes can help assess the anesthetic level. Additional anesthetic must administered if the animal is responding to surgical stimulation.
Postoperative Care
During the immediate postoperative period, the rodent should be observed every 30 minutes until it is fully recovered from anesthesia. Prevent hypothermia by placing the animals in a warm room or cage. If necessary, the cage may be placed on a bedded or padded surface and supplied with supplemental heat as required. Do not place the rodent directly on bedding material until fully awake in order to prevent aspiration of bedding. A paper towel can be used on top of the bedding during the recovery period for this purpose. Be cautious with supplement heat sources especially heat lamps; 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 sterile fluids (0.95% NaCl or Lactated Ringer's) 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. Post-operative pain or distress must be monitored and treated (See appendix) as approved in the IACUC protocol. The IACUC will require scientific justification prior to approval for withholding analgesics if pain/distress is detected. An attending veterinarian should be consulted immediately if an animal is experiencing pain or distress which does not respond to the approved analgesic protocol.
As a general rule, animals should not be returned to the vivarium until they are stable and able to assume a normal posture. To prevent cannibalism or suffocation, house rodents individually until they are ambulatory. External wound clips and sutures should be removed 7-14 days after the surgery.
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. ANIMALS SHOULD BE OBSERVED DAILY UNTIL ALL WOUNDS HAVE BEEN HEALED AND SUTURES OR WOUND CLIPS ARE REMOVED.
Records
A post-operative record should be kept in the room where the animals are housed. 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 federal 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, a composite post-operative record may be used for a group of rodents. Download an example of a Post-Operative Care Record here, this example may be copied or modified as needed. Important information to include in the post-operative record is the animal's identification, surgical procedure summary, any therapeutics given including drugs, doses, and routes of administration, and the observation date and findings. After all wounds have healed and all sutures/wound clips have been removed, the post-operative record requires no further entries. When the study is completed and the animals are euthanized, the record may either be kept by the investigator or discarded.
Further information
The University Laboratory Animal Resources - Surgical Technical Services (292-5494) may be consulted for questions regarding the surgery guidelines. The ULAR veterinary staff may be consulted for questions regarding animal health, analgesia, surgical wound care, animal anesthesia techniques, surgical procedures, and provision of post-operative care. Training on surgical techniques, anesthesia, etc. is available at no charge to investigators and staff upon request. Contact your attending veterinarian or ULAR technician for additional information.
Appendix
Recognition of pain and distress in rodents:
Appropriate analgesia can only be achieved if the signs of pain are recognized in the animal. Since pain assessment in rodents is difficult, it is critical that the investigator be familiar with the normal behavior of the strain/stock they will be using. Some indicators that the mouse/rat is in pain include hyper or hypo-excitability, vocalization, altered ambulation, decreased grooming or rearing, decreased socialization if group housed. In addition to these behavioral changes, one might observe piloerection, dirty coat, or ocular buildup of porphyrin in rats. Decreased eating and drinking may also indicate pain or distress but must be interpreted carefully, especially if the analgesic being used has a sedative effect. Analgesic administration should be utilized if any pain or distress is suspected. Alleviation of clinical signs after analgesic use, helps confirm that the signs were indeed pain/distress related. There are many references available that discuss recognition of pain and distress in rodents, for example: Pain Management in Animals edited by Paul Flecknell and Avril Waterman-Pearson, W.B. Saunders, 2000. Project staff should consult with a ULAR veterinarian for training in recognition and treatment of clinical signs of pain and distress as they apply to the surgery to be done.
Examples of approved analgesic regimes used in rodents:
From: Formulary for Laboratory Animals , 2nd edition by C. Terrance Hawk and S. Leary, Iowa State University Press, 1999.
- Rat
Buprenorphine 0.01-0.05 mg/kg BW IV, SC every 8-12 hours
Butorphanol 0.05-2.0 mg/kg BW SC every 4 hours
Carprofen 5 mg/kg BW SC
Ibuprofen 10-30 mg.kg BW PO - Mouse
Buprenorphine 0.05-0.1mg/kg BW SC twice a day
Butorphanol 1-5 mg/kg BW SC every 4 hours
Ibuprofen 7.5 mg/kg BW PO - Other Rodents
Please consult the Formulary for Laboratory Animals (mentioned above) for additional information on anesthetics and analgesics in other rodent species.
