Preparation of Surgical Instruments and Supplies
Preoperative Considerations/Care
Multiple Surgeries in a Single Session
Post-operative infections in rodents can and do occur. The misconception that rodents have an innate resistance to bacterial infection has not been scientifically substantiated. Such infections, which may not be apparent on casual observation, will cause loss of vessel cannulations1, and numerous changes in physiologic parameters2 . In accordance with good scientific practice and standards set forth in the Public Health Service Guide for the Care and Use of Laboratory Animals (Guide) and the Federal Animal Welfare Act, aseptic surgical procedures must be used.
The NIH Guide provides the following guidelines for rodent surgery:
1) Major survival surgery on rodents does not require a special facility but should be performed using sterile instruments, surgical gloves, and aseptic procedures to prevent clinical infections.
2) Major survival surgery is defined as any surgical intervention that penetrates a body cavity or has the potential for producing a permanent handicap in an animal that is expected to recover.
3) Training in aseptic procedures should be provided for those who require it.
The Animal Welfare Act states:
“Non-major operative procedures and all surgery on rodents do not require a dedicated facility but must be performed using aseptic procedures. Operative procedures conducted at field sites need not be performed in dedicated facilities but must be performed using aseptic procedures.”
A separate room used primarily for aseptic procedures is desirable; however, it is appropriate to perform survival rodent surgical procedures in a conventional laboratory setting using aseptic technique. The investigator should adopt the following standards for aseptic procedures:
•A clean, uncluttered work area and a sanitized work surface should be utilized for the surgery area.
•The work area should be located to minimize laboratory traffic not related to the surgical procedure and dedicated exclusively for surgery, when in use.
•Considerations should also be given to locate the surgery away from potential sources of contamination such as open windows, fans, or fume hoods which can blow dust into the area and increase desiccation of exposed tissues.
•The surgery surface should be disinfected (70% alcohol or a quaternary ammonium compound) before use. It often simplifies the maintenance of asepsis if a sterile drape is then applied over the surgery surface.
Preparation of Surgical Instruments and Supplies
•Survival surgical procedures on all mammalian species must be conducted using aseptic technique which requires the use of sterile instruments, supplies and wound closure materials (suture, wound clips).
•Many supplies such as gloves, surgical blades, and suture material are commercially available as sterile packs. However, it is frequently necessary to sterilize, in house, items such as surgical instruments, drapes, gowns, etc.
•Sterilization kills all viable microorganisms while disinfection only reduces the number of viable microorganisms. High level disinfection will not kill the more resistant bacterial spores. Commonly used disinfectants such as alcohol, iodophors, quaternary ammonium and phenolic compounds are not effective sterilants and, therefore, are not acceptable for use on items intended to be used in survival surgical procedures.
•The following are approved sterilization procedures:
- High pressure/temperature steam sterilization using an autoclave and appropriate monitoring systems to assure sterility.
- High temperature dry heat systems. Since it is difficult to drape instruments prepared in this fashion they cannot be stored for future use. Typically instruments are sterilized and allowed to cool immediately prior to use by the surgeon.
- Gas sterilization with ethylene oxide using an appropriate gas sterilizer and appropriate monitoring systems to assure sterility and personnel safety.
- Cold (chemical) sterilization
- Alcohol with flaming (dulls instruments)
•Effective and proper use of cold sterilization is dependent on many factors including:
- The sole use of chemicals classified as "sterilants". Those classified only as disinfectants (70% alcohol) are not adequate.
- Instruments must be relatively smooth, impervious to moisture, and be of a shape that permits all surfaces to be exposed to the sterilant. Instruments tend to degrade when exposed to sterilants which requires that their integrity be assured prior to each use.
- All surfaces, both interior and exterior, must be exposed to the sterilant. Tubing must be completely filled and the materials to be sterilized must be clean and arranged in the sterilant to assure total immersion.
- The items being sterilized must be exposed to the sterilant for the prescribed period of time.
- The sterilant solution must be clean and fresh. Most sterilants come in solutions consisting of two parts that when added together form what is referred to as an "activated" solution. The shelf life of activated solutions is indicated on the instructions for commercial products.
- Instruments, implants, and tubing (both inside and out) should be rinsed with sterile saline or sterile water prior to use to avoid tissue damage.
- Examples of commercially available sterilants:
•Cidex®- active ingredient: 2% glutaraldehyde, requires many hours of immersion for effective sterilization
•Clidox®- active ingredient: chlorine dioxide, minimum of 6 hours required for sterilization
•Instruments used in pediatric or ophthalmic surgery are sized appropriately for rodent surgery3. These tend to be delicate instruments and the user should examine them prior to each sterilization to insure their integrity.
Preoperative Considerations/Care
- A healthy rodent is a prerequisite to a successful surgery. Rodents undergoing clinical or sub-clinical disease often experience anesthetic complications and are not good candidates for a successful procedure.
- It is recommended that if animals are specifically purchased for surgery that they are housed under maximum isolation conditions on arrival to insure the absence of rodent diseases.
- A minimum of 48 hours is generally required for an animal to recover from the stress of shipping; therefore, surgery should not be performed until this holding period has been completed.
- It is common in larger species to withhold food prior to surgery to prevent the possibility of aspiration pneumonia after regurgitation. This practice is not necessary in rodents since they cannot vomit.
- Fasting for four hours before surgery, however, may promote the absorption of intraperitoneally administered anesthetics .
- Water should never be withheld.
- To decrease tracheobronchial secretions, which may cause obstruction of the trachea, the use of atropine or glycopyrrolate should be considered.
- The investigator should be prepared to aspirate secretions from the trachea if necessary.
- If proper aseptic technique is utilized, antibiotics should not be necessary.
- Antibiotics, in fact, are contraindicated in hamsters and guinea pigs due to the frequent development of fatal Clostridial enteritis.
- If the interior of the intestinal tract is exposed, antibiotics are commonly administered.
- To have the desired effect, antibiotics should be administered prior to surgery to provide adequate blood/tissue levels at the time of surgery.
•Animal preparation includes preparation of the surgical site by removal of the fur by clipping, plucking or using a depilatory. An area approximately 15% larger than the area of the incision should be prepared.
•A vacuum may be used to remove the majority of the fur removed and the use of an adhesive pad will often remove any extraneous fur from the surgical site.
•Clean and aseptically prepare the surgical site by using an appropriate scrubbing technique (e.g. scrubbing in gradually enlarging circular pattern from the interior of the shaved area to the exterior) and an effective disinfectant (e.g. alternating Betadine™ or Nolvasan™ and alcohol scrubs X 3).
•The disinfectant should be in contact for a minimum of 3 minutes before the initial incision is made.
•Do not saturate other areas of the body with disinfectant since this enhances hypothermia which is a common postoperative complication in rodents.
•The surgical area should be draped with sterile drapes to avoid contamination of the incision, instruments and supplies. Opaque and non-opaque materials can be used. Clear materials have the advantage of allowing the investigator to monitor respiration and perfusion through the drape. Autoclavable plastic and sterile adhesive dressings (Steri-Drapes™) are available for use.
•It is recommended that animals be placed on a water re-circulating heating blanket or pads during surgery to prevent hypothermia.
•It is recommended that ophthalmic ointment be placed in the anesthetized animals’ eyes to prevent drying of the cornea.
•The small size of rodents precludes the use of several common methods to evaluate anesthesia used in larger species.
•For rodents, periodic observation of respiration, color of mucous membranes and loss of reflected eye color (in albino animals) will provide the surgeon with a good assessment of the animal's status.
•Except for guinea pigs, the absence of the pedal reflex is a good indication that a surgical plane of anesthesia has been attained in rodents.
•The absence of the pinna reflex is a good indicator in guinea pigs.
Multiple Surgeries in a Single Session
- It may be appropriate to segregate instruments based on potential for contamination. For example, the instruments used to incise the skin could be dedicated solely for that purpose and separate instruments utilized to manipulate exposed tissues and organs.
- Manipulate the tissues with only the tips of the instruments and avoid handling the tissues directly with your hands, which tend to be more easily contaminated.
- Using a dry bead sterilizer to re-sterilize the tips of instruments between surgeries will further insure adequate aseptic technique if precautions are taken to allow cooling of the instruments before re-use.
- Alternating 2 sets of sterile instruments provide the necessary time for instruments to sit in disinfectant for the required time. (N.B. immersion times can be long (up to 6 hours) which may preclude this as a viable option). Instruments must be thoroughly rinsed with sterile saline prior to re-use.
•After surgery the animal should be placed back in a cage that is lined with an absorbent pad. Animal bedding should not be present since the unconscious animal may aspirate bedding into the nares thereby compromising respiration.
•The most common complication that occurs during and after surgery is the development of hypothermia. This is often exacerbated by performing surgery directly on a heat conducting surface (stainless steel). This can be avoided by using sterile pads under the animal or utilizing a circulating water pad. Electrical heating pads cannot be appropriately regulated and should never be utilized as a heat source because they can cause severe thermal burns.
•After surgery an incandescent lamp (50-75 watt) can be placed 12-14 inches away from the animal to provide supplemental heat. The lamp should be positioned so that the animal can escape the light source if desired.
•Many procedures entail the loss of body fluids either through bleeding or drying during surgery. In those cases the administration of warmed sterile saline either subcutaneously or intraperitoneally will hasten the animal's recovery.
•Animals recovering from anesthesia should be monitored and rotated from side to side every 15 minutes until they are able to maintain sternal recumbancy. After full recovery, animals may be returned to their home cage.
•If surgery is to be performed on either lactating females or on pre-weaned pups, please contact the DLAR veterinary staff to discuss methods for re-introduction of pups to the dam.
1) Popp MB, Brennan MF. Long-term vascular access in the rat: importance of asepsis. Am J. Physiol. 241, H606, 1981.
2) Bradfield JF, Schachtman TR, McLaughlin RM, Steffen EK. Behavioral and physiologic effects of inapparent wound infection in rats. Lab Anim Sci 42(6):572-8, 1992.
3) Waynforth HB, Flecknell PA. Experimental and Surgical Technique in the Rat, 2nd Edition; Academic Press, 1992.
4) White WJ, Field KJ. Anesthesia and surgery of laboratory animals. Vet. Clin. North. Am. Small Anim. Pract. 17(5):989-1017, 1987.