New regulations require aseptic surgery even for rodents
The NIH has established survival surgery standards which UMass personnel are obligated to follow in order to remain in compliance with the Animal Welfare Act and Public Health Service Policy. The outlines of rules and procedures governing surgery for all warm-blooded vertebrates are described in the Guide for the care and Use of Laboratory Animals, and are nicely illustrated on a CD entitled “Training in Survival Rodent Surgery” which is available without charge by sending an email request to firstname.lastname@example.org . The IACUC recommends that all UMass students, faculty, and staff intending to perform surgery on animals view this CD before beginning any surgical procedures. In addition, these personnel must complete the surgery module of the web-based course and receive annual certification before beginning surgery.
Types of Surgery
Surgical procedures may be major or minor, and may be considered as survival or nonsurvival surgery. Major survival surgery is defined as surgery that penetrates and exposes a body cavity or produces substantial impairment of physical physiologic function. Thus aseptic technique is required. Minor survival surgery (i.e., without exposure of a body cavity) may require shorter-acting anesthesia or analgesia, but still requires aseptic technique. In all survival surgery it is of paramount importance to minimize pain and distress experienced by the animal. In nonsurvival surgery, the animal is euthanized before recovery from anesthesia. Such surgery should be done in a clean environment but need not be aseptic.
Importance of veterinary consultation before starting procedures
UMass retains a consulting veterinarian (Dr. Joanne Huyler) who is trained in surgical technique and familiar with the regulations. Dr. Huyler can be reached by contacting the Animal Care Office or by email at email@example.com. Before beginning an experiment, the PI and participating students and staff should arrange a consultation with the veterinarian in which they go through the surgical procedure. Suggestions and directives to minimize pain, distress, and the chances of infection as well as to shorten the recovery period are in the interests of the animal, the PI, and the university.
Area where surgery is done
Survival surgery should be performed in a clean and well-lit area that is isolated from foot traffic and not beneath overhanging shelves, ducts, etc. from which dust or dirt might fall into the surgical field. It is recommended that a dedicated area be devoted to surgery, although this space can be used for other purposes when surgery is not in progress. It is of paramount importance that cleanliness of the area be maintained, and disinfection using an appropriate agent (eg clidox; see Table 1) should be carried out before surgery begins.
Surgical garb needed: sterile gloves, packs
The surgeon is required to wear sterile gloves (not conventional latex laboratory gloves) while working. Instructions for hand cleaning with antibacterial soap, and the proper methods for opening the glove pack and donning the gloves are given on the NIH training CD cited above. An instrument pack which is sterilized in an autoclave should be prepared for each animal. Autoclave tape must be placed on the outside of the pack so that complete sterilization is indicated. Hot Bead, gas, or specifically approved chemical sterilization methods are acceptable alternatives (see Table 2). Note that ethanol is a disinfectant, not a sterilant (see Table 3).
A surgical mask is to be worn while performing surgery and a lab coat and surgical cap are recommended to further insure against contamination of the surgical site.
Inhalant and injectable anesthetics are used in rodents and rabbits to accomplish technical procedures causing more than momentary pain. The common inhalants (called gas anesthesia) are Halothane, Isoflurane, and Methoxyflurane. They can be delivered to the animal via a soaked pledget of cotton, or preferably, via a precision vaporizer. A precision vaporizer is especially advantageous in delivering Halothane and Isoflurane, both of which quickly volatilize to lethal levels without a vaporizer. However, a vaporizer is expensive, requires an oxygen source and a dedicated delivery system. In contrast, the inexpensive, anesthetic-soaked pledget can be extremely effective with operator experience and using due diligence in monitoring animal anesthesia depth. This method of anesthesia is suited for quick procedures where the technique requires only a few minutes and the animal is fully recovered in about 5 minutes. Although the pledget delivery system is easy and cheap, one caveat is the need for an exhaust hood in which to perform all manipulations.
Injectable anesthetics are extremely cheap and easy to use, but some of them require a Controlled Substance license, along with additional record keeping. Contact the Director of Animal Care for information and forms to obtain this license. Sometimes a combination of drugs formulated into a “cocktail” is most efficacious for a particular species (Table 5). These drugs are calculated on the animal weight, so record these figures on the Anesthetic Record, along with the administered amounts of anesthetic.
Assessment of plane of anesthesia; record keeping before, during, after surgery
Regulations require that thorough records be kept on each animal on which survival surgery is performed. These records are to be kept near the surgical area and should include the approved protocol number under which the surgery is performed. Record keeping forms are available on the UMass animal care web site and may be downloaded for your use. It is essential to record the animal ID, starting body weight, time at which anesthetic is given, dose of anesthetic, and the animal's response. The animal must achieve a sufficiently deep plane of anesthesia, as assessed by 3 reflexes (typically failure to respond to ear-pinch, toe-pinch, or eye contact [palpebral/conjunctival reflex]) before surgery can begin, and this must be recorded in the surgical record form. If the animal becomes sensitive to pain as the surgery proceeds, the surgeon must stop and administer supplemental doses of anesthetic. S/he must wait until these responses cease before resuming surgery, and the supplemental doses must be recorded on the record form.
Once surgery is complete and wound closed, the time should be noted and analgesics given as appropriate (see below). The time, route, and dose of analgesia administration must be recorded. The animal must be monitored until it resumes activity; its postoperative status should be checked periodically for a minimum of 24h and noted in the record form. It is advisable to record body weight on a daily basis in order to assess whether the animal has resumed eating.
Cleaning of wound site
The wound site should be cleaned by shaving fur or plucking of fur or feathers. This should be done away from the surgical area and before donning gloves to avoid contamination of the area or the surgeon. The wound site should be cleaned with iodophor (e.g., Betadine) or other suitable disinfectant (see Table 4), followed by 70% ethanol. Good technique specifies beginning in the middle of the surgical field and working outwards in a surgical motion; do not move the gauze or swab from a dirty peripheral area to a clean one. This is illustrated in surgery guide disc.
The wound site should be draped before surgery begins. A sterile adhesive drape may be used, or autoclaved paper or cloth drapes may be affixed using towel clamps.
Protecting the animals' eyes
After anesthetic administration, it is advisable to apply a bland ointment to both eyes in order to prevent drying with the loss of blink reflex. It is imperative to monitor anesthetic depth during the induction period, surgical preparation, as well as during the procedure. If pinching a toe, tail, or ear results in a reaction indicating that the animal is too light, either wait a while for the full dose to take effect or give a small additional amount. Also, monitor mucous-membrane color, as well as the respiratory pattern and frequency (can give a dose of atropine if increased fluids are noted in the lungs).
The selection of the type and size of suture material should be done in advance in consultation with the veterinarian, Dr. Huyler. Selection is based on the type of surgery and species of animal. Common suturing methods used in rodents are described and illustrated in the NIH training disc (Module 2). Recommended types of suture are listed in Table 10.
For small animals, a 3 aught (3-0) suture thickness or smaller (4-0 to 6-0) are best. In addition to size, sterile suture material is sold with specific needle types fixed to the suture. Cutting and reverse cutting needles have sharp edges and are best used for skin suturing. Non-cutting, atraumatic, taper or round needles are used for suturing easily torn tissues such as peritoneum, muscle, nerve or intestine. Ligation of vessels or suturing of tissues other than skin requires an absorbable material such as Vicryl®, Dexon®, PDS®, Maxon®, or chromic (see Table 10). For skin closure, non-absorbable suture such as Prolene® or nylon may be used. Stainless steel wound clips are recommended for skin closure because most rodents will gnaw at skin sutures. Veterinary grade cyanoacrylate surgical adhesives, such as Vetbond® or Nexaband®, may be used to close incisions or the area between sutures. The use of silk, a non-absorbable suture material, is not recommended for skin closure. Silk can cause tissue reactions and may wick microorganisms into the wound. It is best used for cardiovascular procedures only.
Prevent hypothermia and dehydration during and after surgery
Once an animal is anesthetized, it loses its ability to thermoregulate so hypothermia is a frequent complication. Hypothermia can be combated by using external heat sources such as a slide warmer, circulating hot-water pad, and chemical (Safe and Warm®) or microwave-activated heat sources. It is advisable NOT to use an electric heating pad since dermal burns can result. Also, if you use a heat lamp as an external heat source, be sure to place a thermometer next to the animal(s) and regulate the temperature between 85-95 degrees F (29-35 degrees C).
Dehydration is another complication that can be countered during the post-op period. Hydration status is assessed by pinching the skin and forming a tent. In an adequately hydrated animal, the tented skin returns quickly upon release, but if the skin is slow to return, some degree of dehydration is indicated. Rehydration is facilitated with either subcutaneous or intraperitoneal administration of warmed Lactated Ringers Solution or Saline (Table 6). The veterinarian should be consulted for appropriate type, volume and route of administration.
Post-operative pain must be allayed with an analgesic, especially if a major body cavity (thorax, abdomen, cranium, joint capsule) was entered. Studies show that analgesics administration via the drinking water are ineffective. Animals either demonstrate a pain-associated avoidance, or are not able to drink a sufficient volume to effect analgesia. Therefore, an analgesic must be administered by either injection or gavage. Table 7 gives potential signs of pain, and Table 8 gives suggestions for specific analgesics for various procedures. Table 9 is a comprehensive list of anesthetics and analgesics for rodents and rabbits. Note that the time and dose of administration of post-operative analgesics as well as the original and supplemental doses of anesthesia must be noted in the surgery and/or postoperative records.
Strategies to obtund pain include administering an opiate, a Nonsteroidal Anti-inflammatory Drug (NSAID), incision line infiltration, and even topical creams. These drugs may be given singly or in combination, depending upon the procedure and protocol requirements. What ever parenteral analgesic is chosen, the initial dose should be administered while the animal is still under anesthesia, but not yet fully recovered. You will note from Table 9 that most opiate analgesics must be administered every 2 to 4 hours. The exception is Buprenorphine that has a longer half life than most. As a result, many scientists are using Buprenorphine for the initial injection, and repeating the dose every 8 to 12 hours until the animal appears to function in a pain-free manner. Finally, remember to annotate drug administrations on the Post-operative Record.
Part of the recovery process is permitting access to highly palatable food. This might be as simple as soaking the regular chow so it is softer and easier to consume. You might add Nutrical to the diet, giving a high energy boost. Be sure the water bottle is functioning properly and is easily reached by a recovering animal.
Occasionally, a protocol may call for the use of an antibiotic(s). If so, contact the Attending Veterinarian at the Animal Care Office , for an appropriate antibiotic and dosage.
Contact the Veterinary Technician, at the Animal Care Office , for drugs and supplies that can facilitate your surgical endeavors.
Animals should be free from the effects of anesthetic before they are returned to their animal facility. Cage cards should be annotated with the date and type of procedures completed. Animals should be monitored for several days following surgery. Measures of food intake where possible, daily body weights and behavior should to be assessed. Providing soft, easily accessible foods may encourage eating in recovering animals. Note, however, that the administration of some necessary analgesics may suppress appetite. Suture or wound clips should be removed within 10 –14 days after surgery. Small scissors should be used with sutures. Specialized instruments are available for removal of wound clips.