Basis

Recommendations for the performance of survival surgery on mice, rats, and birds are based on the 2010 edition of the NIH Guide for the Care and Use of Laboratory Animals (the Guide, pp. 115-119, 144-145) and the USDA Animal Welfare Act Regulations (AWARs) §2.31(d)(1)(ix) and §2.33(a)(5).

Note: For the purpose of these guidelines, the term "rodent" refers to mice of the genus Mus and rats of the genus Rattus. For information on survival surgery in warm-blooded species other than rats, mice, or birds, please refer to the ARC Policy on Survival Surgery in USDA-Covered Species.

Definitions

Aseptic Technique

Aseptic technique is used to reduce microbial contamination to the lowest possible practical level and includes preparation of the animal, such as hair removal and disinfection of the operative site; preparation of the surgeon, such as the provision of decontaminated surgical attire, surgical scrub, and sterile surgical gloves; sterilization of instruments, supplies, and implanted materials; and the use of operative techniques to reduce the likelihood of infection.

Survival Surgery

A survival surgery is any surgical procedure from which an animal regains consciousness for any period of time.

Major Survival Surgery

A major survival surgery is any survival surgical procedure that penetrates and exposes a body cavity, produces substantial impairment of physical or physiologic functions, involves chronic (long-term) indwelling intravascular catheterization, or enters the cranial cavity. Examples of major surgery include laparotomy, thoracotomy, craniotomy, joint or bone replacement, spinal transection, and limb amputation.

Multiple Survival Surgeries

Multiple survival surgical procedures are not permitted on animals unless scientifically justified. For additional information, please refer to the ARC Policy on Multiple Survival Surgeries.

General Guidelines

Location of Surgery

A surgical area for rodents and birds can be a room or portion of a room that is easily sanitized and not used for any other purpose during the time of surgery. However, because dedicated surgical facilities provide advantages with regard to veterinary oversight, the ARC requires the use of a dedicated surgical facility, such as the DLAM Rodent Procedure Room, whenever possible. An investigator's laboratory may be used as a survival surgery area provided such use is scientifically justified by the investigator and the location is inspected and approved by the ARC. In selecting a surgical location, the investigator should bear in mind that "the number of personnel and their level of activity have been shown to be directly related to the level of bacterial contamination and the incidence of post-operative wound infection" (the Guide, p. 144). Thus, every attempt should be made to sufficiently separate the surgical area from other areas in the room to minimize unnecessary traffic and decrease the potential for contamination of the wound. Further, the location should be designed to include the following three areas.

  1. An area should be designated for preparation of the animal, including weighing, hair or feather removal, and initial skin disinfection. The prep area should be sufficiently separate from the surgery table to minimize the potential for contamination of the surgery area by aerosols generated during animal preparation.
  2. A separate area should be set aside for the conduct of the surgical procedures (i.e., from skin incision to wound closure). The surgical table and immediate surrounding areas must be constructed of a material that can be washed with soap and water and then disinfected using appropriate agents (see Table 1 below). The immediate surgical area should be disinfected prior to and between surgeries to decrease dust-borne contamination and may not be used for other purposes during the time of surgery.
  3. Finally, a separate recovery area should be established. This should be a quiet, undisturbed location where the animals can be observed.

Table 1. Recommended Hard Surface Disinfectants

(e.g., table tops, equipment) Always follow manufacturers’ instructions.

Agents Examples Comments
Alcohols 70% alcohol (isopropyl is recommended) Minimum contact time required is 15 minutes. Contaminated surfaces take longer to disinfect. Remove gross contamination before using. Inexpensive. Flammable.
Chlorine

Sodium hypochlorite (Clorox® 10% solution)

Chlorine dioxide (Clidox®, Alcide®)

Corrosive. Presence of organic matter reduces activity. Chlorine dioxide must be fresh (<14 days old). Kills vegetative organisms within 3 minutes of contact.
Aldehydes Gluteraldehyde (Cidex®, Cide Wipes ®) Rapidly disinfects surfaces. Toxic. Exposure limits have been set by OSHA.

Surgical Instruments

Instruments must be sterile at the start of any surgery day, and/or between groups of animal subjects. A list of acceptable methods for instrument sterilization is included in Table 2 below.

Table 2. Recommended Methods of Instrument Sterilization

Always follow manufacturers’ instructions.

Agents Examples Comments
Physical:
Steam sterilization (moist heat)
Autoclave Effectiveness dependent upon temperature, pressure and time (e.g., 121.6°C for 15 min. vs. 131°C for 3 min.).
Chemical:
Gas sterilization
Ethylene Oxide Requires 30% or greater relative humidity for effectiveness against spores. Gas is irritating to tissue; all materials require safe airing time. Carcinogenic. Suitable for catheters and implants.
Chlorine Dioxide Clidox®, Alcide®
Aldehydes Gluteraldehyde Many hours required for sterilization. Corrosive and irritating. Consult Biosafety Officer on proper use. Must be thoroughly rinsed from instruments using sterile distilled water before use.
Dry heat: Hot bead sterilizer Fast (15 to 20 seconds). Instruments must be cooled before contacting tissue. Instruments must be sterilized by one of the above methods prior to first use that day.

Surgical instruments may be used on more than one animal; however, any item used on multiple animals must be carefully cleaned and disinfected between animals (see Table 3 below). Hot bead sterilizers are preferred for this purpose; users must ensure that all blood/tissue debris is wiped from the instrument tip(s) before insertion into the glass beads. (Please refer to the manufacturer’s instructions for the specific model you have to ensure proper usage.)  Soaking in 70% alcohol or disinfectant is also acceptable. Because the effectiveness of disinfection is directly dependent upon the contact time with the disinfectant, the surgeon should anticipate the number of surgical instruments required to guarantee uninterrupted conduct of the procedures while affording ample contact time. Disinfectants should be replaced when contaminated with body fluids or tissues.

Table 3. Recommended Instrument Disinfectants

Always follow manufacturers’ instructions.

Agents Examples Comments
Dry heat Hot bead sterilizer Fast. Instruments must be cooled before contacting tissue.
Alcohols 70% alcohol Minimum contact time required is 15 minutes. Contaminated surfaces take longer to disinfect. Remove gross contamination before using. Inexpensive. Flammable (do NOT use a flaming technique with alcohol to disinfect instruments).
Chlorine Sodium hypochlorite (Clorox® 10% solution)Chlorine dioxide (Clidox®, Alcide®) Corrosive. Presence of organic matter reduces activity. Chlorine-based disinfectants must be fresh (<14 days old). Kills vegetative organisms within 3 min. Must be thoroughly rinsed from instruments using sterile distilled water before use.
Peracetic Acid/ Hydrogen Peroxide Spor-Klenz® Corrosive to instrument surfaces. Must be thoroughly rinsed from instruments using sterile distilled water before use.
Aldehydes Gluteraldehyde Minimum contact time required is 15 min. Corrosive and irritating. Consult Biosafety Officer on proper use. Must be thoroughly rinsed from instruments using sterile distilled water before use.

Monitoring Sterility

Indicators of sterility must be included inside the pack or pouch containing the items being sterilized. These are commercially available strips which change colors to indicate that the internal area of the pack has reached temperatures of sterilization or the gas has penetrated the pack. The date of sterilization must be written on the outside of the pack or pouch.

The autoclave must also be biologically monitored on a quarterly basis. This involves placing a tube containing bacterial spores within a pack. The tubes are removed after the sterilization process and incubated to look for inadequate sterilization. These monitors can be obtained from the DLAM lab.

Sterilization Logs

A log must be maintained showing the dates the instruments were autoclaved. The actual internal pack sterilization monitor, showing adequate sterilization, must be attached to this log.

Pre-Surgical Evaluation & Treatment

Pre-existing health conditions may negatively affect the immediate or long-term success of the surgical procedures. Performing pre-surgical evaluations will help ensure that the animals are not overtly ill. This should include visual inspection of the animal and assessment of the behavioral status of the animal. The animal should be alert and behaving normally, and should have a smooth coat and clear eyes. Physical or behavioral abnormalities must be brought to the attention of the DLAM veterinary staff. Surgery must not be performed on animals exhibiting these signs without prior consultation with a DLAM veterinarian.

Withholding food or water is generally not necessary in rodents or birds unless specifically mandated by the protocol or surgical procedure (e.g., gastrointestinal surgery). Withholding food or water for more than six hours should be discussed with a DLAM veterinarian. Due to the neophobic nature of these species, special diets or drinking water should be introduced to the animals prior to the surgical procedure (up to 1 week in advance for the diets, and 1-2 days in advance for water, including antibiotic therapies).

In some cases, it may be preferable to initiate antibiotic treatment prior to surgery. It is required that the first dose of analgesics be administered prior to making the first surgical incision; please refer to the ARC Policy on Post-operative Analgesia. All antibiotic or analgesic treatment regimens should be discussed with a DLAM veterinarian.

Surgical Preparation

Preparation of the animal should include removal of hair/feather from the surgical site with a generous border (at least 1 cm) to avoid contaminating the incision site. Hair or feather removal should be performed in a location separate from the surgical area. The surgical site should be scrubbed with a povidone-iodine scrub (e.g., Betadine®, Nolvasan®), being careful to scrub from the center of the site toward the periphery. The site should then be rinsed with 70% alcohol. At least three alternating preparations of germicidal scrub and rinse are considered adequate. Finally, the area should be draped with sterile drapes, which will not only help to prevent contaminants from entering the surgical field, but will also provide a sterile area on which to lay sterile instruments during surgery.

The surgeon must thoroughly wash his or her hands with a bactericidal scrub. The use of sterile surgical gloves is required. Gloves dipped in bleach are not acceptable for this purpose. A surgical mask must be worn for major surgeries but is also recommended for minor procedures. Wearing a clean lab coat is mandatory; however, a sterile gown is preferable, especially for major surgeries.

Anesthesia

The anesthetic regimen for any surgical procedure should be determined in consultation with a DLAM veterinarian and must be described in the approved research protocol. Generally, gas anesthesia (e.g., isoflurane) is recommended for longer procedures that would require multiple injections of anesthesia. In any case, it must be determined that the animal is fully anesthetized prior to initiating the procedure and that a consistent plane of anesthesia is maintained throughout the duration of surgery. Anesthetic depth may be monitored in a number of ways (e.g., respiration rate, muscle relaxation, lack of response to toe pinch, etc.) and may vary depending upon the species and the anesthetic agent used. Anesthetic depth must be continuously monitored throughout the procedure.

For guidance in selection and use of anesthetics, please contact a DLAM veterinarian.

Surgical Procedures

All surgical procedures must be conducted as described in the approved protocol. Evaluation of the animals during surgery is critical. In addition to monitoring anesthetic depth as described above, maintaining normal body temperature is of particular importance, as anesthetics can directly or indirectly induce hypothermia. Water-circulating heat pads are recommended for this purpose. The use of electric heat pads may overheat the animal; if these are used, the pad should be set on low, a light cloth covering should be placed between the animal and the pad, and the animal must be observed frequently for signs of hyperthermia. Because heat lamps may cause severe hyperthermia or other thermal injury, their use is prohibited.

To prevent corneal desiccation, bland ophthalmic ointment should be placed in the eyes following the onset of anesthesia and prior to clipping or shaving any fur or hair, or plucking of feathers. If animals will undergo survival stereotaxic surgery, blunt ear bars must be used to prevent damage to the tympanic membrane.

Paralytic agents may not be used without anesthesia. If a neuromuscular blocking agent is required for the surgical procedures, please refer to the ARC Policy on Neuromuscular Blocking Agents.

Suture Selection

Closure of the internal tissues should be completed using an absorbable suture material. A nonabsorbable monofilament suture material should be used for skin closure. Placement of subcuticular sutures, although more technically challenging, is acceptable for skin closure and may be performed using absorbable materials. The smallest gauge suture material should be used as practicable. Lists of acceptable suture materials and their gauges are is included in Tables 4 and 5 below. Please note that these are general guidelines and suture size will be dependent on the size of the animal and the location of the surgery. More specialized surgery, such as microsurgery, will require smaller sutures. Please contact a DLAM veterinarian for more personalized recommendations.

Table 4. Acceptable Suture Materials

Suture Characteristics and Frequent Uses
Vicryl®, Dexon® Absorbable; 60-90 days. Suitable for internal wound closure.
PDS®, Maxon® Absorbable; 60-90 days. Suitable for internal wound closure.
Prolene® Nonabsorbable. Suitable for skin closure.
Nylon Nonabsorbable. Suitable for skin closure.
Stainless Steel Wound Clips, Staples Nonabsorbable. Suitable for skin closure. Requires instrument for removal from skin.

Table 5. Recommended Suture Gauges/Sizes

Species Location or Function
Mouse

Abdominal or peritoneal area: 4-0 to 5-0
Skin (subcuticular): 5-0 to 6-0
Skin (external): 4-0 to 6-0

Rat Abdominal area: 4-0 to 5-0
Skin (subcuticular): 4-0 to 6-0
Skin (external): 3-0 to 5-0
Bird 3-0 to 6-0 (depending on animal size)
Skin (external): as for mice

Because silk and chromic gut may cause tissue inflammation, these materials are not acceptable for wound closure.

Sutures, staples, or wound clips must be removed 7-14 days following surgery. If animals will be euthanized within 14 days following surgery, removal of sutures prior to euthanasia is not necessary. Any foreign substance left in the incision for a long period of time serves as a nidus of irritation and infection. A DLAM veterinarian should examine incisions that do not appear to be healing.

Post-Operative Recovery

Observation during post-surgical recovery is imperative. The animal, whether it will recover in or out of its cage, must be kept warm. Water-circulating heat pads are recommended for this purpose. The use of electric heat pads may overheat the animal; if these are used, the pad should be set on low, a light cloth covering should be placed between the pad and the animals/cages, and the animal must be observed frequently for signs of hyperthermia. Somnolent animals should be turned periodically to prevent burns or other thermal injury. Provisions must also be made so that a conscious animal can escape the heat source when it becomes too warm. Because heat lamps may cause severe hyperthermia or other thermal injury, their use is prohibited.

A recovering animal must be watched continuously until in sternal recumbency and able to ambulate. Unconscious animals must not be left unattended. To prevent undue risk, rodents should be housed individually following surgery until suture/wound clip removal or two weeks after the surgical procedure if there is no need to remove wound closures.

Post-Operative Analgesia

As described in the ARC Policy on Post-Operative Analgesia, analgesia must be provided to all animals following survival surgery, unless scientific justification for withholding such agents is approved by the ARC as part of the investigator’s research protocol, or if a veterinarian examines the animal and determines that analgesic administration is no longer necessary.

Pain has been defined by the International Association for the Study of Pain as "[a]n unpleasant sensory and emotional experience associated with actual or potential tissue damage or described in terms of such damage. It is always subjective."

The fact that different anatomical and neurobiological substrates for pain exist supports the use of targeted pain therapy and explains the variable effectiveness of analgesic drugs depending upon the characteristics of pain. In the research laboratory setting the characteristics of pain will vary with the particular animal (species, age, strain) and the type of potentially painful procedure.

The following lines reflect a brief description of the mode of action, indication, contraindication and side effects of the most commonly used type of analgesics in the UCLA animal facilities. For alternative options and for dosage and regimen recommendations please contact a DLAM veterinarian.

  1. p-Aminophenol Derivatives – Included in this group is acetaminophen. Acetaminophen has analgesic and anti-pyretic activity with weak anti-inflammatory action. This drug is contraindicated in cats due to increased sensitivity to toxicosis. Methemoglobinemia and hepatotoxicity characterize acetaminophen toxicosis. Available in the form of tablets, capsules, suppositories, chewable tablets, wafers, elixirs and solutions, in combination with other drugs. Effective against pain of low to moderate intensity; recommended only for minor surgical procedures.
  2. Non steroidal anti-inflammatory drugs (NSAIDS) - Agents that reduce inflammation and provide analgesia primarily by inhibiting one or more steps in the metabolism of arachidonic acid through inhibition of inflammatory cytokine synthesis. NSAIDs do not contain the molecular steroid ring structure and don’t have the immunosuppressive and metabolic side effects associated with corticosteroids. Most NSAID act primarily to reduce the biosynthesis of prostaglandins (PG) by inhibiting cyclooxygenase (COX). NSAIDs most commonly used in veterinary medicine include:
    1. Propionic acid derivatives – Ibuprofen, ketoprofen and carprofen are included in this group. Ketoprofen and ibuprofen are available in parenteral form. Carprofen and ibuprofen are available in oral tablets. Ketoprofen and ibuprofen inhibit both isoforms of cyclooxygenase (COX-1 and COX-2), with ketoprofen having a higher selectivity for COX 1 than ibuprofen in dogs. Carprofen is claimed to be predominately COX-2 selective drug, but it does inhibit COX-1 slightly. Efficacy against orthopedic and postoperative pain compares favorably in clinical studies with some opioids such as buprenorphine. Drugs in this group can have GI side effects including GI erosion and altered permeability. Most of these drugs can alter platelet function and altered bleeding time. Carprofen and Ketoprofen did not affect bleeding time in dogs based on previous studies. Carprofen has been reported to cause minor changes in certain renal parameters, but most studies have not observed significant risk to renal function.
    2. Oxicams – The most commonly used drug used in veterinary medicine in this group is Meloxicam. Meloxicam is available in parenteral, tablet and oral suspension. This drug is a selective inhibitor of COX-2 in humans and dogs and non selective in cats. It is as effective an analgesic as buprenorphine for post-operative pain resulting from soft tissue surgery. Meloxicam does not affect bleeding time in dogs, however there’s no current evidence of this in other species. GI side effects and other typical NSAID toxicities have been reported.
    3. Nicotinic acid derivatives – Flunixin meglumine is included in this group of drugs. This drug is available in oral and parenteral forms. It demonstrates COX-1 selectivity in dog and horse blood. Flunixin is a potent analgesic agent after parenteral administration in mice, rats and monkeys. This drug demonstrates potency similar to that of other NSAIDS such as ketoprofen and carprofen. The most common potential side effect is gastroduodenal ulceration and perforation.
  3. Opioids - Opioids are the most effective analgesics available for the systemic treatment of acute pain in many species. Opioids combine reversibly with specific receptors (mu, delta and kappa) in the brain, spinal cord, and periphery, altering the transmission and perception of pain. The clinical effects of opioids vary between the µ opioid receptor agonist, partial mu agonists, and agonist-antagonists. Repeated use of opioids may induce tolerance in some species. The most common side effects associated with the use of opioids include respiratory depression, constipation, sedation, euphoria, dysphoria, excitement and pica. Buprenorphine is a partial mu agonist and kappa antagonist that is widely used in Laboratory animal medicine due to its relatively long duration of action. Buprenorphine is available in parenteral formulation.

Antibiotic Treatment

Post-operative antibiotic treatment should be discussed with a DLAM veterinarian to determine whether routine administration of antibiotics is necessary. In general, post-operative antibiotics should be provided if the animal will survive long enough to develop severe infection, but may also depend upon other factors such as the invasiveness of the procedure and the immune status of the animals. The administration of antibiotics prior to commencing a procedure can further minimize post-operative infections.

Post-Operative Treatment Cards

Every cage containing a post-operative mouse or rat must have displayed on it a post-op treatment card or sticker, placed on the back of or behind the regular cage card (see below). Treatments (such as analgesics and antibiotics) following procedures must be indicated on this form until treatment is discontinued (for example, until the drinking water containing antibiotics is changed out for fresh water) or until any sutures or wound clips are removed, whichever is longer. These cards are available either from the DLAM Pharmacy (for a fee) or the template can be downloaded from the DLAM website, under Technical Operations (file name: Timed Post-Op Labels), and printed onto adhesive labels (such as OfficeMax® White Multipurpose labels, Order No. A5OM97793, Size 2x4, or Avery® 5163) or other cards for use on the cages. Please do NOT apply the adhesive labels directly to the cage surfaces, but rather to the back of the cage card or onto another card that will be placed in the cage card holder.

Post-Operative Treatment Card

Long-Term Recovery and Monitoring

Post-surgical observations include a minimum daily observation, including weekends and holidays, of the condition of the animal and the surgical site. Animals should be observed for continued recovery, which may include state of arousal; indices of pain or discomfort; condition of the surgical wound; appetite; hydration status; capillary refill time; mucous membrane color; or fecal and urine production.

Some surgical manipulations may require an extended period of post-operative monitoring. The appropriate duration and extent of monitoring can be determined by the DLAM veterinary staff in consultation with the investigator. Some situations that may constitute prolonged monitoring periods include animals with chronic debilitating disease states (e.g., diabetes mellitus), animals undergoing organ transplantation or immunosuppressive therapy, and animals with chronically implanted instruments or catheters.

The on-call veterinarian pager number (pager #96545) should be kept readily available in the event that post-surgical complications are observed.

Record-keeping Requirements

In accord with recommendations of the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) International, surgical records are required for rodents and birds. These records should include the administration of anesthetics, fluids, and any drugs; details of the procedure, including intra-operative monitoring; daily post-operative recovery observations and treatment, including administration of analgesics and antibiotics; monitoring of incision healing, including suture/staple removal if applicable; and the initials of the individual performing these tasks. All medications, including the name, dose, route, and time of administration should be recorded. Additionally, any adverse outcomes should be noted. A sample "Surgical and Post-Operative Evaluation Record for Mice, Rats, and Birds" is included with these guidelines.

So that the veterinary staff can better evaluate the post-operative healing process and to ensure that sutures are appropriately removed, each cage card should be clearly marked with the date of surgery, and the post-op treatment card used (see Post-Operative Recovery section, above). If the research staff will administer antibiotics in the drinking water, a blue "Investigator Will Feed/Water/Medicate" tag must be placed on each cage to communicate this special care to DLAM staff (if the animals are being housed in a DLAM-managed area).

All records relating to surgical procedures and post-operative care may be subject to review during inspection or audit.

References

  1. Fish, R.E.B., Marilyn J.; Danneman, Peggy J.; Karas, Alicia Z., Anesthesia and Analgesia in Laboratory Animals. 2nd ed. American College of Laboratory Animal medicine Series, ed. A. Press. Vol. 1. 2008, Amsterdam, Boston, Heidelberg, London, New York, Oxford, Paris, San Diego, San Francisco, Singapore, Sydney, Tokyo. : Elsevier. 268-275.
  2. MERCK, Nonsteroidal Anti-Inflammatory Drugs, in The Merck Veterinary Manual, S.E. Aiello, Editor. 2005, Merck & Co., INC. New Jersey. p. 2131-2137.
  3. Sam McMillan, V.A., DAVN(Med), RVN. NSAIDS: What are the Options? in British Small Animal Veterinary Congress. 2008. Birmingham, UK: Quedgeley : British Small Animal Veterinary Association 2008.
  4. Bill, R.P. NSAIDs--Keeping Up With All the Changes, in Atlantic Coast Veterinary Conference 2008. 2008. Atlantic City, New Jersey.
  5. Budsberg, S.C. Review of NSAIDS: COX Selectivity and Systemic Effects Beyond Analgesia (S22C). in Western Veterinary Conference 2009. 2009. Las Vegas, NV.

Approved 9/22/03; Revised 11/24/03, 6/14/10; Updated 1/18/11; Revised 6/23/14; Revised 8/26/14

Replaces Rodent Surgery Guidelines 6/02