Institutional Animal Care & Use Committee (IACUC)

When is an Animal Use Protocol Required?

  • You plan to use live vertebrate animals for teaching, testing, or research at NEOMED.
  • You receive funding administered by the University for studies of live vertebrates at another institution (i.e., the animals are owned by NEOMED or a University-based investigator).
  • You submit a grant application that includes the use of live vertebrate animals.  No animal work can proceed until a protocol has received full Institutional Animal Care and Use Committee (IACUC) approval.
  • You use invertebrates.
  • You are involved in a project at another institution which involves vertebrates for which NEOMED provides no funding, no aspect of the animal research takes place at NEOMED, the animals are not owned by the University or a University-based investigator, and the project is reviewed by that institution’s IACUC.
  • You plan to use cultured cells or tissues from vertebrates that have already been euthanized under approval by NEOMED or other IACUC or regulatory body. Because of the zoonotic potential, we require that you inform the Comparative Medicine Unit (CMU) if you are importing tissues from an outside source, such as another institution or from an abattoir (slaughterhouse).

You plan to administer or expose live vertebrates to radioactive materials, carcinogens, infectious agents, recombinant nucleic acid, highly toxic compounds (an LD50 of > 50 mg/kg), breeding of genetically engineered animals to normal animals, the cross-breeding of two lines of genetically engineered animals, radiation generating equipment,  etc.

Anesthesia

There is little published information on the effectiveness and safety of these agents in the wide range of bat species. Their inclusion here is intended only as a very general guide. Although the anesthetic regimens described below either have been used as indicated for anesthetizing bats or are generally used for many mammalian species, their utility and safety for many bat species and procedures has yet to be determined.

Injectable Agents

Ketamine + acetylpromazine – 11 mg/kg of ketamine and 1.1 mg/kg of acetylpromazine by intramuscular injection or possibly intraperitoneal injection. COMMENTS: Used for restraint in giant fruit bats, intramuscular injection may not be suitable for some small bats, intraperitoneal or subcutaneous injection may be suitable.

Ketamine + xylazine: 20 mg/kg of ketamine and 1.5-2.0 mg/kg xylazine by intramuscular injection. COMMENTS: Used in fishing bats, intramuscular injection may not be suitable for some small bats, intraperitoneal or subcutaneous injection may be suitable.

Pentobarbital: 25-50 mg/kg by intraperitoneal injection, recommend 30 mg/kg as an initial dose. COMMENTS: Sometimes combined with acetylpromazine at 1-2 mg/kg, however, if this tranquilizer is used, the amount of pentobarbital should be reduced by 1/3 to 1/2.

Inhalant Agents

Halothane: Halothane should be administered with a vaporizer. Anesthesia can be induced with 3-5 percent Halothane and maintained at 1.0-1.5 percent of inspired gas. COMMENTS: Rapid induction and changes in levels of anesthesia make Halothane dangerous when used without a vaporizer; waste gases should be scavenged for personal safety reasons; microsomal enzymes are induced to a greater degree than with methoxyflurane; suitable for prolonged procedures when administered with a vaporizer; rapid recovery.

Isoflurane: Administer with a vaporizer. Anesthesia can be induced with 3-5 percent isoflurane and maintained at 1-2 percent of inspired gas. Preanesthesia with butrophanol (5 mg/kg by intramuscular injection) may be suitable for some species of bats. COMMENTS: Rapid induction and changes in levels of anesthesia make isoflurane dangerous when used without a vaporizer, waste gases should be scavenged for personal safety reasons; suitable for prolonged procedures when administered with a vaporizer; rapid recovery; can be cardioprotective in some species.

The following list includes only the commonly used anesthetic agents in cats. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, extension 6555.

For both injectable and inhalant procedures, cats should be medicated with an anticholinergic such as atropine (0.02-0.04 mg/kg intramuscularly or subcutaneously) or glycopyrrolate (0.02 mg/kg intramuscularly or subcutaneously) to reduce salivation and maintain heart rate.

Injectable Agents

Ketamine: 10-11 mg/kg by intramuscular injection for restraint; 22-33 mg/kg for diagnostic or minor surgical procedures. COMMENTS: Poor muscle relaxation and analgesia; when used alone it is not suitable for anything more than minor surgical procedures.

Ketamine + diazepam: 5 mg/kg of ketamine combined with 0.25 mg/kg of diazepam by intravenous injection. COMMENTS: Suitable for intubation or procedures requiring light anesthesia.

Ketamine + xylazine: 2.2 mg/kg of xylazine followed in ten minutes by 11-17.6 mg/kg of ketamine, both by intramuscular injection. COMMENTS: Xylazine will induce emesis in cats; adequate muscle relaxation and analgesia for short duration mild to moderate surgical procedures, can have prolonged recovery requiring significant supportive care, inhalant agents are preferred; commonly used to anesthetize cats for training in neonatal intubation.

Pentobarbital: 20-30 mg/kg by intravenous injection, initially give 1/2 the calculated dose as a bolus and an additional amount as needed. COMMENTS: Best suited for nonsurvival procedures because of respiratory depression, poor analgesia, and prolonged often violent recovery.

Thiopental: 8-12 mg/kg by intravenous injection. COMMENTS: Suitable for only very brief procedures such as intubation prior to inhalation anesthesia; like pentobarbital it has poor analgesia and shows respiratory depression.
Local anesthesia – see Analgesics.

Inhalant Agents

Note: Although inhalant anesthetics can be delivered to cats in an induction chamber (provided the animal is adequately tranquilized), they must be administered via an endotracheal tube for maintenance. For intubation, cats can be preanesthetized with thiopental (see dose above) or inhalant agents administered via an enclosed chamber or a face mask. Please contact the CMU for instruction.

Inhalation anesthesia is augmented by the use of nitrous oxide delivered a proportion of 30-70 percent of the inspired gases. It will reduce the amount of inhalant agent required.

Isoflurane: 3-5 percent of inspired gas used for induction, 1-4 percent of inspired gas for maintenance, delivered through an endotracheal tube and a nonrebreathing apparatus with vaporizer. COMMENTS: Requires a vaporizer, waste gases should be scavenged for personal safety reasons, rapid induction and recovery; preferred for survival procedures.

The following list includes only the commonly used anesthetic agents in dogs. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, extension 6555.

For both injectable and inhalant procedures, dogs should be medicated with an anticholinergic such as atropine (0.02-0.04 mg/kg intramuscularly or subcutaneously) or glycopyrrolate (0.02 mg/kg intramuscularly or subcutaneously) to reduce salivation and maintain heart rate.

Injectable Agents

Ketamine + diazepam: 5-10 mg/kg of ketamine combined with 0.25-0.5 mg/kg of diazepam by intravenous injection; if doses in the higher range are used, half of the calculated dose is given as a bolus and the rest as needed. COMMENTS: Suitable for intubation or procedures requiring light anesthesia.

Pentobarbital: 20-30 mg/kg by intravenous injection, initially give half the calculated dose as a bolus and an additional amount as needed. COMMENTS: Best suited for non-survival procedures because of respiratory depression, poor analgesia, and prolonged often violent recovery.

Propofol: 4-6 mg/kg by intravenous injection; draw up 6 mg/kg and give slowly (over 30-60 seconds) to effect. COMMENTS: Can cause vasodilation and apnea; short duration of action – recovery usually occurs in 10-20 minutes; can be given as a continuous infusion to maintain anesthesia.

Thiopental: 8-12 mg/kg by intravenous injection. COMMENTS: Suitable for only very brief procedures such as intubation prior to inhalation anesthesia; like pentobarbital it has poor analgesia and shows respiratory depression.

Tiletamine + zolazepam (Telazol®): 4 mg/kg by intravenous injection to effect or as a bolus intramuscularly. COMMENTS: Suitable for intubation; produces deep sedation to light anesthesia.

Local anesthesia, see Analgesics.

Inhalant Agents

Note:

Inhalation anesthetics are administered to dogs via an endotracheal tube. Consequently animals must be pre-anesthetized and intubated prior to administration of the gas. Intubation is usually accomplished with the administration of a short acting barbiturate (see Thiopental above).

Inhalation anesthesia is augmented by the use of nitrous oxide delivered at a proportion of 30-70 percent of the inspired gases. It will reduce the amount of inhalant agent required.

Isoflurane: 3-5 percent of inspired gas used for induction, 1-4 percent of inspired gas for maintenance, delivered through an endotracheal tube with a vaporizer. COMMENTS: Requires a vaporizer, waste gases should be scavenged for personal safety reasons, rapid induction and recovery, preferred for survival procedures.

The following list includes only the commonly used anesthetic agents in gerbils. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, ext. 6555.

Injectable Agents

Chloral hydrate + ketamine: 350 mg/kg chlora lhydrate and 30 mg/kg ketamine by intraperitoneal injection; supplemental doses of 50 percent of the original calculated dose can be given as needed to maintain anesthesia.

Chloral hydrate + telazol: Administer 20 mg/kg of Telazol by intramuscular injection followed by 200 mg/kg of a 5 percent solution of chloral hydrate by intraperitoneal injection; anesthesia lasts 1 to 2 hours.

Ketamine + dexmedetomidine: 75 mg/kg of ketamine and 0.25 mg/kg of dexmedetomidine by intraperitoneal injection. COMMENTS: Produces approximately 30 minutes of safe, reproducible anesthesia.

Ketamine + xylazine + acetylpromazine: Mix 1.5 ml of ketamine (100 mg/ml) with 1.5 ml of xylazine (20 mg/ml) and 0.5 ml of acetylpromazine (10 mg/ml), give 0.5-0.7 ml/kg by intramuscular or subcutaneous injection. Intraperitoneal injection may also be suitable. COMMENTS: Only suitable for very minor procedures in gerbils; other combinations using ketamine are also more suited for restraint than surgical anesthesia in gerbils.

Telazol: 20-40 mg/kg provide sedation, 60 mg/kg by intramuscular injection provides surgical anesthesia. COMMENTS: Produces 4-6 hours of anesthesia and prolonged recovery, athetoid movement (involuntary movement of toes, limbs or other body parts) may occur even though the animal is anesthetized, males have more prolonged anesthesia and recovery than females.

Inhalant Agents

Isoflurane – Administer with a vaporizer. Anesthesia can be induced with 3-5 percent isoflurane and maintained at 1-2 percentr of inspired gas. COMMENTS: Rapid induction and changes in levels of anesthesia make isoflurane dangerous when used without a vaporizer, waste gases should be scavenged for personal safety reasons; suitable for prolonged procedures when administered with a vaporizer; rapid recovery; can be cardioprotective in some species.

The following list includes only the commonly used anesthetic agents in hamsters. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, extension 6555. The use of an anticholinergic (atropine, 0.04 mg/kg, intramuscularly, subcutaneously, or intraperitoneally) may help reduce salivation and maintain heart rate.

Injectable Agents

Ketamine + xylazine: 150-200 mg/kg ketamine combined with 10 mg/kg xylazine given intraperitoneally. COMMENTS: Adequate level of anesthesia for most procedures.

Pentobarbital: 50-90 mg/kg by intraperitoneal injection. COMMENTS: Decreases tidal volume and respiratory rate.

Telazol® (tiletamine + zolazepam) + xylazine: 30 mg/kg of telazol combined with 10 mg/kg xylazine given intraperitoneally. COMMENTS: Safe and reliable surgical anesthesia.

The following list includes only the commonly used anesthetic agents in mice. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, extension 6555. The use of an anticholinergic (atropine, 0.04 mg/kg, intramuscularly, subcutaneously, or intraperitoneally) may help reduce salivation and maintain heart rate.

Injectable Agents

Note: The doses of injectable anesthetic agents can vary depending upon the genetic background of the mouse. In some mice, the therapeutic dose is very close to the lethal dose. The doses provided here are intended as a guide, and they may have to be adjusted for different strains or stocks of mice.

Ketamine: 100-200 mg/kg by intraperitoneal injection. COMMENTS: Poor muscle relaxation, incomplete analgesia, not suitable for procedures requiring deep anesthesia, lasts about 30 minutes or less, time to recovery is longer.

Ketamine + acetylpromazin: 100 mg/kg ketamine + 2.5 mg/kg acetylpromazine by intramuscular injection (can be mixed together in the same syringe). COMMENTS: More muscle relaxation than #1, incomplete analgesia, not suitable for procedures requiring deep anesthesia, lasts less than 30 minutes, time to recovery is longer.

Ketamine + xylazine: 100 mg/kg ketamine + 5 mg/kg xylazine by intramuscular injection. COMMENTS: Good analgesia and muscle relaxation, suitable for more invasive procedures than #1 or #2, up to 80 minutes of surgical anesthesia, causes hypothermia, bradycardia and hypotension, can be reversed at least partially with 1.0-2.1 mg/kg yohimbine by intraperitoneal injection.

Ketamine + xylazine + acetylpromazin: Mix 1.5 ml of ketamine (100 mg/ml) with 1.5 ml of xylazine (20 mg/ml) and 0.5 ml of acetylpromazine (10 mg/ml), give 0.5 – 0.7 ml/kg by intramuscular or subcutaneous injection. Intraperitoneal injection MAY also be suitable. COMMENTS: Suitable for most surgical procedures, good muscle relaxation and analgesia.

Pentobarbital: 40-80 mg/kg by intraperitoneal injection. COMMENTS: Highly variable effect in mice depending on numerous factors including genetic background, sex, diet, environmental temperature, and type of bedding used for housing; poor analgesia; suitable for mildly invasive procedures at higher dose range; can last 20-30 minutes.

Tribromoethanol (Avertin): 0.2 ml/10 g (approximately 250 mg/kg) of a 1.25 percent w/v solution by intraperitoneal injection. COMMENTS: Produces anesthesia for about 16 minutes with full recovery in 40-90 minutes; commonly used in embryo transfer, vasectomy and distal tail amputation for Southern blot analysis; high mortality associated with toxic breakdown products if stored improperly – must be protected from light and kept at 4EC; contact the CMU for instructions on mixing.

Inhalant Agents

Carbon dioxide: Administer in an enclosed container such as a bell jar using bottled CO2, given to effect, optimal concentration is 70 percent CO2 + 30 percent O2. COMMENTS: Generally safe so long as not overdosed, suitable only for very brief procedures – one minute or less.

Halothane: Halothane should be administered with a vaporizer. Anesthesia can be induced with 3-5 percent Halothane and maintained at 1.0-1.5 percent of inspired gas. Mice are usually maintained with a face mask. COMMENTS: Rapid induction and changes in levels of anesthesia make Halothane dangerous when used without a vaporizer; waste gases should be scavenged for personal safety reasons; microsomal enzymes are induced to a greater degree than with methoxyflurane; suitable for prolonged procedures when administered with a vaporizer; rapid recovery.

Isoflurane: Administer with a vaporizer. Anesthesia can be induced with 3-5 percent isoflurane and maintained at 1-2 percent of inspired gas. Mice are usually maintained with a face mask. COMMENTS: Rapid induction and changes in levels of anesthesia make isoflurane dangerous when used without a vaporizer, waste gases should be scavenged for personal safety reasons; suitable for prolonged procedures when administered with a vaporizer; rapid recovery; can be cardioprotective in some species.

The following list includes only the commonly used anesthetic agents in pigs. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, ext. 6555.

For both injectable and inhalant procedures, pigs should be medicated with an anticholinergic such as atropine (0.04-0.05 mg/kg intramuscularly or subcutaneously) or glycopyrrolate (0.02 mg/kg intramuscularly or subcutaneously) to reduce salivation and maintain heart rate.

Injectable Agents

Ketamine + acetylpromazine: 22 mg/kg ketamine and 1.1 mg/kg acetylpromazine (can be mixed in the same syringe) by intramuscular injection. COMMENTS: Suitable for minor procedures including intubation for inhalation anesthesia; large injection volumes may make for difficult administration.

Ketamine + xylazine: 20 mg/kg ketamine and 2 mg/kg xylazine by intramuscular injection. COMMENTS: It is important to give atropine with this combination because xylazine causes heart block and hypotension in pigs; suitable for minor surgery.

Pentobarbita: 24 mg/kg by intravenous injection, given to effect. COMMENTS: Reduce dose by one-half to two-thirds when given with other agents (as is usually the case because pigs normally have to be chemically restrained for intravenous injections); apnea may occur so a ventilator should be available; for procedures lasting longer than two hours pentobarbital is undesirable due to prolonged recovery; best suited for non-survival procedures because of respiratory depression and poor analgesia.

Telazol: Administer 6.6 mg/kg by intramuscular injection. COMMENTS: Suitable for minor procedures including intubation for inhalation anesthesia; can be supplemented with Thiopental (2-3 mg/kg by intravenous injection to effect) or pentobarbital (3 mg/kg by intravenous injection to effect).

Telazol + xylazine: Dilute Telazol with large animal Xylazine (100 mg/ml) and administer the combination at a dose of 5.5-6.5 mg/kg of Telazol. COMMENTS: Suitable for minor procedures including intubation for inhalation anesthesia.

Thiopental: 6.6-8.8 mg/kg by intravenous injection. COMMENTS: Suitable for only very brief procedures such as intubation prior to inhalation anesthesia; like pentobarbital it has poor analgesia and shows respiratory depression – ventilatory support should be available.

Inhalant Agents

Note: Inhalation anesthetics are administered to pigs via an endotracheal tube. Consequently the animals must be pre-anesthetized and intubated prior to administration of the gas. Intubation is usually accomplished with the administration of Telazol (alone or in combination with Xylazine) and, if necessary, a short acting barbiturate (see Thiopental above). The dose of Thiopental is reduced by half to 2/3 when used with another agent. Intubation can be difficult in pigs and requires both training and practice. Please contact the CMU for instruction.

Inhalation anesthesia is augmented by the use of nitrous oxide delivered at a proportion of 30-70 percent of the inspired gases. It will reduce the amount of inhalant agent required.

Isoflurane: 3-4 percent of inspired gas for induction, 0.5-2.0 percent of inspired gas for maintenance, delivered through an endotracheal tube with a vaporizer. COMMENTS: Requires a vaporizer, waste gases should be scavenged for personal safety reasons, rapid induction and recovery; may be cardioprotective.

The following list includes only the commonly used anesthetic agents in rabbits. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the CMU office, extension 6555. For routine anesthetic procedures, rabbits should be premedicated with 0.1 mg/kg of glycopyrrolate to control salivation and maintain heart rate.

Injectable Agents

Ketamine + dexmedetomidine: 15-20 mg/kg of ketamine and 0.125-0.25 mg/kg of dexmedetomidine given intramuscularly. COMMENTS: 1/4 to 1/3 of the original dose can be given as needed to maintain anesthesia. Anesthesia should be reversed with Atipamizole given at a volume dose equivalent to that of dexmedetomidine.

Ketamine + xylazine: 5-10 mg/kg of xylazine by intramuscular injection followed in 10 minutes by 35-50 mg/kg of ketamine by intramuscular injection. COMMENTS: Injection volumes greater than 1 ml should be divided into multiple sites; exercise care in injecting to avoid perineural injury which is manifest as impaired reflexes and self-mutilation in the limb distal to the site of injection; higher doses are necessary to achieve surgical anesthesia; can be partially reversed with 0.2 mg/kg yohimbine by intravenous injection.

Pentobarbital: 20-60 mg/kg by intravenous injection, suggest use 28 mg/kg as a starting calculated dose, initially give 1/3 of the calculated dose and evaluate the response before giving more. COMMENTS: Barbiturates generally produce respiratory depression which is particularly profound in rabbits, the anesthetic dose and the dose that produces apnea and death are very close, one must be prepared to ventilate the rabbit if necessary, experience is important when using barbiturates in rabbits.

Thiopental: Doses vary: 12 – 25 mg/kg by intravenous injection is recommended. Initially, the animal should be given half of the calculated dose rapidly followed by the balance slowly until anesthesia is achieved. COMMENTS: You may have a narrow margin of safety in some animals depending on breed and condition. Please be prepared to intubate the animal and provide ventilatory support. Usually this method is used as a means of induction with maintenance provided by an inhalant anesthetic agent.

Ketamine + xylazine + acetylpromazine: 35 mg/kg of ketamine, 5 mg/kg of xylazine and 0.75 mg/kg of acetylpromazine all by intramuscular injection, can be mixed together. COMMENTS: Injection volumes greater than 1 ml should be divided into multiple sites; exercise care in injecting to avoid perineural injury which is manifest as impaired reflexes and self-mutilation in the limb distal to the site of injection; suitable for most surgical procedures; good muscle relaxation and analgesia.

Inhalant Agents

Note: Inhalation anesthetics are generally administered to rabbits via an endotracheal tube. Intubation may be difficult in rabbits and requires both training and practice. For intubation, rabbits can be preanesthetized with the low doses of ketamine and xylazine (given above), ketamine and dexmedetomidine, or inhalant agents administered in an enclosed chamber. Please contact the CMU for instruction.

Isoflurane: 3-5 percent of inspired gas used for induction, 1-4 percent of inspired gas for maintenance, delivered through an endotracheal tube and a nonrebreathing apparatus with vaporizer. COMMENTS: Requires a vaporizer, waste gases should be scavenged for personal safety reasons, rapid induction and recovery; may be cardioprotective.

The following list includes only the commonly used anesthetic agents in rats. A list of other anesthetics suitable for specific procedures and additional information about the effects, mode of action, metabolism, and side effects of these agents are available in the Comparative Medicine Unit (CMU) office, extension 6555. The use of an anticholinergic (atropine, 0.04 mg/kg, intramuscularly, subcutaneously, or intraperitoneally) may help reduce salivation and maintain heart rate.

Injectable Agents

Note: The doses of injectable anesthetic agents can vary depending upon the genetic background of the rat. In some rats, the therapeutic dose is very close to the lethal dose. The doses provided here are intended as a guide, and they may have to be adjusted for different strains or stocks of rats.

Ketamin: 44-100 mg/kg by intramuscular injection. COMMENTS: Lower dose produces sedation, higher dose range produces light surgical anesthesia, poor muscle relaxation, incomplete analgesia, not suitable for procedures requiring moderate to deep anesthesia, lasts about 30 minutes or less, time to recovery is longer.

Ketamine + acetylpromazine: 75 mg/kg ketamine + 2.5 mg/kg acetylpromazine by intramuscular injection (can be mixed together in the same syringe). COMMENTS: More muscle relaxation than #1, incomplete analgesia, not suitable for procedures requiring moderate to deep anesthesia, lasts about 30 minutes or less, time to recovery is longer.

Ketamine + xylazine: 40-87 mg/kg ketamine + 5-13 mg/kg xylazine by intramuscular or intraperitoneal injection. COMMENTS: Good analgesia and muscle relaxation, suitable for more invasive procedures than Ketamine or Ketamine and Acetylpromazine and those of short to medium duration, causes hypothermia and hypotension, can be reversed at least partially with 1.0-2.1 mg/kg yohimbine by intraperitoneal injection.

Ketamine + xylazine + acetylpromazine: Mix 1.5 ml of ketamine (100 mg/ml) with 1.5 ml of xylazine (20 mg/ml) and 0.5 ml of acetylpromazine (10 mg/ml), give 0.5 – 0.7 ml/kg by intramuscular or subcutaneous injection. Intraperitoneal injection MAY also be suitable. COMMENTS: Suitable for most surgical procedures, good muscle relaxation and analgesia.

Pentobarbital: 30-40 mg/kg by intravenous injection; 30-60 mg/kg by intraperitoneal injection. COMMENTS: Some variation in effect in rats depending on genetic background; poor analgesia; suitable for mildly invasive procedures at higher dose range; can last 20-30 minutes.

Inhalant Agents

Isoflurane: Administer with a vaporizer. Anesthesia can be induced with 3-5 percent isoflurane and maintained at 1-2 percent of inspired gas. Rats may be intubated (using a 14-18 gauge over the needle catheter; please see the CMU staff for specific guidance and instruction) or anesthetized with a face mask. COMMENTS: Rapid induction and changes in levels of anesthesia make isoflurane dangerous when used without a vaporizer, waste gases should be scavenged for personal safety reasons; the anesthetic concentration of isoflurane required to anesthetize hypertensive strains (e.g., SHR and Wistar-Kyoto) is significantly lower than that for normotensive rats; suitable for prolonged procedures when administered with a vaporizer; rapid recovery; can be cardioprotective in some species.

Analgesia

The following analgesics are intended for general use in the species of laboratory animals commonly used at the University. The animals’ genetic background and other factors may have a profound effect on the efficacy, safety, and incidence of side effects of these agents in these species. Although most of the information below is taken from published reports, some is based upon the clinical experience of other investigators.

Other less commonly used analgesics and combinations are available for use in the listed species and others. Blank entries indicate that no information was identified for use of the analgesic agent in the species indicated or that the agent is not well suited for the species. Please feel free to contact the Comparative Medicine Unit (CMU) office at ext. 6555 for other agents and additional information regarding effects, mode of action, metabolism, and side effects.

Download a list of commonly used analgesics [PDF]

Small Animal Survival Surgery SOPs

The United States Department of Agriculture (USDA) and the Public Health Service (PHS) require that survival surgery performed on animals be conducted using aseptic technique. The requirements of the USDA and the PHS, as defined by the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) for survival surgery in rodents and other small animals, are summarized below in the “Requirements for Small Animal Survival Surgery.” Survival surgery is defined as any surgical procedure from which the animal will recover, even if only for short periods of time.

These requirements apply to survival surgical procedures in small mammals belonging to the Order Rodentia and the specific Chiroptera (bat) species, Pteronotus parnelli, Carollia perspicillata and Antrozous pallidus with additional conditions as noted in the document. The conditions are also applicable to survival surgery in small amphibians, reptiles, and fish with appropriate modifications. Other species may be added as needed with IACUC approval.

The Institutional Animal Care and Use Committee (IACUC) recognized that unusual circumstances may necessitate the development of alternative approaches to address specific research needs. However, IACUC approval is necessary before implementation of variations from the “Requirements.”

As with any activity involving animals the individual carrying out the procedure must be adequately trained. In some cases it may be necessary for the surgeon to practice the technique on cadavers before performing it in a living animal. Please feel free to contact the Comparative Medicine Unit (CMU) at ext. 6555 or at 330. 325.6555 to arrange for guidance and practice.

In the list below explanatory information is provided in italics for several requirements. Recommendations intended to assist in the implementation of this policy and enhance the well being of the surgical patient follow the requirements.

  1. The surgery area must be dedicated to that purpose while surgery is conducted. Surgery on bats must be performed in an area dedicated solely for that purpose.
  2. The surgery table and associated equipment (e.g., stereotaxic apparatus) must be sanitized prior to use. Suitable products for disinfecting the surgery area include a disinfecting soap and water rinse, 15 percent sodium hypochlorite solution, 70 percent alcohol, and quaternary ammonium based disinfectants.
  3. Surgical instruments and implantable materials must be sterilized by an approved method. Instruments used in bat surgery must be autoclaved. Sterilization may be achieved by autoclaving (minimum 121 °C, 15 PSI, for 15 minutes), dry heat (171°C for one hour), ethylene oxide, irradiation, chemical sterilization according to the product manufacturer’s recommendations (e.g., Cidex®, or 10% providine iodine) or other means as approved by the IACUC. Autoclaving generally is the preferred method for sterilization because of its convenience, low cost, and efficacy. Depending upon the nature of the surgical procedure, the degree of instrument contamination during the surgery, and the type of animal, a sterile set of instruments may be used for up to 2-5 small animals during the same surgery session. Between animals the instruments must be decontaminated using a chemical disinfection or a point heat source such a glass bead sterilizer.  Chemical sterilants should be removed from instruments by wiping or rinsing with sterile water or saline to avoid chemical tissue damage.
  4. The surgery site must be aseptically prepped including removal of hair and disinfection by an approved method. For larger rodents such as rats or guinea pigs washing with an iodine- or chlorhexidine-based surgical soap (e.g., Betadine® scrub or Nolvasan® scrub) followed by rinsing with clean water and disinfecting with 70 percent alcohol or iodine solution is acceptable. For mice and small bats, three applications of 70 percent alcohol or two of alcohol followed by an iodine solution can be used. Animals should not be excessively wetted due to the potential for hypothermia.
  5. Sterile gloves, a surgical mask, and a clean outer garment (e.g., lab coat or scrub top – not street clothes) are required. For surgery on bats, a surgical cap is also necessary.
  6. Volatile anesthetic agents must be suitably scavenged.
  7. Animals must be monitored until they have recovered satisfactorily from anesthesia, i.e., normal respiration, sternal posture and moving about.
  8. The date and a brief description of the surgical procedure, including any drugs administered and the anesthetic agent(s) used, must be noted on the animal’s cage card.
  9. Animals must be monitored post-surgically as often as necessary to assure their well being.  Any abnormal findings must be recorded on the cage card.
  1. Covering the surgical surface with a clean paper (e.g., plastic-backed lab bench paper) or cloth will help prevent hypothermia and absorb fluids.
  2. The surgery area should be separate from high traffic areas and free of unrelated equipment and supplies. Laminar flow workbenches such as the Stay-Clean L/F Workbench by Lab Products are useful for small animal survival surgery.
  3. If possible, the surgery area should be subdivided into separate areas for animal preparation, surgery and recovery.
  1. The same methods used to sterilize surgical instruments are applicable to implantable materials. Some materials may be commercially available (e.g., polyethylene tubing) as a sterile product.
  2. Instruments and other autoclavable supplies such as gauze pads and drape material can be easily wrapped with disposable paper wraps designed for that purpose or reusable cloth towels/drapes. Autoclave confirmation tape should be used on each pack that is autoclaved. Autoclaved instruments are considered sterile for variable lengths of time depending on the manner in which they are wrapped. Double cloth wrapped instruments stored in an enclosed cabinet are recognized as sterile for up to seven weeks unless the integrity of the wrapping is compromised. Scalpel blades should be purchased sterile and not autoclaved as they are dulled by autoclaving.
  3. Surgical instruments should be cleaned with an instrument cleaner, rinsed, and dried after each surgery session. A soft toothbrush is often useful for delicate instruments. Instruments should be stored such that the cutting edges, tips, and points are protected from damage.
  1. Food and water are not usually withheld from rodents unless there is concern about ingesta within the gastrointestinal tract as may occur for abdominal surgery.
  2. Preparation of the animal is usually best done in an area close to, but separate from, the surgery area.
  3. Plucking or shaving with electric clippers is preferred techniques for removal of hair. Depilatories and razor shaving should be used carefully due to the potential for dermal irritation. Loose hair can be removed with a vacuum, tape or wet gauze.
  4. The depilated area should extend beyond the surgical margins so as to facilitate the maintenance of aseptic technique during surgery, but not so far as to contribute to hypothermia.
  5. Gauze sponges and Q-tips are convenient means to wash, rinse, and disinfect the surgical site.
  1. The surgery should be conducted so as to minimize trauma to the tissues and preserve the sterility of the instruments and the surgical field. It should be completed as quickly as possible without compromising technique; tissues should be handled delicately and, depending on the nature of the surgery, kept moist with sterile saline. Sutures and staples should not be placed too tightly. A subcuticular skin suture pattern will often preclude the chewing and removal of sutures by the animal.
  2. Whenever possible the surgery site should be draped with sterile drapes of cloth, paper, surgical gauze, or clear adhesive vinyl to minimize the risk of contaminating the surgery site. Care should be taken to avoid placing a drape such that the animal cannot be monitored. Drapes can have the added benefit of keeping the animal warm.
  3. Animals should be kept warm using an external heat source, particularly for procedures of any significant length (i.e., longer than 30 minutes). An electric or water circulating (preferred) heating pad or an overhead heat source such as a lamp can be used. Great care must be taken to prevent overheating or burning the animal. Some heating pads come with a rectal temperature probe that acts as a thermostat to turn the pad on and off.  In all cases the external heat source should be separated from the animal by a towel or other protective barrier.
  4. Also for prolonged procedures, particularly those accompanied by blood loss, warmed fluid therapy should be administered.  The recommended amount is equal to 1-2 cc per 100g body weight per hour of anesthesia plus any blood loss. Because of the small size of the patients covered by this policy, the intraperitoneal or subcutaneous routes are usually used.
  5. During the surgery the animal’s respiration, tissue color and response to noxious stimuli should be monitored so that corrective action can be taken promptly if necessary.
  1. Depending upon the nature and duration of the surgery, it may be necessary to provide the post-operative patient with an external heat source during the recovery period. As described above, steps should be taken to protect the animal from the heat source. At the very least, animals should be placed in their bedding or provided with other external insulating material (e.g., a towel). Animals recovering from anesthesia or surgery should not be recovered in a wire bottom cage.
  2. During the recovery period, the animal’s clinical condition should be monitored. Specific observations should include the body temperature, respiratory pattern, condition of the surgical wound, and strength and rate of heartbeat.
  3. Rodents can often be stimulated to breathe in the case of apnea using gentle chest compression or inflating the lungs with a rubber bulb (from a pipette) applied to their nostrils. An oxygen rich environment also may be beneficial. The drug doxapram can be used to stimulate respiration and heart beat when administered orally at a dose of 0.5-1 mg/100g. Yohimbine (approx. dose 0.1-0.2 mg/100g by intraperitoneal injection) can be used to hasten the recovery in animals anesthetized with anesthetic combinations containing xylazine.
  4. Animals cannot remain in investigator laboratories or other unapproved housing areas for longer than twelve hours without IACUC approval.
  1. Depending upon the nature of the procedure and the condition of the animal, post-surgical monitoring may range from once daily for one or two days to multiple times per day for extended periods.
  2. Conditions of observation are reviewed by the IACUC at the time of protocol review. In some cases, such as when the same procedure is conducted on many animals, alternative methods of record keeping (other than on a cage card) can be used.  Please contact the CMU at ext. 6555 or at 330.325.6555 to discuss alternative means of record keeping.

Species Characteristics

Contact

Stanley Dannemiller, D.V.M.
Director, Comparative Medicine Unit
Phone: 330.325.6558
Email: sdannemiller@neomed.edu

Institutional Animal Care & Use Committee (IACUC)

Research at NEOMED