Rationale:

An integral component of veterinary medical care is the prevention and alleviation of pain associated with procedural and surgical protocols.  The selection of the most appropriate anesthetic should reflect professional judgment as to which best meets clinical and humane requirements without compromising the scientific aspects of the research protocol.

The choice of anesthetic will depend on the procedure, the length of the procedure, the equipment available, and the expertise of those administering the anesthetic.  There are a number of options available for rodent anesthesia, including injectible and inhalation anesthetic agents.  The standards will be listed and discussed within this policy, as will standards of monitoring and recovery of the anesthetized animal.

Policy:

It is required that written documentation be kept on use of anesthetics, analgesics and tranquilizers by the Principal Investigators in the form of Animal Use Records.  If injectible anesthetics are being utilized, keep a record of the anesthetic cocktail that you are using as well as how you are dosing the animals (route, i.e. IP, SQ or IV, and how many mls per animal).  This record must be kept in a place that is easily accessible by ARC veterinarians, IACUC, and Environmental Health and Safety personnel. If inhalation anesthetics are being utilized, ensure that the liquid forms of the anesthetics are not out of date and are available for review by the above mentioned personnel.

Inhalation anesthetics:

Independent of the use of precision calibrated vaporizers or the open drop technique, proper disposal of waste gases should be determined prior to use.

• Waste gases must not be vented into the room.
• Acceptable scavenging of waste gases include:
• 1. the Use of a downdraft table (Be sure to not cover the downdraft table, as this will cause a loss in its ability to effectively scavenge gases. Also note that downdraft tables are usually only effective up to 6-8 inches from the surface, so induction chambers should not be taller than this.),
  2. working in a chemical fume hood,
  3. working in a type IIB Biosafety cabinet ducted to the outside, or
  4. the use of Charcoal canisters (Canisters must be weighed regularly and replaced when they increase by 50 grams in weight.).
• When using an anesthesia induction chamber, the system must be flushed for a minimum of 10-15 seconds after use to evacuate all anesthesia-saturated gas from the chamber.
• When using a bell jar, waste gas should be vented outside of the building by a fume hood or type IIB Biosafety cabinet, or can be fitted with an active waste gas scavenging system.

Precision Calibrated Vaporizer Anesthesia systems

Open Drop Technique

1.       A bell jar or induction chamber can be created by using a large covered glass or plexiglass container with anesthetic soaked cotton balls or gauze squares located at the bottom.  The mouse or rat must be prevented from coming into direct contact with the liquid inhalant by placing a mesh grid over the cotton/gauze, or by placing the cotton/gauze in a meshed container.

2.       Because the anesthetic concentration cannot be controlled and can rapidly reach toxic or fatal concentrations, this method must be reserved for very short-term procedures such as retro-orbital bleeding.

3.       Inspect and clean bell jars immediately before and after use with a suitable disinfectant to prevent cross-contamination.

Injectible Anesthetics

• Injectible anesthetics are widely used in rodent surgeries.  The use of controlled substances requires additional record keeping.
• There are four basic routes of administration for injectible anesthetics.  They are intraperitoneal, subcutaneous, Intramuscular, and intravenous.

• 1. Intraperitoneal (IP): For mice, hold the animal in a head-down position and insert a 22-25 guage needle into the lower right or left abdominal quadrant just off of midline.  For rats, hold the animal in a head-down position and insert a 20-22 gauge needle into the lower left abdominal quadrant just off of midline.
  2. Subcutaneous (SQ or SC): For mice and rats, tent the skin of the back and inject into the space between the skin and underlying muscle.
  3. Intramuscular (IM): Intramuscular injections are NOT recommended in mice. For rats, injections are usually delivered into the caudal thigh muscles or the muscles lateral to the spine.  IM injections should be reserved for non-survival surgeries as many of these injections are irritating and may cause lameness, self-mutilation, or cutaneous ulcerations.
  4. Intravenous (IV): For mice and rats, injections are most often given in the lateral tail veins.  These vessels may be dilated by placing the tail in warm (not hot) water for 1 to 2 minutes prior to injection.  For mice, a 25 or 27 gauge needle will be required.  For rats, a 25 gauge needle will be required.
• When preparing your own anesthetics, such as tribromoethanol, be cognizant of the proper methods for preparation and storage.
• Ophthalmic ointments should be administered to the eyes when anesthetics such as Ketamine or any other medication that abolishes the blink reflex are used.
• Please refer to the IACUC website page entitled Information on Anesthetics and Analgesics regarding any questions on types or doses of anesthetics available for use in rodents.  See the link to this page in the Instructions section.

Monitoring anesthetized animals:

Anesthetized animals must be monitored during the procedure to ensure that they remain in the proper anesthetic plane.  The anesthesia must not be allowed to get too light or too deep.  Once the anesthetic has had time to take effect, the following items must be addressed.

• Assess the anesthetic plane by pinching the toes, tail or ear of the animal.  Any reaction from the animal indicates that the animal is anesthetized too lightly and that additional anesthesia is required before beginning the procedure.  Other general signs of inadequate surgical anesthesia include:
• • reflex activity (corneal reflex),
  • increase in muscle tone (i.e. increased jaw tone as surgery progresses),
  • increase in respiratory or heart rate,
  • increase in blood pressure, and
  • movement and/or vocalization during the surgery.
• Care must be taken to prevent anesthetic planes that are too deep, as this may lead to death.  High doses of injectible anesthetics or high doses of inhalation anesthetic given for a prolonged period of time can result in excessively deep planes.  Signs that may indicate deep planes of anesthesia (decreased heart rate, decreased respiratory rate, decrease blood pressure) can often be difficult to see, so appropriate use of anesthetics with experience will minimize this possibility.
• Note that the most frequent complication of small animal anesthesia is hypothermia, which significantly prolongs anesthesia, impairs recovery and can lead to the death of the animal.  Provide the animal with a heat source during the pre-operative, intra-operative and post-operative periods. Doing the procedure in chemical fume hoods or biosafety cabinets increases the risk of hypothermia because of high air flow.  Acceptable forms of heat sources include:
• • re-circulating warm water blankets (ideal),
  • slide warmers (can be used during recovery),
  • instant heat devices like Safe-n-Warms or hand warmers (can be used with caution), and
  • heat lamps and electric heating pads (should be used with caution as varying temperatures can be seen across the surface.  Only 50% of the cage should be covered by a heat lamp or heating pad to allow animals to escape.  The animal must always be separated from the heat source by a towel or drape to prevent thermal injury to the animal.  Hyperthermia can be as detrimental as hypothermia.)

Recovery of the anesthetized animal:

 The recovery period begins with completion of the surgery or procedure and if inhalation anesthetics are utilized, when the animal is removed from the inhalation anesthetic source. Appropriate recovery monitoring will help to ensure timely attention to problems that may arise.  Trained personnel and appropriate facilities and equipment must be available for postsurgical care.

• Additional recovery requirements
• • Animals should be recovered in a clean cage.
  • Recovering animals should not be placed onto loose bedding material until they are fully awake, as suffocation can result.  A paper towel or sterile drape can be placed between the bedding and the animal until it awakens from anesthesia.
  • Dehydration can be ameliorated by the administration of appropriate fluid therapy.
  • • Initially, 2mls of warm fluids (0.9% NaCl, saline or lactated Ringer’s solution) per 100 grams of body weight can be given by subcutaneous injection.
    • If blood loss occurred during the surgical procedure, additional fluids may be administered.
  • Animals must be monitored continuously until they are sternally recumbent and every 15 minutes after until they are mobile.  Animals can be moved back to their home cage only after they can maintain sternal recumbency without assistance and can ambulate on their own with appropriate balance.
  • • It is important that the animals be able to ambulate on their own so that they can make it to the food or water available within their home cage, and/or find a warmer or cooler area within their home cage.
    • Ensuring mobility will also ensure that the animal is recovered enough to not aspirate bedding material which is another concern of placing an animal in their home cage too early.
    • Monitoring of the animal can be discontinued when the animal is able to ambulate on its own with appropriate balance, and moved back to its home cage.
  • Nutritional support in the form of moistened food being placed on the cage floor can be given after the animal has been moved back to its home cage.

Applicability:

This policy applies to all animals used for research and teaching purposes at UT Southwestern Medical Center.

Instructions:

Preanesthetic fasting is not necessary for mice and rats.

For anesthesia or analgesia options, please log onto the IACUC Drug Information page at

http://www.utsouthwestern.edu/utsw/cda/dept238828/files/358261.html#ranalgesia

If you have any concerns about exposure to anesthetic gases, please contact Environmental Health and Safety at 8-2250

The IACUC recommends that an anti-spill device be used to pour Isoflurane into the vaporizer. An example of an anti-spill fill device can be found on the Vet Equip Inc. website at http://www.vetequip.com/item.asp?cat=3&catalogID=911111-3

Exceptions:

Exceptions to this policy may be granted by a decision of the IACUC in an approved protocol.

Contact Information:

If you have any questions concerning inhalation or injectible anesthesia agents, please contact the ARC veterinary staff at 8-4100.