Introduction

The appropriate use of anesthetics, analgesics and tranquilizers in research animals is a critical concern to all. Under all circumstances, the use of these drugs is reviewed during the IACUC protocol approval process. However, since these issues are so important to humane research and they involve a highly complex variety of topics, this document has been produced in an effort to provide additional guidance and consultation to research personnel on these issues.

Proper use of pain alleviating drugs involves sophisticated control of all key physiological systems (nervous, cardiovascular, and respiratory systems). General anesthesia is filely a controlled near-death state for the anesthetized animal. A plethora of factors influence the process each time it is performed. Health of the animal can have a major impact. An animal that appears quite normal on initial inspection may have a serious pneumonia that can lead to uncontrolled anesthesia and death. A procedure area that is cold can dramatically change the manner that an animal will react to administered drugs. An individual animal's response to an anesthetic drug will vary during a procedure in response to the research manipulations as well as it's changing physiological state. It is known that age, sex, strain, previous drug exposure, and even time of day of exposure can have important impacts on anesthetic drug responses. These effects can be dramatic; different strains of mice given 40mg/kg pentobarbital IP display sleep times varying from 10 to 300 minutes! The knowledge of physiology and pharmacology required for precisely managed anesthesia is enormous. This required expertise is recognized in clinical medicine by highly trained and specifically certified physicians and veterinarians. Conversely, in the research environment, persons administering anesthetics may have only cursory training in these areas. Fortunately through the use of modern drugs with wide safety margins and the impressive resilience of healthy animals, anesthesia in the research setting is generally quite routine. However, always remember that anesthesia is a complex system and many factors can abruptly crop up to cause serious problems.

The following consists of a short compilation of information on anesthesia, analgesia and tranquilization of typical research animals. If the information does not meet your research needs, please contact the Campus Veterinarian for further assistance.

Definitions Commonly Used in Animal Anesthesia

• Analgesia: Loss of sensation of pain.
• Anesthesia: Total loss of sensation in a part or in the whole body, generally induced by the administration of a drug that depresses nervous tissue activity.
• Local Anesthesia: Loss of sensation limited to a local area.
• General Anesthesia: Loss of consciousness and loss of sensation throughout the body.
• Surgical Anesthesia: Loss of consciousness and sensation with sufficient muscle relaxation and analgesia to allow surgery to be performed without pain or struggling.
• Dissociative Anesthesia: A central nervous system state characterized by muscular rigidity, peripheral analgesia and altered consciousness (e.g. ketamine anesthesia).
• Balanced Anesthesia: Surgical anesthesia produced by a combination of two or more drugs or anesthetic techniques each contributing differing pharmacological effects.
• Terminal Surgery: The animal is anesthetized, a procedure is performed and euthanasia is conducted before the animal regains consciousness.
• Survival Surgery: Procedure performed with the intent of recovering the animal to consciousness from the anesthesia.
• Major Surgery: Procedures involving exposure of a body cavity or which have a potential for creating a permanent impairment.
• Minor Surgery: Surgical procedures that do not meet the definition of Major.
• Controlled Substances: Drugs regulated by the Drug Enforcement Administration. They are classified according to abuse potential.
• Neuroleptanalgesia: Hypnosis and analgesia produced by a combination of a neuroleptic drug and an analgesic drug (e.g.Innovar Vet).
• Sedation: A mild degree of central nervous system depression in which the patient is awake but calm. The patient may be aroused with sufficient stimuli. Sedatives act by dose dependent depression of the cerebral cortex.
• Tranquilization: A state of tranquillity and calmness in which the patient is relaxed, awake and unconcerned with its surroundings. With sufficient stimuli the patient is aroused. Analgesia is not a component of tranquilization. Tranquilizers act by depressing the hypothalamus and the reticular activating system.

Considerations In Choosing An Anesthesia Regimen

When choosing anesthetic agents consideration must be given to providing appropriate depth and length of anesthesia and analgesia as determined by the procedure. The agents used should be safe and effective for the species under study given its age, sex, history and physical condition. The agent should also be safe and relatively convenient for the personnel that are administering and monitoring anesthesia.

When choosing an anesthesia regimen careful consideration must be given to the type of procedure being performed, the duration of the procedure, the amount and type of pain that may accompany the procedure, and the goals of the project. Each agent has a different duration of action, length of recovery time, amount and site of analgesia, degree of muscle relaxation, site and route of detoxification or excretion, etc. The type of procedure also greatly influences the anesthesia requirements. Some procedures cause more pain than others. An understanding of the agents being used is also necessary to assure that they are safe for the species under study and that they will not interfere with the goals of the project. Drugs may be used in terminal procedures that are not appropriate for survival procedures.

The animal under study will also influence the anesthesia agents and route of administration. The size and anatomy of the animal subject will affect the route and consequently the type of anesthesia administered. Tracheal intubation with inhalation anesthesia is the route of choice for most procedures. Tracheal intubation can be performed on most species. However, it is difficult and requires considerable expertise for small rodents and rabbits.

It seems obvious that the health and general condition of the animal will affect the outcome of anesthesia but the investigator must be aware that animals may hide or compensate for disease processes to such an extent as to appear normal.

Understanding the anesthetic agent or agents that are to be used is extremely important in assuring success. Often an investigator's experience with a particular agent and the agent's previous use on a research project are the factors that determine its choice. Experience with an agent or anesthesia regime is important but it should not preclude consideration of other factors and consideration of new anesthesia regimes.

Pre-anesthesia Evaluation

Prior to induction of anesthesia, animals should be carefully evaluated for good health. If there are any doubts about the animal's health, anesthesia should not be initiated until a thorough veterinary exam has been conducted.

The pre-anesthesia exam should be noted on an anesthesia record. The exam should contain but is not limited to: confirmation of animal's number or other identification; sex; age; body weight; body condition; skin condition; estimate of hydration; color of mucous membranes; heart rate and rhythm; respiratory rate and any indicators of respiratory disease; signs of diarrhea; body temperature.

The body condition of the animal is very important to anesthesia induction, maintenance and recovery. Very thin animals may react rapidly to some agents and become overdosed at what should be an appropriate dosage. Very fat animals may be slow to respond to drugs. The skin and fur of an animal is a good indicator of the animal's general state of health. With practice, the skin can be used to roughly evaluate an animal's hydration status. If "pinched up" skin does not flatten out when released, the animal is dehydrated. Dry, tacky mucous membranes of the mouth are also an indication of dehydration. The color of the mucous membranes around the teeth should be pink in most animals.

Heart and respiratory rates and character can provide valuable information about an animal's health. They should be routinely checked so that abnormalities will be recognized. Gastrointestinal problems frequently cause changes to the stools. Stools should be noted as an assessment of the animal's health. Body temperature can be very useful as well; however, it is difficult to obtain from most laboratory animals under routine circumstances.

Withholding food from an animal (fasting) prior to induction of anesthesia is recommended with some species to reduce the incidence of vomiting and the potential for aspiration during anesthesia. Pigs, cats, primates, and ferrets should have food removed at least 12 hours prior to induction of anesthesia. Water should be removed one hour prior to induction. It is not recommended to remove water sooner than this because it may lead to dehydration. Rodents and rabbits typically are not fasted prior to anesthesia. If anesthesia is required for a pregnant animal, the need to withhold food should be carefully evaluated.

Preanesthetic Medications

It is often advisable and sometimes necessary to premedicate an animal before induction of anesthesia. In general anticholinergic drugs, tranquilizers, and narcotics are used for premedication. Anticholinergic drugs are used to decrease oral and respiratory secretions; maintain the heart rate; and decrease gut motility. Tranquilizers relieve anxiety and produce calmness, aid in restraint, calm postoperative recovery, and reduce dose of anesthetic needed. Narcotics are used as preoperative medications to: quiet an animal; produce vomiting and defecation; reduce the amount of anesthesia needed; smooth recovery; and to provide postoperative pain relief.

Administering Anesthesia

Injectable Anesthesia

Intraperitoneal (IP) injections are the most common site of anesthesia injections in rodents. This has become a preferred site for rodents because of difficulty finding small veins and muscles. Although this technique has been used in larger species such as rabbits, ferrets and cats, it leads to unpredictable dose responses and is not consistent with good veterinary practices.

IP injections in rodents are relatively simple. Learning videos covering handling techniques are available at Kerr Learning Center and the laboratory animal facility staff are available for hands-on training and assistance. In general, IP injections in rodents require firm restraint of the animal with control of the head. Injections should be made slightly off the midline in the caudal (towards the tail) quadrant of the abdomen. Use the smallest needle that will allow easy injection of the anesthesia product. For small rodents, dilution of the anesthesia agent is recommended for accuracy of dosing. Problems may arise if the injection goes into the subcutaneous space, the urinary bladder, gut or a major organ such as the liver. If the handler is reasonably familiar with the animal's anatomy and is adept at restraint, these generally are not problems.

Certain agents can be given by the intramuscular (IM) and subcutaneous (SQ) routes. Generally, these routes result in a slower induction time. IM and SQ are often used for sedation and tranquilizing prior to induction of surgical anesthesia. Intramuscular (IM) injection is relatively simple in "large" animals (e.g. greater than one kg). In rodents, IM injections should be done carefully and the volume of injection must be small. IM injections can be administered in any large muscle mass; the usual site is the back of the hind leg. Generally the site of injection need not be disinfected; but if the animal is dirty, the area must be cleaned. The largest gauge (smallest diameter) needle that allows easy injection of the solution but that resists bending should be used. Subcutaneous (SQ) injections are simple in most animals. SQ injection deposits the injected substance in the space below the skin. This site is appropriate for administration of fluids when IV administration is not available. Irritating or hypertonic solutions should not be given SQ. As with IM injections, the needle size will depend upon the animal, viscosity of the solution and the volume needed.

Inhalation Anesthesia

Inhalation anesthesia has advantages and disadvantages when compared to injectable anesthesia regimes. The disadvantages include: necessity of an adequate scavenger system to prevent human exposure; expense of a calibrated vaporizer for larger animals; expense of anesthesia agents in open systems; need for trained and experienced personnel to run anesthesia equipment. The major advantages of inhalation anesthesia are: rapid induction; rapid changes in level of anesthesia; safety for the animal (with experienced operator); good analgesic effects and fewer adverse physiologic affects (generally speaking). In general, for long procedures and major operative procedures, inhalation anesthetic agents would be preferred.

Anesthesia may be induced with an injectable agent such as ketamine or pentobarbital and then maintained with inhalation agents. Rodents may be induced using inhalation agents by placing the anesthetic on cotton or gauze placed in the bottom of a covered container. A glass desiccator jar is ideal. They have a perforated platform and a lid that seals and they provide good visualization of the interior. A screen or perforated platform must be used to cover the moistened cotton. Animals must not come in contact with the liquid anesthesia agent because anesthetic agents are irritating to the skin and mucous membranes. The animals must not be left unattended as overdose or suffocation may occur. Once the animal is unconscious and respiration has slowed, it is removed from the induction chamber and anesthesia is maintained.

The best method of maintaining anesthesia using inhalation agents is with a gas anesthesia machine. Such machines have calibrated delivery vaporizers that deliver precise quantities of the anesthesia agent in oxygen. Anesthesia may also be maintained by delivering additional anesthesia via an "open drop" system. Anesthesia may be administered via a tight fitting mask or nose cone or preferably through an endotracheal tube. If intubated with an endotracheal tube, the animal's ventilation can be supported and personnel exposure can be more readily limited. The level of anesthesia must be closely monitored and the cone moved away from the animal or vaporizer concentration decreased as depth of anesthesia increases. Nose cones used in an open drop system require holes in the end so that room air is pulled across the anesthetic agent. Halothane and isoflurane both have high vapor pressures at room temperature and animals can easily be overdosed with these agents when precision vaporizers are not used. Methoxyflurane has a maximum vapor pressure of 3% which is generally a safe dosage.

All inhalation agents affect humans and may cause drowsiness and headaches. It is particularly important to avoid repeated exposures over a long period of time. Specific scavaging systems that draw waste anesthesia out of the building or that capture the agents are necessary. Cabinets and hoods that draw gases away from personnel can be effective in protecting personnel from exposure.

Monitoring Anesthesia

Monitoring to assure that the level of anesthesia is adequate for the procedure being conducted is a major component of proper care. Monitoring the anesthetized animal involves four body systems: the central and peripheral nervous system; the respiratory system; the cardiovascular system; and the musculoskeletal system. Normal values of heart rate and respiratory rate should be obtained prior to anesthesia. Basic patient parameters that should be evaluated throughout the operative procedure include: heart rate (pulse rate), rhythm and pulse intensity; respiratory rate, depth and character; mucous membrane color, capillary refill time; body temperature; muscle tone; and reflexes. The standard measure of the cardiovascular system is the heart rate. Heart rate will generally decrease as the animal goes deeper under anesthesia. Monitoring the animal's body temperature throughout the operative procedure is extremely important. Anesthesia causes reduced muscular activity and depressed central thermoregulatory control leading to lowered body temperature. Peripheral perfusion is also decreased. Poor circulation and hypothermia potentiate anesthesia and increase recovery times. Care must be taken to maintain an animal's body temperature in a normal range. Thermal water circulating pads should be used whenever possible. Anesthetized animals should always be placed on towels or pads, never directly on stainless steel tables. Animals should never be placed directly on a standard human heating pad. These pads can produce thermal burns even at low settings. As with the heart rate, respiratory rate generally decreases (fewer breaths per minute) with depth of anesthesia. The breaths will initially become deep and regular in interval; they then become shallow and irregular. If the animal is getting "light", respiration rate may increase and the breaths may appear shallow. In addition, a very light animal that is perceiving pain may consciously hold its breath. Since different conditions can lead to similar presentations, it is imperative to track the heart and respiratory rates throughout an operative procedure so that the trends can be used to evaluate each situation. Monitoring depression of the central nervous system in an anesthetized patient takes experience. Muscle tone is an indicator of depth of anesthesia for agents other than ketamine (by itself). A variety of reflexes can be evaluated to monitor the depth of anesthesia. Reflexes of the eye are most commonly used. The eyelids will close (blink) when the cornea or eyelids are touched. These are the corneal and palpebral reflexes, respectively. Generally, a brisk palpebral reflex is "too light" and a slow corneal is "too deep". Withdrawal of the feet when stimulated shows the pedal or withdrawal reflex. The stimulus applied must be relatively intense to elicit this reflex in an anesthetized animal. Pinching the base of the toenail or the web of skin between the toes is the method used. If a toe pinch results in an increased heart rate, increased respiratory rate, movement of the animal's body or vocalization, the animal actually perceives the stimulation as pain and clearly, the anesthesia is insufficient.

Anesthesia Recovery

The anesthesia recovery period commences at the end of the surgery and ends when the animal is fully awakened. Surgery guidelines available from the IACUC office and the laboratory animal facility discuss anesthesia recovery and postoperative care. These guidelines are consistent with the AWA, PHS Policy, routine veterinary practices and several other UC campuses' animal care policies. In general, an unconscious animal should not be left unattended. When the animal reaches a semi-conscious state it should be monitored at least every 15 minutes.

A major component of post operative care is monitoring and alleviating post operative pain. As part of the protocol review process, the expected extent of postoperative pain, how that pain will be assessed and drugs that will be used to alleviate pain must be considered by both the investigator and the IACUC. Assessment of pain in certain laboratory animals can be difficult. In general pain is indicated by: "guarding" (protecting) the surgical site; vocalization (whining, growling, squeaking) when the animal moves or if the surgical area is touched; limping, a "tucked up" abdomen, or other abnormal stances or posture; reclusive behavior; lack of appetite; increased inspiratory rate or abnormal breathing patterns.

Records

To assure that all aspects of anesthesia and surgery are conducted in an appropriate manner, written records must be maintained. The USDA regularly asks for anesthesia and postoperative care records. Even if the investigator keeps a logbook of animal procedures, complete records of preoperative evaluation and care, surgical monitoring, anesthesia recovery and post operative care must be maintained in the animal facility for species covered by the Animal Welfare Act. Records for rats and mice need not be as extensive and the animal's cage card may be used. The surgery guidelines indicated above contain information on required records and forms are available for use.
Suggested Anesthesia Regimes and Analgesia Dosages
As indicated in the rest of this document, there is a tremendous variety of anesthetic regimes for laboratory animals. A limited list of starting possibilities are below. Analgesia use should always be considered when painful procedures, such as surgery, are performed. Unless it is known that analgesia use will affect the research results, it may be better to use the drug in cases when it is "not needed" and thereby give the animals the benefit of the doubt about whether our pain assessment is accurate or not. Some consideration should be made on whether or not to use DEA controlled drugs [C_]. Properly secured storage, strictly limited access and accurate inventory and use records are required for their use.

rats: (1) inhalation anesthetics, to effect (2) ketamine 90mg/kg IM with xylazine 10mg/kg IM (3) pentobarbital [C2] 40mg/kg IP (4) flunixin meglumine 2.5mg/kg SC 2 times/day (5) buprenorphine [C5] 0.01-0.03mg/kg SC 2 times/day (6) butorphanol 2mg/kg SC 6 times/day

mice: (1) inhalation anesthetics, to effect (2) ketamine 100mg/kg IM with acepromazine 2.5mg/kg IM (3) pentobarbital [C2] 40mg/kg IP (4) flunixin meglumine 2.5mg/kg SC 2 times/day (5) buprenorphine [C5] 0.05mg/kg SC 2 times/day (6) butorphanol 1mg/kg SC 6 times/day

hamsters: (1) inhalation anesthetics, to effect (2) ketamine 100mg/kg IP with xylazine 10mg/kg IP (3) pentobarbital [C2] 60mg/kg IP (4) buprenorphine [C5] 0.01-0.03mg/kg SC 2 times/day

rabbits: (1) ketamine 35mg/kg IM preceded by xylazine 5mg/kg SC and acepromazine 1mg/kg SC (2) pentobarbital [C2] 35mg/kg IV (3) ketamine 35- 40mg/kg IM with xylazine 5mg/kg IM (4) acetaminophen 20mg/kg PO 2 times/day (5) flunixin meglumine 1.1mg/kg SC 2 times/day (6) buprenorphine [C5] 0.01- 0.05mg/kg SC 2-3 times/day

guinea pigs: (1) inhalation anesthetics, to effect (2) ketamine 40mg/kg IP with xylazine 5mg/kg IP (3) pentobarbital [C2] 40mg/kg IP (4) buprenorphine [C5] 0.05mg/kg SC 2-3 times/day (5) morphine [C2] 2.5mg/kg SC or IM 6 times/day

Approved Euthanasia Methods

Pentobarbital overdose is the most preferable euthanasia method in all cases. As a general rule, a dose that is at least 3 times an anesthetic dose will be effective. Ketamine is not acceptable for euthanasia when used alone but can be humane when used in conjunction with sedatives and tranquilizers. However, it is not very efficient as it requires very high doses. Carbon dioxide overdosage is commonly used. Physical methods of euthanasia have a high potential for being inhumane and are only acceptable when scientifically necessary and must be performed by carefully trained personnel. Physical methods are acceptable for fully anesthetized animals. In fact, physical assurance of euthanasia is critically important in all cases. Very deeply anesthetized animals may appear dead; yet, they may recover from the anesthesia at a later time. A convenient method of assuring euthanasia is to create a bilateral pneumothorax by permitting air to enter the chest cavity. This is best accomplished by making a small incision through each side of the chest. Physical assurance of euthanasia of mice and other similarly small rodents can be accomplished by cervical dislocation.

Acceptable euthanasia methods are periodically reviewed by the American Veterinary Medical Association "Panel on Euthanasia".

Selected References:
HB Waynforth, PA Flecknell, eds., (1992). Experimental and Surgical Technique in the Rat. Academic Press, San Diego
PA Flecknell (1987). Laboratory Animal Anaesthesia: An introduction for research workers and technicians. Academic Press, San Diego.

ILAR (1992). Recognition and Alleviation of Pain and Distress in Laboratory Animals. National Academy Press, Washington.

NH Booth, LE McDonald. Veterinary Pharmacology and Therapeutics. Iowa State Press, Ames.

RE Borchard, CD Barnes, LG Eltherington (1990). Drug Dosages in Laboratory Animals: A handbook. Telford Press, Caldwell, NJ.

LS Goodman, A Gilman. The Pharmacological Basis of Therapeutics. Macmillian, New York.