I. Definitions
Before we begin a detailed discussion of each of these topics, it is prudent to review the definitions listed below. A clear understanding of each of these terms is needed to comprehend why each anesthetic agent has been placed in its respective category.

• Analgesia - loss of sensitivity to painful stimulation without loss of consciousness.
• Anesthesia - loss of feeling or sensation, which is often accompanied by loss of consciousness.
• Hypnosis - a state of artificially induced sleep or trance.
• Local Anesthesia - loss of sensation to a section of the body.
• Narcosis - a sedated state in which the patient has some analgesia.
• Neuroleptanalgesta - a state of profound sedation and muscle relaxation that occurs following administration of a narcotic analgesic combined with a tranquilizer.
• Pain - a sensation that is commonly aroused by an unpleasant stimulus and leads to avoidance reactions.
• Restraint - state of being controlled, confinement.
• Surgical (General) Anesthesia - unconsciousness with adequate analgesia and muscle relaxation to al-iow surgical manipulation without struggling or pain perception by the patient.
• Sedation - mild degree of central depression in which the patient is awake but calm (used interchangeably with tranquilization).

Later I will specifically discuss assessment of analgesia and analgesic drugs. These definitions were constructed for use in man, are based on a verbal description of perception, and are quite difficult to universally apply to both man and nonverbal animal subjects. Nonetheless, until alternative definitions are available, we must do our best to establish criteria for assessing anesthesia, analgesia and a surgical plane of anesthesia in animals.

II. Types of Anesthesia

There are six major types of anesthesia, which include:

• parenteral (injectable)
• inhalation
• local
• stress-induced
• hypothermia
• hypnosis

III. Selection of Anesthetics, Analgesics and Tranquilizers

There are several criteria which should be used when selecting anesthetics, analgesics and tranquilizers for use in various species of laboratory animals.

1. Species, strain of patient
2. Age, size, sex of patient
3. Physical condition of patient
4. Temperament of patient
5. Pre-existing disease (health)
6. Temporary tolerances
7. Previous administration of other drugs
8. Management practices (prior to, during and after anesthesia)
9. Purpose of anesthesia/analgesia, i.e. what do you want the anesthetic to do?
   • restraint, examination
   • blood sampling
   • dosing
   • surgical anesthesia - what is the duration and degree of manipulation involved in the surgical procedure?
10. Type of study or procedure

    - the choice of anesthetic should provide
    the least adverse effects, thereby minimizing the influence of anesthesia on the data obtained in the study (e.g., when performing a respiratory study opioids which cause respiratory depression should be avoided).

11. Is post-operative analgesia required?

    Some anesthetics, such as Innovar-Vet®, can have prolonged analgesic effects which continue into the post-operative period.

12. Available equipment/laboratory environment
    • Is inhalation equipment available?
    • Is the study to be conducted in a unidirectional flow (UDF) room? The latter is important in many animal facilities, as UDF rooms use recirculated air, therefore we prohibit the use of volatile agents in these rooms unless a ventilated hood is available.
13. Skill and experience of the anesthetist

    Some anesthetics are quite safe to use and can thereby be used by relatively inexperienced personnel. Other agents, however, have a narrow margin of safety and should only be used when an experienced and knowledgeable anesthetist is present.

14. Availability/cost of anesthetics
    • Some anesthetics (barbiturates, narcotics) are controlled substances and require a D.E.A. license for purchase.
    • Some anesthetics, such as Safran®, are not approved for use in the U.S

IV. Stages of Anesthesia

1. Stage 1: Voluntary Excitement
   
   
   1. Excitement, struggling, salivation, lacrimation, urination and defecation by the animal may occur during induction of anesthesia.
   2. Lasts from initial administration until loss of consciousness.
   3. Can be caused by forced inhalation of strange, irritant gas under unusual circumstances.
   4. Epinephrine released which produces increased heart rate, rapid and deep respiration and struggling against restraint.
   5. Minimized through the use of preanesthetics.
   6. Variable degrees of analgesia.
      
      
2. Stage II Involuntary Excitement
   1. Characterized by loss of consciousness.
   2. Exaggerated reflex, struggling or purposeless muscular movement by the animal occurs during slow induction of anesthesia that is characteristic of inhalant anesthetics.
   3. Respiration is uneven in depth and rate, and breath holding may occur. The eyelids are wide open and the pupils are dilated.
   4. The administration of rapidacting and potent agents, such as thiamylal, thiopental or other similar agents, avoids these undesirable characteristics.
      
      
3. Stages I and II together constitute the period referred to as induction.
4. Stage III: Surgical Anesthesia
   1. The depressant action of the anestheticis extended from the cortex and midbrain to the spinal cord.
   2. Consciousness, pain sensation and spinal reflexes are abolished.
   3. Muscular relaxation occurs and coordinated movement disappears.
   4. The hypothalmic centers are paralyzed progressively during this stage resulting in inactivation of the heatregulating mechanisms so that loss of body heat fails to evoke such protective responses as shivering and vasoconstriction. The animal's body temperature falls markedly unless protected against heat loss.
   5. Major reflexes and methods of assessment during surgical anesthesia:
      • Palpebral (lid) Movement of the eyelids when the inner canthus is touched.
      • Corneal Closure of the eyelids upon lightly touching the cornea.
      • Skin
      • Swallowing The act of swallowing.
      • Cough
      • Pedal Withdrawal of the limb on painful stimulation.
      • Thermoregulatory function of animals is suppressed as early as the initial phases of Stage III, with shivering only present in Stage ii. The presence of shivering is quite often indicative of very light anesthesia, when recovery from anesthesia is in progress.
      • Skeletal muscle tone, which is nearly normal at the beginning of light surgical anesthesia, decreases due to a depression of the ordinary posrural reflexes.
   6. Deep Surgical Anesthesia
      • In deep surgical anesthesia, reflexes such as the palpebral, corneal and pedal are completely depressed. These reflexes, especially the pedal, may persist throughout all planes following barbiturate anesthesia. Skeletal muscle tone rapidly disappears, leaving the animal flaccid. The pulse is rapid and weak.
      • in the latter part of this stage, as the level of anesthesia becomes deeper, the disappearance of intercostal respiration indicates a dangerous depth of anesthetic depression.
      • Most major surgical procedures can be performed under fight to moderate surgi cal anesthesia.
5. Stage IV: Medullary Paralysis
   1. This stage is characterized by paralysis of the vital regulatory center of the medulla, and death soon ensues.
   2. The heart usually beats weakly for a short time after respiration ceases.
   3. There is a complete absence of all reflexes and complete dilation of the pupils.
   4. Anesthetic(s) must be withdrawn and resuscitative actions initiated to save the patient.

V. Unique Features of Rodent Anesthesia and Analgesia

Rodent anesthesia offers unique challenges that are encountered infrequently in any other veterinary setting. Due to the common practice of providing anesthesia for multiple patients simultaneously, a 'herd' anesthesia approach is necessary.

Additional features that require consideration in planning an anesthetic protocol include:

• dosing in mg/kg volume quantities from premixed dilutions or combinations of drugs.
• poorly accessible peripheral vessels for IV Induction
• "one drug bias" by certain scientific specialties
• difficulty in extrapolating dosages from humans and other animals to rodents
• multiple species and strains which show differing sensitivity, efficiency and duration with various anesthetics
• comparative lack of scientific studies focusing upon anesthetic efficacy in hamsters, gerbils and guinea pigs
• infrequent monitoring of physiologic parameters in rodents
• the use of anesthetic agents in rodents which have no corollary in veterinary or human medicine, thus physiologic, toxicologic and pathologic effects have not been determined
• greater difficulty in judging anesthetic and analgesic depth and muscle relaxation is present in rodent species
• Federal, regulatory agency and activist review of rodent welfare during anesthesia and analgesia is less focused, hindering development of improved guidelines. Anesthetic drugs may be chosen based on expense, available equipment and s 'kill of the anesthetist
• requirements for aseptic surgical procedures influence ability to administer and monitor anesthesia

VI. PreOperative Patient Evaluation and Care

A. Rodents

Rodents require appropriate pre-operative evaluation and care when planning experimental manipulations, including those requiring anesthesia. The colony history should be carefully reviewed to determine the original source of the animals and the original health status. The current age of the animals and the length of tithe they have been present in the facility should be determined, and the most current information available from serological monitoring of sentinel animals or other available health surveillance reviewed. Newly arrived animals should be quarantined and allowed to rehydrate and acclimate after shipping for at least three days (Landi, 1982). Body weights for the group should be consulted when selecting dosages for a particular strain, sex or age of rodent, and the time of day selected for anesthesia. Older or otherwise compromised animals will benefit from biochemical screening. Preanesthetic fasting should be considered/contemplated/ restricted in light of the high metabolic rate and rapid dehydration and blood glucose detriments that occur rapidly in animals of small body mass. Feed restriction should be limited to two hours prior to surgery. Animals scheduled for early morning procedures could be given a limited but adequate feeding in the prior late afternoon (15gm.) Water should not be restricted. Animals scheduled for anesthesia should be individually weighed and their weights recorded on the cage card or ID tag prior to induction.

B. Rabbits

Prior to experimental manipulations, animals should be quarantined to allow acclimation to their surroundings as well as to permit health surveillance by the veterinary staff. The preanesthetic evaluation of rabbits includes a physical examination with chest auscultation and rectal temperature measurement. Because blood samples are easily obtained from conscious rabbits, it is also prudent to obtain baselines hematologic and biochemical parameters prior to inducing anesthesia. Other preanesthetic screening measures may be employed to satisfy the scientific needs of the particular study. In addition, animals may require training to specialized equipment and techniques prior to initiation of an experimental study.

VII. Methods of Anesthetic Delivery/Equipment

A. Rodents

There are basically two methods of anesthetic delivery to rodents, parenteral and inhalation, or a combination of both. 'Balanced anesthesia' is a relatively novel method for producing anesthesia in humans that is becoming popular for us to use in animals. This format utilizes combinations of drugs, each in a sufficient amount to produce its major or desired effect(s) to the optimum degree, and yet keep its undesirable or unnecessary effects to a minimum.

Parenteral Anesthesia
Parenteral anesthesia refers to intraperitoneal, intramuscular and intravenous routes of administration.

Techniques for Administering Injectable Agents
The three critical factors to be considered when proposing parenteral drug delivery to rodents are: 1) the drug volume to be delivered, 2) the site(s) for drug delivery, 3) the irritant properties, if any, of the anesthetic compound, and 4) the method of administration. Parenterai anesthetics can be delivered by single bolus injection, intermittent injection as needed and by continuous infusion; the latter method requiring some form of syringe infusion pump. Because the volume of drug to be injected is often quite small in rodents, it is preferable to dilute many agents to a 50% or less solution. This technique will not only assure that a significant percentage of the chosen agent will not remain within the 'dead space' of the needle and syringe, but will also often decrease the irritation attendant with many acidic drug solutions, such as ketamine and Innovar-VetO. To further decrease the irritation caused by injection of irritating solutions, required dosages may be divided and smaller volumes injected at multiple sites or, alternatively, diluted with an equivalent volume of physiological saline or sterile water prior to injection.

a. Technique of intraperitoneal (IP) drug injection

The IP route of drug injection is probably the most popular method of drug delivery to rodents because 1) minimal skills are required and those needed are easily mastered, 2) easily accessible peripheral blood vessels for IV injection are nonexistent in several species of rodent (e.g. hamsters, guinea pigs), 3) many anesthetics that create irritation when given by the intramuscular or subcutaneous routes do not cause lesions or clinical evidence of pain (e.g. limping) when given IP and 4) many published regimens for anesthesia of rodents have been formulated for IP drug delivery.

To properly administer an intraperitoneal injection to a rodent, the animal should be restrained so that it is held in a headdown position with the needle inserted into the lower left abdominal quadrant. Errors in IP drug injection can be minimized by both fasting the animal for 4-8 hours prior to drug injection and by use of a 20-22g needle, rather than a smaller (e.g. 25 or 26g) needle which may not penetrate the subcutaneous tissue, fat and abdominal wall and thus never deliver the anesthetic to the large peritoneal surface for absorption. Scientists should also be aware of potential peritonitis caused by IP drug administration, particularly when irritating solutions are administered or bacterial contamansion is introduced.

b. Technique of intramuscular (IM) drug injection

Intramuscular drug injections are usually delivered into either the caudal thigh muscle mass or the epaxial muscles along the spine.

c. Technique of intravenous (IV) drug injection

In mice, rats and gerbils, IV anesthetic or analgesic drug injections are most often made into the lateral tail vein, particularly after this vessel has been vasodilated using a heating lamp or a heated restrainer (e.g. the type of heated restrainers used to perform noninvasive blood pressure monitoring in rats). With smaller rodents (e.g. mice), a 27g needle is required for IV administration, while a 22g needle is often adequate for IV access in a larger rat. Other vessels, such as the dorsal metatarsal vein and, in particular, sublingual vein, require marked restraint of the animal such that they are rarely used for induction of anesthesia. The latter vessels would, however, be useful for post-operative analgesic drug delivery at the conclusion of surgery, before the animal has regained consciousness. When IV access is desired for long-term drug infusion, a thin, flexible catheter can be inserted into the tail vein through a small bore needle, and connected to a suitable perfusion system.

Intravenious anesthetic delivery to a guinea pig or hamster is rarely performed. Recently, a technique has been described for catheterization of the lateral saphenous vein in the anesthetized guinea pig (Nau, 1993). After placing a torniquet above the hock, a 22 gauge over-the-needle catheter is inserted 2-3 mm distal to the protruding vessel. Using this technique, the cannula remains patent for IV injections for up to 5 hours, although blood collection may not always be possible. Additionally, small volumes of drugs can be delivered into the dorsal metatarsal vein or into a catheterized femoral or jugular vein in these species.

d. Techniques to minimize irratancy/tissue response to anesthetics

Various degrees of tissue damage almost inevitably result from IM drug injections of several irritating anesthetics, e.g. ketamine and Innovar-Vet® (Gaermer, 1987, Smiler, 1990). In the case of small laboratory animals this irritation may or may not be clinically evident. In some cases, however, sufficient tissue damage can occur which leads to lameness, selfmutilation of a limb or cutaneous ulceration, thus undeniably causing some magnitude of pain and/or distress to the animal. Techniques which may be useful to minimize tissue irritation from IM drug delivery include deep rather than superficial IM delivery of the drug, use of small gauge (e.g. 22-25 gauge) needles, delivery of multiple small injections into several IM sites rather than injection of a large volume at one site, and dilution of the drug prior to injection with physiological saline.

e. Technique for continuous intravenous infusion

When delivering anesthetics by continuous intravenous infusion, a concept known as the 'minimum infusion rate (M.I.R.)' should be utilized. The M.I.R. is the minimal infusion of the injectable agent that keeps the patient asleep. Anesthetics are delivered by this method following in dwelling catheterization as previously described. Drug administration requires a syringe infusion pump or similar equipment calibrated to continually deliver the relatively small drug volumes required for use in rodents.

Inhalation Anesthesia

Inhalation anesthesia involves the delivery of volatile anesthetic agents to the patient via the respiratory tract. In the past, delivery of volatile anesthetic agents to rodents was not considered to be feasible due to their small size and the unavailability of appropriate equipment. From a practical standpoint, anesthesia of rodents often involves induction of multiple animals simultaneously or in sequential fashion to permit surgical manipulations on a group of animals. This format is often more amenable to injectable anesthetics, which can be rapidly administered, rather than investing the greater time required for either chamber, mask or endotracheal tube induction using inhalant agents. Greater technical competence is also needed to deliver inhalant agents, particularly when endotracheal intubation of these small subjects is required. Unless skilled personnel are available, problems encountered with intubation of small rodents may include trauma to oropharyngeal structures by insertion of the laryngoscope, local irritation or laceration of the trachea by catheters used as endotracheal tubes, and possible subsequent induction of subcutaneous emphysema. Furthermore, in contrast to the minimal cost incurred in parenteral anesthetic delivery (i.e. drug + syringe + needle), the use of volatile agents is accompanied by the attendant costs of vaporizers, oxygen, endotracheal tubes, laryngoscopes, face masks, anesthesia chambers and delivery circuits. Finally, rodent anesthetic and surgical procedures have frequently been performed outside of dedicated surgical areas (e.g. laboratories), which may not have scavenging equipment available for operator safety and comfort.

Despite these disadvantages, investigators and veterinarians are increasingly learning that inhalation anesthesia can be reliably and safely delivered to rodents with moderate temporal efficiency and cost effectiveness. Major advantages of inhalant techniques are: 1) increased operator control over depth and duration of anesthesia, thus greater survivability, and 2) ability to choose agents (e.g. isotlurane) that require minimal metabolism, biotransformation or excretion, and thus contribute minimal variability to overall goals of the research project. Because equipment purchased and/or fabricated to deliver volatile agents to rodents can be reused indefinitely, costs incurred are rapidly recovered when contrasted with replacement animal costs for the higher mortality often attendant with parenteral agents as well as reimbursement for personnel required to support animals during the often prolonged recovery period which follows the use of parenteral agents (e.g. Pentobarbital). Obviously, greater survivability coupled with a decreased probability that anesthesia contributes untoward variability to experimental results is both humanely and scientifically beneficial.

1. Methods of Delivery of Inhalant Agents to
Rodents

• Open drop - ("bell jar") does not permit control over anesthetic; high exposure for personnel.
• Nonrebreathing (open) system - exhaled anesthetic mixture is released into the at-mosphere. Methods of delivery include endotracheal intubation, use of an anes-thesia chamber, or delivery using a face (head) mask.
• Rebreathing (closed) system - exhaled mix-ture flows through soda lime which removes all the carbon dioxide. Uses leas anesthetic and less oxygen. Due to the small tidal volume of rodents, this method is rarely practical or useful in these species.

Ancillary Equipment

Additional equipment that is useful for administering inhalant anesthetics to rodents includes an anesthesia chamber, oxygen tank, vaporizer, varioussized masks, endotracheal tubes (can be fabricated from over-the-needle catheters), fiberoptic light source, mouth speculum or customized laryngoscope and miniaturized breathing circuits.

A. Delivery of fithalant anesthetic gases using various types of anesthesia chambers and face masks


The easiest method for delivery of volatile anesthetic agents to rodents is by use of an anesthesia chamber. 'Chambers' can be fabricated simply and inexpensively by use of a large covered glass container ("bell jar'). The liquid anesthetic is volatilized by placing moistened cotton balls or gauze squares into file bottom of the jar. Prior to placing the rodent in this crude chamber, it is advisable to cover the anesthetic-impregnated cotton with a woven mesh grid to prevent local irritation of file anitaars feet by direct contact with the liquid anesthetic. The animal is then placed within the jar and visually observed for cessation of movement and recumbency, thereby signifying the onset of anesthesia. Because this system of induction involves no calibrated vaporizer, the anesthetic concentration within tile jar cannot be controlled and lethal concentrations can rapidly accumulate. For this reason, the 'bell-jar' or open drop method is usually reserved for use with methoxyfiurane which reaches a maximum concentration of approximately 3% after full volatilization, in contrast to levels of approximately 30% inhalant gas which can be reached upon volatilization of halothane or isofiurane with this method. Because this method contains no provisions for scavenging excess gases, it should be used within a fume hood or otherwise ventilated area to minimize inhalation of methoxyflurane (with its attendant nephrotoxic potential) by surrounding personnel. An additional disadvantage of the 'bell-jar' technique is that no apparatus is present to maintain anesthesia once the animal is removed from the jar, thus this method only provides anesthesia for very shortterm procedures, e.g. orbital bleeding, subcutaneous tumor implantation, tattooing.

A more sophisticated method for delivering volatile agents to rodents involves use of an anesthesia chamber alone or in combination with a facemask appropriately sized for rodents. Standardly, anesthesia chambers are constructed with a fresh gas inlet as well as outlet for exhaled and waste gases. Ideally, the rodent is placed within the chamber for induction, then removed from the chamber with anesfilesia maintained by delivery through a face or head mask. Both chamber and mask delivery incorporate the addition of a calibrated vaporizer into the circuit for precise control of the concentration of anesthetic gas delivered to the patient. Because oxygen flow is required to volatilize the liquid anesthetic placed within the vaporizer, oxygen is also delivered to the patient and helps to maintain the blood oxygen saturation. Fairly high fresh gas flows are required for either chamber or mask delivery, therefore adequate suction of waste anesthetic gases is necessary to avoid exposure to personnel. Anesthesia chambers are commercially available or can be fabricated from plexiglass or other clear materials or by modification of a fish tank or terrarium. Face or head masks can be easily made from a funnel or proximal end of a 20, 50 or 60 cc syringe barrel, depending on the size of the rodent. Several references for construction of rodent anesthesia chambers or circuits are included at the conclusion of the chapter (Dardai, 1987, Franz, 1988, Glen, 1980a, Levy, 1980, Mulder, 1984, Norris, 1981 & 1982, Querldo, 1985, and Tarin, 1972).

B. Methods of Anesthetic Delivery to Rabbits

Anesthetics can be delivered to rabbits by either parenteral or inhalant methods, or a combination of both. Balanced anesthesia is a technique for producing anesthesia in humans that is now becoming popular for use in animals. This format utilizes combinations of drugs in doses selected to maximize their major or desired effect(s) and minimize undesirable or unnecessary effects (1). Balanced anesthetic regimens often include one or more parenteral agents plus a muscle relaxant and inhalant. Parenteral anesthetics are most often delivered to rabbits by the subcutaneous (SC), intramuscular (IM) and intravenous (IV) routes. Although intraperitoneal (IP) anesthetic injection is certainly feasible in rabbits, it provides a slower onset and greater variability than IM and IV administration. Because some anesthetics may be irritating when delivered subcutaneously, care should betaken to dilute the injectate. Although subcutaneous administration may be the safest method of delivery, the onset of anesthesia will be slower and the anesthetic plane achieved will show greater variability as compared to other methods. The very accessible marginal auricular (ear) vessels simplify intravenous anesthetic delivery to rabbits. For IM injection, anesthetics can be delivered into either the semitendinous or semi membranous muscles of the rear limbs or into the epaxial spinal musculature. Iatrogenic nerve irritation or injury from IM injection may lead to acute pain and rear leg paresis. In the author's experience, this is a particular problem with anesthetic injection into the spinal epaxial muscles.

Recently, anesthetic regimens are being designed for administration by intermittent or continuous intravenous administration to rabbits. This method of delivery is somewhat more complex that simple bolus delivery in that intravenous access is required and, in the case of truly continuous infusion delivery, some type of infusion or syringe pump is required. The physiological advantages of this method suggest that it is one of the most progressive anesthetic methods available, particularly for prolonged procedures. The primary advantage of continuous IV infusion delivery is the ability to supply and withdraw each component of

anesthesia in a consistent and continuous manner, such that wide variations in drug absorption and their physiologic manifestations do not occur. Using this method, many of the severe adverse effects of anesthesia, such as hypotension, apnea, cardiac arrhythmias and prolonged recovery, are minimized. Rabbits are ideal candidates for this method of delivery due to their easily accessible marginal auricular veins.

Inhalant anesthesia requires use of an anesthetic machine and various ancillary equipment. A non-rebreathing pediatric circuit (e.g. Bain, McGill, or Mapleson) should be used to accommodate the relatively small tidal volume of the rabbit. Inhalant anesthetics can be delivered by anesthetic chamber, mask or endotracheal tube. Specialized plexiglass chambers are commercially available or can be fabricated to be appropriately sized for rabbits. Depending on the breed and size of the rabbit, cat, infant or child-sized masks can be used for induction. Because inhalant anesthetics can be irritating and thus objectionable to the rabbit, animals should be properly restrained during mask induction to avoid kicking and injury to the animal as well as attendant personnel.

VIII. Intubation Of Rabbits And Rodents

A. Intubation of rats, guinea pigs and hamsters

As with larger species, endotracheal intubation is most easily accomplished in rodents by direct visualization of the pharyngeal and laryngeal structures. Customized rodent laryngoscopes are not commercially available; however, there are reports in the literature describing the equipment needed and techniques for fabricating laryngoscopes suitable for use in rats, hamsters and guinea pigs. Although there appear to be no publications describing endotracheal intubation of the gerbil, it seems likely that techniques and equipment suggested for use in similarly-sized rodents would be applicable.

Intravenous over-the-needle catheters (sizes 14-20, depending on the size of the rodent), are usually suitable for intubation of rats, hamsters and guinea pigs [(size 14, (60 mm) for guinea pigs, 16 (55 mm) for hamsters and sizes 14-20 for various sizes of rats)]. When using over-the-needle intravenous catheters as endotracheal tubes, it is advisable to file down the pointed stylet prior to insertion in order to decrease the likelihood of oropharyngeal and/or tracheal trauma.

Guinea pigs, in particular, are a challenge to intubate due to their large tongue which hides the larynx and their small epiglottis which is angled upward and positioned anteriorly. Intuhation of many species of rodents can be accomplished by direct visualization using a Wisconsin pediatric laryngoscope size 0 blade attached to a pediatric laryngoscope handle and lightsource. When using this equipment in guinea pigs, it is advisable to bilaterally file the anterior aspect of the blade to facilitate its insertion into the narrow mouth of this species. This modification is not needed for laryngoscope insertion into the short and wide mouth of hamsters.

An alternative to intubation of rodents using a purpose-made laryngoscope is to position the animal on its hack and insert an oral speculum (e.g. an operating otoscope fitted with a small, short coned tip). Alternatively, one can transilluminate the pharynx by illuminating the ventral neck with a powerful light source. Commercially-available fiberoptic penlights can be used for this purpose. When placed on the ventral neck, the light source penetrates the tissues underlying the larynx and illuminates the oropharynx, thereby permitting direct visualization of the larynx.

When using an intravenous catheter as an endotracheal tube, some modification of the Luer fitting is needed to provide connection to the appropriate anesthesia circuit. Connectors and tubing components of the anesthesia circuit should be miniaturized such that the smallest possible volume of dead space is present within the anesthesia circuit.

B. lntubatlon of Rabbits

Endotracheal intuhation of rabbits presents a formidable challenge to veterinarians and technicians. Anatomically, rabbits have a narrow oropharyngeal opening with minimal excursion of the jaws, a small posteriorly situated larynx and a narrow, small tongue that is difficult to grasp to allow visualization of the larynx. Due to this anatomy, many veterinarians settle for delivery of inhalant anesthetic gases by mask rather than to risk penetration of oropharyngeal structures with subsequent air leakage and subcutaneous emphysema. Alternatively, tracheostomies are frequently performed on rabbits ancillary to administration of inhalant agents; however, complications (such as stricture) can result from this surgical procedure. Clearly, it is most advantageous to the animal for the veterinarian and/or technical staff to possess the skills to accomplish endotracheal intubation in rabbits expediently and reliably, with minimal attendant trauma.

Equipment is available to assist the veterinarian in successfully intubating rabbits. This includes a 3.0 mm uncuffed endotracheal tube, a pediatric intuhation stylet, a pediatric laryngoscope (size 0-1 Wisconsin blade, depending on the size of the rabbit) and 1% lidocaine for application by cotton swab or aerosol onto the laryngeal opening.

Prior to intubation, rabbits should be sedated with parenterai anesthetic(s) or by mask delivery of volatile anesthetic agents plus oxygen. Positioning of the animal varies with the preference of the person performing intubation; however, my personal preference is for the animal to be in lateral recumbency with the head of the animal in extension and the tongue pulled forward by gripping it with a dry gauze pad. A pediatric laryngoscope containing a size 0 to # 1 Wisconsin blade on a penlight handle is then used to visualize the larynx and glottis. Because rabbits will easily exhibit laryngospasm upon attempts to introduce the endotracheal tube, I standardly either swab the laryngeal opening with a cotton swab moistened with 1% lidocaine or, alternatively, spray 1/4 ml of 1% lidocaine (without epinephrine) onto the vocal folds. A homemade atomizer can be fabricated for this purpose by attaching a 20 g over-the-needle catheter (stylet removed) to a 3 ml syringe, the latter containing 1/4 ml lidocaine and 3 mls air (the air is needed to nebulize the lidocaine). The endotracheal tube with stylet in place is now inserted into the oropharynx.

Unfortunately, introduction of the endotracheal tube largely obscures the view of the vocal cords at the moment of intubation. I find it quite helpful to watch the animal for 30 seconds or so to become familiar with its respiratory pattern and rate so that I can then time advancement of the endotracheal tube into the laryngeal opening during inspiration. Despite !idocaine application to the vocal cords, the rabbit will still often cough following insertion of the endotracheal tube into trachea, which signifies proper tube placement. Additionally, the endotracheal tube will opacify with exhaled air (particularly when a narrow stylet or no stylet is used) if the tube is properly placed. A modification of this technique is to use a small gauge (#5 or #6 French) intravenous catheter as a guidewire. Using this technique, the catheter is passed through the endotracheal tube such that the catheter extends beyond the lip of the tube. The catheter-tube is then held in the operator's dominant hand, and laryngoscopy performed. When a stable view of the larynx is achieved, the tip of the catheter is advanced into the trachea during inspiration. The endotracheai tube is then advanced over the catheter (the latter acting as a guidewire) into the trachea. The introducing catheter is then withdrawn and the endotracheal tube is secured. Advantages of this method are that intubation is accomplished with visual evidence of translaryngeal placement of the guide wire in contrast to the 'blind intubation," particularly with small rabbits, that is unavoidable when the endotracheal tube itself is introduced.

An alternative method for blindly intubating rabbits is to insert the tube without the stylet into the oropharynx and position the tube in the supra epiglotic region (i.e. just proximal to the laryngeal opening) by watching the tube cloud up upon expiration. By 'getting the pace' of inspiration/expiration, the operator can then quickly advance the tube during inspiration into the open trachea. Although I find the latter method more serendipitous than scientifically derived, it does seem to reliably work for some technicians and veterinarians.

As with cats and ferrets, an improperly placed endotracheal tube can be palpated externally by noting two rigid, cylindrical neck structures, e.g. the trachea and the intubated esophagus. Because rabbits have a broad, flat nose, the properly positioned endotracheal tube should be secured with umbilical tape or gauze tied to the tube and behind the ears.

IX. Monitoring Anesthesia In Rabbits And Rodents

At first glance, it seems impossible to apply the sophisticated monitoring techniques which are commonplace in human medicine and with larger animals to rabbits and rodents; however, equipment and techniques are available for monitoring anesthesia in these small species. Despite the limitations of monitoring and supportive care that are applicable to these species, anesthetic overdose, metabolic and respiratory acidosis, blood loss and hypothermia are often contributing factors to cardiovascular collapse and subsequent death of these animals. Thus, it is advisable to perform some degree of monitoring of anesthetized rabbits and rodents so that therapeutic intervention can occur or anesthesia can be terminated prior to a fatal outcome.

Parameters which can be used to monitor the physiological status of anesthetized rodents and rabbits include:

• monitoring of anesthetic depth
• respiratory rate and pattern
• arterial blood gases, pH and oxygen saturation
• pulse
• arterial blood pressure
• ECG
• core and surface body temperature
• analysis of inhaled and exhaled gas for anesthetic agent and oxygen delivery, as well as end tidal CO2 content.

Parameters which can be used to assess the depth of anesthesia in rodents and rabbits include:

• recumbency and loss of purposeful movemerits (caution - 'swimming' or purposeless limb movements often are retained in hamsters and gerbils, in particular, even at a deep plane of anesthesia!)
• loss of reflexes (e.g. corneal, pedal, pinnae reflexes). Note: In terms of anesthetic-monitoring in rabbits, the best reflexive response to use for evaluation of a surgical plane of anesthesia is the 'ear pinch reaction' or loss of pinnae reflex. Loss of the corneal reflex is a sign of a dangerously deep plane of anesthesia in rabbits.
• muscle relaxation and lack of vocalization
• response to averslye stimulation (e.g. pinching the abdominal skin with a hemostat, pinching the toes or the tail)
• alterations in respiration, cardiovascular function and EEG's.

In clinical practice, cardiovascular and respiratory assessments are limited to observations of chest wall movement to determinig respiratory rate and palpation of the apical pulse through the chest wall. However, more sophisticated methods are available for monitoring of anesthetized rodents and rabbits. These methods include both invasive (i.e. using implanted arterial catheters connected in turn to transducers and recorders or using telemetry) and noninvasive (using tail or leg band cuffs connected to pneumatic pulse transducers, sphygmomanometers and recorders) blood pressure measurement. With either invasive or noninvasive blood pressure measurement, data obtained is usually limited to the mean rather than both systolic and diastolic pressures. Commercially-available portable monitors can also be used to noninvasively record the EKG, heart rate and respiratory rate of rabbits, and will even accurately monitor the heart rate in anesthetized rodents. Arterial catheters can also be used to monitor electrocardiographic changes and to provide arterial blood samples for blood gas measurements. Recently, pulse oximetry is being adapted to rabbits as well as rodents. It is possible to modify commercially-available neonatal human digital and foot sensors as well as adult nasal probes for pulse oximetry on animals. The color of the eyes under reflected light can also be used as a crude measure of perfusion and oxygenation. To avoid aberrations in respiratory parameters, ventilation can be controlled using a commercially-available ventilator that is customized for rodents or rabbits.

Because the ratio of body surface area to body mass is greater in rodents than in larger species, thermal support may be critical to the successful recovery of rodents, in particular, from anesthesia. Particularly with rats and mice body heat may be dissipated from the tail, soles of the feet and ears to the environment, with a resultant profound decline in the core and surface body temperature. This hypothermia may, in turn, lead to a decline in both anesthetic metabolism and urinary excretion of the anesthetic agent. It is customary for us to consider hypothermia as inevitable in almost all anesthetized rodents and rabbits. We make special efforts to provide thermal support to minimize this adverse effect.

X. Supportive Care of Anesthetized Rodents and Rabbits

Similar to the techniques and equipment available to monitor anesthetized rabbits and rodents, it is not currently practical or feasible to provide the same intensity of supportive care, particularly with regard to anesthetized rodents, as is commonplace with that applied to humans or larger research animals. Techniques that are available, however, for supportive care of rabbits and rodents are briefly discussed below.

Methods to minimize heat loss to the environment during anesthesia include techniques such as increasing the ambient temperature of the operating room, placement of a thermal blanket (e.g. recirculating warm water blanket) or drape between the animal and the stainless steel operating table, use of heat lamps (carefully placed!) during surgery, minimization of organ exposure from body cavities during surgery, use of warmed irrigation solutions throughout surgery, recovery of the animal upon a warming blanket or within a temperature-supported cage or infant warmer, administration of warmed subcutaneous, intraperitoneal, or intravenous fluids intra and/or postoperatively, recovery housing on bedding to provide thermal insulation (rodents), and recovery with cagemates (when permitted by the experimental protocol) to permit animals to huddle together and thus provide thermoregulation. A microwave is a must for your OR prep area such that intraoperative and postop fluids can be warmed! Commercially available avian ICU cages are ideal for group housing of rodents. These cages can be supplied with oxygen, increased humidity and an increased ambient temperature. They are relatively small in size, yet can be used for recovery of multiple anesthetized rodents simultaneously. Commercially-available canine ICU cages are ideal for recovery of anesthetized rabbits. Again, temperature, humidity and oxygen support are available. Portals are also present for exteriorizing IV fluid administration sets. Intravenous fluid warmers are also commercially available. These units maintain IV fluids at 37'C and thus minimize hypothermia due to the administration of cold IV fluids.

Rabbits and rodents have high energy requirements due to their small size and high metabolic rate, yet they have minimal fat reservoirs which can be mobilized to supply needed energy. Nutritional support is critical upon recovery to avoid hypoglycemia, particularly if the animal was fasted prior to induction. Nutritional support can be provided by simply providing a high-quality pelleted diet as soon as the animal has recovered sufficiently to ambulate and eat (remember - rodents and rabbits don't vomit!). Additionally, calories can be supplied to anesthetized animals by injecting small volumes (10-25 mis, depending on the size of the animal) of warmed 5% dextrose solution subcutaneously. Rabbits can also be provided with IV calories present in 5% dextrose or commercially-available parenteral diets.

Volume deficits can be corrected by subcutaneous or intraperitoneal injection of warmed saline, Lactated Ringers solution or replacement fluids (e.g. Normosol®)· If significant blood loss has occurred, blood transfusions can be administered, using the tail or jugular veins (rodents) or auricular veins (rabbits). Particularly with inbred rodent strains and transfusions into naive animals, blood typing is not usually needed and transfusion reactions infrequently occur.

Because rodents and rabbits are frequently anesthetized with injectable agents that inhibit blinking (e.g. ketamine), ocular lubrication is important to protect against corneal ulceration. I have found the C57/B! mice, in particular, are sensitive to corneal ulceration after ketamine anesthesia if lubrication is not applied.

Table 1. Anesthetics and Tranquilizers for Use in Mice

Table 2. Anesthetics and Tranquilizers for Use in Rats

Table 3. Anesthetics and Tranquilizers for Use in Guinea Pigs

Table 4. Anesthetics and Tranquilizers for Use in Hamsters

Table 5. Anesthetics and Tranquilizers for Use in Gerbils

Table 6. Analgesics for Use in Rodents

Table 7. Analgesics for Use in Rabbits

Table 8. Sedatives, Tranquilizers, Anesthetics and Analgesics for Use in Rabbits

References:

• Rabbits