Human responses to pain vary widely, not only among individuals but for any individual depending on circumstances. (For discussions of the complex topic of pain in animals, see Bateson 1991; Elzanowski and Abs 1991; Gentle, 1992; Andrews et al. 1993.) The evident psychological component is usually aggravated by fear. Similarly, various species respond to traumatic experiences differently, and either restraint or disorientation may elicit more evident distress than such physical injuries as punctures or small incisions. Unfortunately, an animal's fear of the unknown cannot be lessened by assurances. Hence, anesthetics may be used not only to ameliorate pain but to reduce the total stress of a procedure. On the other hand, there may be circumstances in which an investigator judges that any stress from the procedure will be increased and/or the bird's chances of survival decreased by administration of anesthetic. A decision to forego anesthesia must always be justified in terms of the benefit to the animal, never in terms of convenience to the investigator. For a general reference, see Smith and Swindle (1994).

Anesthesia ideally creates minimal stress in both administration and safety of the bird, but also provides adequate restraint and/or analgesia for the intended procedure. Choice of agent is based in the animal's general body condition and on working conditions in the laboratory or field. AWA mandates consultation with a veterinarian for laboratory animals, and AAALAC supports this practice for all vertebrates. Perhaps the most important message on this subject is that there are no easy answers and no single agent that is ideal for all situations. The agent of choice for one species may be ineffective, or lethal, for other, sometimes closely related, species. Dosage ranges vary widely among taxa. Within species, effects may vary with age, sex, season, or fat content of the bird. Prolonged recovery time or the necessity of special equipment may render an anesthetic of choice in the laboratory totally inappropriate for field use. Every anesthetic agent has specific advantages and disadvantages. The investigator, in consultation with a practicing avian veterinarian, must take the time to determine which agent or combination of agents is appropriate to the study and to justify that decision.

The effects of drugs on many species are unknown, but great variation of response among species is common. When information concerning the effect of a drug on the species under consideration is unavailable, pre-experimental testing with low dosages is advised, [see X.FJ A good starting point for those working with exotic species is Samour, et al 1984, which surveys the effects of CT-1341, ketamine and ketamine plus xylazine on 154 species in 15 orders. Genevois et al. (1983) also considers several agents and 35 species. Various techniques are summarized in Fedde (1978), Sinn (1994), Chapter 23 of Muir et al. (1995), Heard (1997) and several chapters in Redig et al. (1993).

An anesthetic is an agent that may be used for one or more of 1 ) analgesia (reduction of pain), 2) immobilization, or 3) tranquilization. General anesthetics can perform alt of these functions, but not necessarily equally well, and the effects of many are dose-dependent. Hence, it is important to recognize the level of the effect.

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Some anesthetics, e.g., ketamine hydrochloride (VetalarÆ Parke-Davis), are classified as "dissociatives." They relax the subject and decrease the emotional response to pain, but have little or no true analgesic properties. Humans given such drugs report that they can feel pain, but that they don't care.

General anesthetics are administered either as a gas or as an injection. Gaseous anesthetics have the large advantage that dosage is easily changed during the procedure, and, because of the clearing properties peculiar to the avian respiratory system, recovery can be extremely rapid. Many of the best are better administered with, or even require, special equipment for precision delivery. Light-weight, portable systems are available for field use (Exotic Animal Medical Products). For a practical reference to field use of gaseous anesthetics, see Olsen et al. 1992. Injected anesthetics may be placed in a muscle mass (I.M.) or in a vein (I.V.). The latter provides more predictable reactions, faster induction and usually faster recovery, and should be used when possible, but it requires some skill even with large species and is inappropriate for small species. The dosage for most injected anesthetics varies inversely with weight (Boever and Wright 1975), i.e., small birds require relatively more. Hence, the weight of the subject must be accurately measured prior to administration of the drug, and the drug may need dilution.

Anesthetics may be combined with each other or other drugs for synergistic or antagonistic effects. Ketamine is often used as a sedative to be followed by a gaseous anesthetic using endotracheal intubation. Ketamine is the injectable anesthetic of choice for many birds, but recovery is often violent. Hence, a common practice is to mix equal volumes of ketamine and xylazine, which has strong muscle relaxing as well as weak analgesic properties. This procedure eases but also prolongs recovery. The prolongation can be countered by administration of yohimbine (Hsu 1985; Heaton and Brauth 1992). Such complex procedures require special skills. Investigators wishing to use them should consult, or practice with, an avian anesthesiologist.

Given the difficulties of administering some of the common general anesthetics, the use of local anesthetics is attractive, especially if the procedure is simple and the bird is to be released quickly. However, the advantages may be more apparent than real. Dosages are uncertain, and the effects may be general and prolonged (Graham-Jones 1965). To some extent, the problem is one of size (Gandal 1969; Klide 1973), with small species (or perhaps simply budgerigars) susceptible to overdose. Further, studies on mammals indicate that several common local anesthetics, including 1% procaine, 0.2% tetracaine. 0.5% lidocaine (with and without epinephrine), 2% chloroprocaine, 0.25% dibucaine, 2% mepivacaine, and 2% piprocaine, have temporary but severe myotoxic effects (Basson and Carlson 1980; Foster and Carlson 1980; Carlson and Rainin 1985). Hence, the intramuscular use of local anesthetics should be undertaken with caution.

Chilling has sometimes been used for topical analgesia. Ethyl chloride can temporarily numb a small area for quick incision, such as in laparotomy (Risser 1971). Refrigerants such as dichlorodifluroromethane may also be used for cryosurgery. However, because it is difficult to control the degree of local chilling, and frozen tissue may be permanently damaged or rendered inoperable, and because a relationship between hypothermically induced immobility and analgesia has not been clearly established, the use of chilling as a general anesthetic is strongly discouraged.

Avian surgery is considerably different from mammalian surgery (Ritchie et al. 1994; AItman et al. 1997). In part, the differences are due to avian structure, especially the airsacs and flow-through respiratory system, and/or physiology, e.g., blood-pH and proclivity to fall into hypothermia. These can generally be accommodated by altered techniques. However, two differences require further comment. First, birds may not show an inflammatory response to infection associated with cuts or punctures. This does not mean that birds are not subject to infection. The physiological response is simply different from that of mammals, so standards of antisepsis cannot be relaxed. Second, many birds show little behavioral evidence of pain or discomfort from punctures or incisions over much of the body, especially in the apteria (Green 1979; Steiner & Davis 1981). The head and bill, scaled por-

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tions of the legs, and vent area are exceptions. However, most birds show strong evidence of discomfort from pinching or pulling of the skin. Because of this lack of response, because anesthesia can be stressful to a bird, and because birds can be severely stressed by prolonged handling (Gandal 1969), many aviculturists and investigators perform some surgical procedures, including laparotomy and muscle biopsy, with little or no anesthesia and close incisions without sutures (Risser 1971; Wingfield & Farner 1976; Baker 1981). Such procedures need not affect the survival or reproductive potential of the subject (Ketterson and Nolan 1986; Westneat 1986; Westneat et al. 1986). Given the availability of local analgesics and the rapidity with which birds can recover from such gaseous general anesthetics as isoflurane, the practice of invasive techniques without the use of anesthesia requires special justification, e.g., the bird is to be released into the wild immediately, [see F] In no case should such procedures be performed by those who have not developed the necessary skills.

The acceptability of a procedure varies with the experience and skill of the investigator. Any invasive procedure more complicated than a simple injection should be rehearsed with an appropriate model (mock-up, cadaver, generally anesthetized subject), and the most conservative limitations on techniques should be maintained until they can be performed quickly and smoothly. As a major portion of surgical trauma for many birds is the necessary restraint, rapid, but not hasty performance can markedly reduce distress. Again, individuals who are not familiar with a technique should learn it directly from the tutelage of an expert.

The conditions governing the adoption of procedures may depend on the intended fate of the bird. We can distinguish four categories of subjects:

a. wild birds in the field that are to be released immediately upon recovery;

b. wild birds that have been brought into a laboratory and will be released after recovery in a holding facility;

c. wild or captive bred birds that are to remain captive permanently or for an indefinitely long period after the procedure;

d. birds that will be euthanized without recovery.

For any animal that is to be released to the wild, the prime consideration shall be that the procedure will have a minimal effect on the subsequent survival and reproductive potential of the subject. If the purpose of the experiment is to alter survivability or reproductive potential, then the interference should be no more than necessary, as judged and justified by the investigator, to test the issue in question.

High standards of antisepsis should be practiced routinely during invasive procedures. Many procedures need only chemical sterilization of instruments. No single procedure for sterilization is appropriate to all materials and all situations. Procedures for chemical disinfection and sterilization have been reviewed by the American Dental Association (Council on Dental Therapeutics et al. 1985). Precautions should be taken to reduce the possibility of disease transmission. Disposable blades and needles should be used, and instruments should be immersed in a strong disinfectant between subjects (household bleach or 90% ethanol plus flaming). Instruments should be rinsed in sterile, distilled water after immersion in either bleach or ethanol (unless flamed) before use. A plastic board can serve as a temporary, portable surgical area. This, too, should be sterilized with bleach or ethanol between uses.

Aseptic conditions are not required in the laboratory, but the surgical area, which should be specifically designated and set aside for that use only, at least at the time of use, should be scrubbed with a strong disinfectant, e.g., dilute sodium hypochlorite (household bleach, dilute 1/10), quarternay ammonium compound, or an iodoform compound (followed by alcohol to remove the residue), before and after procedures. All organic debris from previous procedures must be removed or sterilized. Special precautions, such as color coding and separate storage areas, must be taken to insure that surgical instruments are used for that purpose only. They must not be mixed with autopsy, dissection, or skinning instruments. The investigator should wear disposable surgical gloves during the procedure.

AAALAC regulations require that a sterile field be maintained as effectively as possible -even in the field. One can use plastic boards for surgical procedures. These can be dis-

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infected with 1/10 (or even 1/32) dilution of household bleach solution, quaternary ammonium compound, chlorine dioxide based sterilant (ClidoxÆ), or chlorhexidine (NolvasanÆ). Alternatively, the board can be wrapped with pre-sterilized cloths or disposable paper covers. Sterilizing instruments under field conditions is a vexing problem. A variety of chemical sterilizing solutions exist, but these may require that the instruments be rinsed in sterile water before use. Techniques that may be appropriate to specific situations can be garnered from any of the publications mentioned above and from studies on rodents (Cunliffe-Beamer 1993; Callahan et al. 1995). The best general solution is to use disposable instruments and blades. As long as they are not contaminated with blood or other tissues, non-disposable instruments can be soaked in 70% ethanol between uses. The investigator should wear surgical gloves, which can be obtained from medical supply stores; they are light and compress into a small storage space for field use. Whenever possible, the procedures should be carried out in some sort of shelter that reduces airflow and the possibility of wind-borne contaminants.

Repeated surgeries on a single subject are discouraged unless they are part of a single experiment and have scientific justification. Although repeated surgeries may be desirable toreduce the number of birds to be removed from the wild, especially if the investigation involves imported species, that argument is not recognized for primates, and approval must be sought from USDA.

Wound closure may present difficulties as the skin of many birds is not suitable for holding sutures. Fortunately, the small incisions used in most field procedures will heal effectively without sutures. Cyanoacrylic tissue glues (e.g., Tissu-GluÆ, Ellman International, or VetbondÆ, 3M Corp.; N.B. household super glues are toxic to tissues) may be used. A disadvantage is that the necessary drying time markedly increases handling time. However, such procedures may be useful to protect the cleanliness of the wound if the bird is to be returned to a relatively dirty environment such as a nest or open water. Surgical staples are an effective and rapid means of closing large incisions in medium-sized and large birds. However, as they must be removed mechanically, they are not advised for field use.

Other issues specific to birds:

a. Because of the high metabolic rates characteristic of birds, pre-surgical fasting is not advised for small birds and should be of a duration sufficient only to empty the crop in large birds (overnight for large birds, 4-6 hrs at most for small birds).

b. Hypothermia is a common avian response to general anesthesia. Therefore, the surgical recovery areas should be warm, and special heating arrangements may be necessary for prolonged recoveries.

c. If recovery is prolonged, the bird should be rotated to lie on alternate sides every few minutes. An anesthetized bird should not be allowed to lie on its back except as necessary for the surgery. The cage should be covered to reduce stress.

d. Some anesthetics, especially ketamine-plus-xylazine, do not induce eye closure. In such cases, the opened eyes should be bathed with an optical wetting agent every few minutes or should be protected with an opthalmic ointment.

Although considered a minor procedure by some, laparotomy penetrates a body cavity and, thus, is here considered a major surgical procedure. Exploratory laparotomy has several uses. It can provide information on sex in monomorphic species and stage of gonadal development, as well as indicate presence of parasites, gross condition, and activity of other organs. Topical application of xylocaine cream may reduce discomfort of laparotomized birds (Ritchie et al. 1994). Many experts perform this procedure with only a topical anesthetic or no anesthetic at all, especially in the field, where speed of operation is important so that the bird can be released quickly and in a condition to avoid predators. [see E] Such usage is not recommended for anyone lacking abundant practice on anesthetized or recently deceased birds. Even skilled practitioners should practice following any significant hiatus in performance. Several reports have shown that laparotomy has no effect on survival and does not disrupt breeding activity or winter foraging (Bailey 1953; Miller and Miller 1968; Wingfield and Farner 1976; Ketterson and Nolan 1986). In the laboratory, and in the field where practicable, isoflurane is ideal anesthetic for this procedure [see D].

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In many cases, such as small landbirds, sealing the laparotomy wound is not necessary, and healing occurs within 3-7 days (Wingfield and Farner 1976). In spite of the fact that small birds heal swiftly, sealing the wound might be advisable in some instances. Wounds in waterbirds should be sutured to reduce infection. In those species that dive, the wound must be sealed to avoid injection of water into the body cavity as pressure increases with depth. Surgical glues serve this purpose well. [see E] An unpublished study reveals that, after laparotomy of diving alcids and closure of the wound with non-toxic glue, adults returned to the breeding colony and nested normally. Body mass was not reduced, suggesting that the birds were able to dive and feed normally.

Recently, several less invasive techniques have become available for sexing birds. These are summarized in Halverson (1997). Especially promising are techniques using DNA to assess the presence of the W-chromosome, e.g., Quinn et al 1990, as these could be used to sex nestlings and even eggs (Langenberg et al. 1997; Nuechterlein and Buitron 1997). These appear to be highly effective and accurate. However, the easier and cheaper polymerase chain reaction techniques are species specific (Griffiths and Tiwari 1993). Preparation and testing of a probe may take several trials, even for an expert (P. Parker, pers. comm.). Hence, one should be sure that the proper probe is available before using the technique on critical specimens. Similarly, flow cytometry is relatively easy, but may not provided totally unambiguous data (Tiersch et al. 1991; McLain and Roth 1997). All of these techniques require the return of tissue samples to the laboratory and some delay during processing. Nevertheless, investigators are encouraged to explore the appropriateness of these new techniques to their studies.

The technique for euthanasia should not interfere with post-mortem analysis and should be as swift and as painless as possible. The technique adopted will be considerably influenced by what one wishes to do with the cadaver (i.e., use it for a museum specimen or for tissue chemistry or just dispose of it). Many techniques for euthanasia have been reviewed by the American Veterinary Medical Association (Andrews et al. 1993). Relatively few are appropriate for birds, and none consider the needs of field studies. Hence, we make some general comments on the nature of what might be done and list only those procedures that are not acceptable. This is another area in which a close working relationship with an avian veterinarian will be useful.

Generally acceptable techniques involve overdose with an anesthetic, either injected or gaseous (including carbon dioxide) or administration of a specific euthanasia compound (usually based on barbituates). Such procedures pose little problem for laboratory studies, but may be impractical in the field. Field investigators who normally include hypodermic syringes as part of their equipment, e.g., for tissue sampling, will probably find that a small bottle of anesthetic or euthanasia compound adds little burden. Others may find the advantages of a technique that provides a specimen with minimal damage and can be easily adjusted to any size specimen are attractive. There will remain, however, field conditions in which carrying the equipment for administering a drug is impractical, or perhaps even illegal. Such situations require mechanical means for dispatch. The traditional technique of cardiac (thoracic) compression approaches the limits of present standards of speed and minimal stress and may not be accepted by an investigator's IACUC. Thus, although the technique is permissible for field use, we recommend use of an alternative whenever practical. A mechanical alternative is cervical dislocation, in which the neck is quickly stretched (not twisted) until the spinal cord snaps. This technique is easily learned and can be used on birds as large as pheasants or small geese.

Unacceptable methods include curare, decamethonium, gallamine, magnesium or potassium salts, nicotine, pancuronium, strychnine, and succinylcholine. These chemicals are stressful or do not induce rapid analgesia. Carbon monoxide and ether are undesirable because of danger to personnel. On the other hand, if an animal has been anesthetized, or is unconscious from trauma, the mechanical means of euthanisia are of less consideration. It is important to remember that the primary purpose of euthanasia is to terminate suffering. Hence, speed is important. An animal that is already dying from severe trauma, e.g., from

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gunshot, should be terminated by the fastest available method compatible with preserving the desired portions of the specimen.

Finally, euthanasia is not a technique for the disposal of animals at the end of an experiment but a procedure to end chronic distress or pain. Investigators should seek ways to provide healthy experimental subjects with an opportunity for a continued, comfortable existence.