CARE AND USE OF ANOLIS CAROLINENSIS
IN THE LABORATORY
adapted from Appendix I to Greenberg, N. 1992. “The Saurian Psyches Revisited: Lizards in the Laboratory,” in The Care and Use of Amphibians, Reptiles, and Fish in Research, D.O. Schaeffer, K.M. Kleinow, L. Krulish, editors. Scientist’s Center for Animal Research, Bethesda, MD. Suggestions for surgical procedure provided by Dorcas Schaeffer, DVM, IACUC consulting veterinarian, are italicized; last change: August 28, 1996.
Department of Ecology and Evolutionary Biology
University of Tennessee
Knoxville TN 37996
Reptiles, while intrinsically fascinating, are also “exemplars of alternative tactics for solving ecologically important problems of survival and efficient energetics. But beyond these traditional zoological concerns, recent research … indicates that several unique qualities of reptiles may provide models useful for research on a diverse array of problems of biomedical interest”
These general guidelines represent open documentation for the attainment and maintenance of “physical and psychological well being” in captive green anoles, Anolis carolinensis maintained at the University of Tennessee, Knoxville for research purposes. The term “open” identifies these notes as regularly reviewed and perpetually amenable to modification and extension. The relevant indices of “well being” are physical appearance (including external indications of physiological stress) and changes in the known behavioral repertoire, particularly those indicative of species-typical coping strategies and diminished reproductive activity). Comments and suggestions for additional inclusions are welcome
1. SOURCE and MAINTENANCE OF LIZARDS
1a. Geographical source. To maintain consistency in the research on this wide- ranging species (Conant 1975), lizards are sought only from the New Orleans area. Lizards can be collected readily in the field where they are seasonally and locally abundant. The most reliable commercial collector is The Snake Farm, La Place, LA; This facility was visited in 1980 and in 1988 and his capture, handling, and shipping procedures appeared satisfactory and reliable. very few lizards shipped frpom this source show indications of stress or trauma; mortality is approximately 1%.
1b. Climatic and habitat variables. Temperatures between 18C and 32C are within this species adaptive scope and are thus easily tolerated if at least an hour is allowed for acclimatization. Critical elements of laboratory habitats for naturalistic behavior are adjusted periodically based on data obtained on climate and the structural habitat used by this species at the Tulane University Field Station, with the cooperation of Terry Christianson, Department of Psychology, Tulane University.
2. RECEIPT of LIZARDS
2a. Mode of Receipt. Lizards are generally received by US Air Express Mail in packages specially designed for their safe transit. Transit time is generally less than 24 hours. Typically they are received at ambient temperature wrapped with moist paper or cloth; they are quiet and healthy but become active rapidly when exposed to light or additional warmth. Lizards are to be kept cool until they are transferred to large (20-60 gallon) all glass aquaria that have been prepared to receive them.
2b. Alert all people that may handle mail that temperature and humidity are the crucial survival variables for lizards in transit and in the first hours of receipt and unpacking.
2c Examine immediately examine lizards for indications of trauma or mite infestation (see 5b, below). If mites are observed, they are removed by swabbing the lizard with 70% alcohol in a manner that penetrates sites such as flaps of skin where mites might be sequestered but avoids eyes and mouth. After careful cleansing in this way, no reinfestations have been noted.
2d. Sort and distribute lizards to holding aquaria or specific experimental habitats as quickly as possible. If subjects need to be retained in holding tanks for convenience, they are to be maintained at reduced temperatures and/or photoperiod, depending on whether or not the proposed experiments involve behavioral patterns that are controlled by hormones facilitated by specific light or temperature regimens (see below). Do not reduce ambient temperature below 25C for longer than a few days to avoid compromising food assimilation.
3. CAGES, SUBSTRATE, and PERCHES
3a. Vivaria. Vivaria are adapted from all glass aquaria; these are easy to clean and sterilize, although silicone cement and plastic retaining frames may react with certain cleansers or disinfectants.
3b. Vivarium size and design. The sizes of vivaria are dictated by the nature of specific experiments and no generalization can be made about them because of the profoundly different Umwelt (sensory world) of reptiles (Delong, Greenberg, and Keaney 1986); size must be ascertained empirically using indications of stress (below) as a guide. Cages employed successfully in past studies range from 1 gallon glass jars through 60 gallon modified glass aquaria, and even very large glass fronted museum display cabinets.
If reproductive function is a consideration, animals are to be placed in relatively small habitats under conditions known to initiate and maintain gonadal recrudescence throughout the year (Crews and Garrick, 1980; Crews, Rosenblatt, and Lehrman 1973; Licht 1967, 1971). If not housed individually, male dominance aggression will compete with reproductive activities and can retard gonadal development in losing males and nearby females (Crews, Rosenblatt, and Lehrman 1973). Careful attention to microclimate is important (see Section 4, below)
The development of naturalistic, ethologically valid habitats was described by Greenberg (1978b) and involves, briefly, successive reductions in habitat complexity beginning from a starting point approximating conditions in the field, until the least complex environment in which the animal manifests the behavior of interest is attained. Such behavior is invariably precluded by even mild physiological stress which, in this species, is immediately manifest in altered body color (see below). Ordinarily, subjects are isolated in 3-5 gallon glass vivaria, and maintained in pairs or triplets in 10 gallon vivaria. Group living is permitted in larger cages but never exceeding a population density of 1 subject per “gallon” of space with perches provided to allow subjects to maintain at least one body length (approximately 60 mm, snout-to-vent) between them. Large cages often result in more social strife than small cages, apparently because “subordinate” behavior is reinforced in small but large tanks; the effect can, however, be ameliorated by visual barriers.
3c. Substrate. Sphagnum (long stems, provided sterilized by any good nursery) is the substrate of choice. Sphagnum moss is relatively expensive but absorbs water enthusiastically, is slightly antiseptic, and easy to sterilize (autoclave) and reuse; further, it gives partial cover to live prey that compel lizards to be alert and attentive in order to feed. Sphagnum substrate is to be sterilized before being reused or whenever the individuals maintained in the habitat are changed.
Avoid substrate that provides dust or small particles that might be accidentally ingested with prey or irritate eyes (puffed clay (“kitty litter”), vermiculite, crushed mica, and playbox sand). Sterilized soil, pea gravel, or coarse sand are little better. These and other common animal substrates such as wood or corncob shavings may cause mouth irritations which can lead to stomatitis, digestive tract impactions, and cloacal irritations.
3d. Perches. Lizards can perch on glass walls, but seek more naturalistic sites when they are available, a habit which facilitates their distribution in the habitat. Stone surfaces, such as rough rocks, brick or cinderblock are effective, but crevices are sought when available. Natural bark, twigs, and houseplants also provide effective perch-sites that are attractive and naturalistic. Perch selection is, however, a significant dependent variable in many experiments (e.g., Greenberg and Crews 1990), requiring that subjects be provided with standardized water-resistant sticks, placed diagonally between the substrate and the wall of the cage in a manner to facilitate observations. Generally we use oak; aromatic woods while resistant to prolonged intensive use and frequent cleaning may compromise experiments that involve chemosensory variables. Normally, the inside glass surfaces of cages are spayed with non- toxic (“food-grade”) silicon to force lizards to use only those perches provided by the caregiver or specified by the experiment. Plastic plants may be used if you are confident they will not emit toxic fumes in humid cages; “silk” plants may be stiffened with water-soluble toxic substances that are ingested as lizards lick water from them.
3e. Cleaning. Lizards produce relatively dry feces, much like birds. There is evidence that many species utilize chemical stimuli incorporated into secreted or eliminated substances for social signals. A. carolinensis is constantly testing its environment with tongue-licks (Greenberg 1985). Ordinarily, a moderate accumulation of fecal material may ameliorate cage-novelty-induced stress. It is certainly less stressful than the intrusions of too-frequent cleaning. For cleaning, perches should be rapidly replaced with prepared replacements. Cleaning agents that leave a residue must be avoided.
3f. Frequency of cleaning. Holding cages containing more than 2 individuals per gallon of living space should be cleaned weekly. Those in experimental cages should not be subjected to the stress of having their habitat cleaned too often. Once every three weeks is generally satisfactory, but cages should be inspected at every feeding or at least weekly for evidence of potentially hazardous deviations from the initial condition. Fecal material is typically dry, but high humidity, often due to spilled water or water wicked from a water dish by a stray strand of sphagnum moss, creates an environment favorable to mold or fermentation. Any detectable ammonia, methane, or substrate mold signals the need for an immediate cleaning.
Our principal research interest is in patterns of behavior associated with seasonal activity such as aggression and reproduction. Thus, seasonal climatic variables must be monitored and controlled while subjects are being prepared for or involved in specific experiments. Temperature and light may be adjusted for individuals beyond the bounds of normal maintenance or experimental procedure for specific individuals under intensive observation, such as in post-operative care. Although A. carolinensis gonads manifest spontaneous cycles of regression and recrudescence, they are ordinarily cued by climatic variables (Fox and Dessauer 1957, Licht 1973). Thus the management of light, heat, and moisture will facilitate or disrupt reproductive cycles and behavior.
4a. Light. Photoperiod can be from 6 hr through 18 hr. A. carolinensis is a seasonally active species, in which changes in or absolute amounts of light or levels of temperature, and their interaction, are the key environmental stimuli. Paul Licht (1973) has established that a minimum 13.5 hr photoperiod is needed to promote gonadal recrudescence after their seasonal or photically induced regression. A recommended stimulatory regimen is 14h light:10h dark. The lights contribute heat which if necessary may be supplemented to provide a corresponding daily temperature fluctuations of 32o:22oC.
Although A. carolinensis are arboreal and live in a world of visual barriers (foliage), they are often exposed to full sunlight. At least some infra-red light appears to be important for manifesting the complete range of spontaneous color change; an ultra-violet component (such as that in “Vita- Lites” (Duro-Test Corp) is believed important for the full expression of photo- modulated physiological and behavioral traits and may be important to the health of hatchlings.
4b. Temperature. Thermoregulatory sources are ambient air temperature or photothermal supplementation. The thermal needs of individuals are related to diet, health, experience, daily or seasonal rhythms and, of course, the habitat in which the species evolved (Greenberg 1978b). Anolis carolinensis are poikilothermic ectotherms and tolerate range of temperature from 5 C through 35 C, and light levels from approximately indoor fluorescent lighting through outdoor sunlight, midday, midsummer. Rooms have temperature/humidity chart recorders and/or temperature alarms (set at 33 C). Temperature is known to be a significant variable in the responses of lizard adrenal to ACTH (Licht and Bradshaw 1969), on spermatogenesis (Licht 1971), and in the production and action of androgens (Pearson, Tsui, and Licht 1976).
4c. Humidity. Humidity does not appear to be a variable in the behavior of males beyond its association with thermoregulation. Humidity of 60-85% is recommended for maintenance of females in reproductive condition (Summers and Norman 1988). In a closed humid cage humidity will vary inversely with temperature ranging from 70 (day) to 90 (night) percent RH. For effective egg maintenance during incubation (detailed in Greenberg and Hake, 1990), humidity must be 90-99%.
With sphagnum as a substrate, humidity cycles regularly with temperature. In our small rooms with stable air temperature, lighting alone can raise cage temperatures ten degrees and humidity varies inversely with the temperature change from approximately 60-90%. The cycle is obvious as dew on the glass evaporates in the first hours of photoperiod. Although standing water is provided ad lib, dew is water source of choice. The temperature-humidity cycling requires closed cages with reduced airflow, a condition easily maintained with glass lids with one or two 3-5 cm diameter holes covered by screen.
In 15 years of maintenance of thousands of experimental lizards, this technique has not been associated with any distress or pathology. On the contrary, comparisons with screen-topped cages which allow low humidity conditions to prevail and/or excessively rapid drying, indicates subjects maintained in this way are more healthy and vigorous, presumably attributable to the regularity of the microclimate. The main difficulty of glass-topped cages is excessive humidity attributable to water inadvertently wicked or spilled from a water dish (see 3f).
5. PHYSICAL and PSYCHOLOGICAL HEALTH
5a. Disorders/disease. A. carolinensis are remarkably robust and manifest few spontaneous disorders or diseases. Basic and adapted husbandry procedures of Frye (1973) suffice for maintenance of health. Most common disorders are infraorbital abscesses (treated by incision, drainage, or removal of caseated debris) and chalk gland tumors. The responsible animal care facility veterinarian should be consulted in all cases.
5b. Parasites. Ectoparasites such as Ophionyssus (Sec 2b) are easily removed when lizards are first received and reinfestations have not been observed. Lizards are frequent hosts to the larvae of trombiculid mites. These are frequently sequestered in “mite pockets” in the skin of the neck, axillae, groin, or the base of the tail. These pockets protect as well as provide convenient blood supply for the parasites. While the chiggers may be a source of irritation or lesions leading to secondary infections, the pockets limit their possible damage by coping better with minor trauma and healing more quickly. (see Arnold, 1986)
5c. Stress. This species readily indicates several parameters of physiological stress by means of hormone-mediated body color (Greenberg, 1977, 1990; Cooper and Greenberg, 1992; Hadley and Goldman, 1969; Kleinholz 1938). Briefly; melanotrophic hormones are released in acute and chronic stress. Light green, brown, or mixed body color, repeatedly observed (minimum of 4 observations daily for three days) reflects a tonic level of melanotrophic hormones that are invariably associated with maintenance or increase in weight and “normal” levels of spontaneous activity. Dark green or brown or “blotchy” lizards, often with a postorbital darkening (“eyespot”) are indicative of severe adrenal activation. (In many taxa “stress” responses can be a constructive and benign variable in animal health (“eustress” as opposed to “distress”). It is manifest in both the establishment and maintenance of social dominance and other relationships important in reproduction (Greenberg and Wingfield 1987).)
5d. Psychological “well being.” A significant indication of psychological well being in this species is light body color (see above and Cooper and Greenberg, 1991), spontaneous activity (exploratory behavior, Greenberg 1985), and the latency to elicited activity. Given a relatively complex environment (see 3b, above), A. carolinensis will manifest a repertoire of approximately 40-50 distinctive units of behavior in social contexts (delineated in Greenberg 1977), which are very sensitive to social status (see Bels 1984)
5e. Reproduction. Female green anoles can manifest a complete repertoire of parental behavioral patterns if unstressed. Eggs may be maintained exposed in humid cages on sterile substrate, but the maintenance of a naturalistic light and temperature regimen is apparently important to successful hatching (Greenberg and Hake 1990). Still, only occasionally are young fledged; although neonates feed and behave normally at first, most do not survive a month; the cause of death has not been determined but muscular spasms observed in their last hours indicate a profound metabolic problem. Some workers improve the odds by calcium supplementation (dusting prey) and ultra-violet light supplementation.
5f. Deaths. A. carolinensis are relatively short-lived (4 seasons is unusually long) and in a large colony of field-caught individuals some will always be in an irreversible decline. Apparently healthy individuals occasionally die suddenly; in a dry vivarium they will rapidly desiccate/mummify; in more moist habitats they will decay, emit an offensive smell, and possibly host gnat maggots.
6. FOOD AND WATER
6a. Food. Live, moving food is required. For adults, size should be no larger than 2 cm; the exclusive provision of prey with chitinous or other hard-to- digest components (such as mealworms, Tenebrio molitor) must be avoided. Commercially supplied 3/8 inch crickets are ideal; larger ones irritate lizards. Too many crickets also irritate the lizards, and excess should be removed. Hatchlings do well on pinhead crickets or a vestigial-winged strain of Drosophila.
6b. Frequency. Two-four 3/8 inch crickets or their equivalent should be provided two-three times a week. When naturalistic levels of arousal are sought in subjects, sphagnum moss substrate through which crickets can easily move, hide, and evade lizards, should be provided and 5-10 crickets supplied once a week.
6c. Alternative food. When sweep-net contents are provided, lizards appear to be good self-selectors, but large insects that might injure a lizard should be removed. Mealworms are satisfactory, but only occasionally; they do not contain all the necessary amino acids and the chitinous exoskeleton of larger worms eventually create a serious digestion problem. Locally available mealworm colonies may provide food when crickets are unavailable due to late shipments or bad weather.
6d. Fasting. For some procedures, lizards must be fasted 2-5 days. When food- deprived for longer than 4 days, or in the event of morbid anorectic decline for other reasons, animals manifest (1) clearly defined pelvic girdle bones and (2) a bilateral depression of flesh or fat pads near the base of the tail.
6e. Cricket handling. Crickets may be received live each week and must be transferred to a “cricket bin” with a ventilated lid within two days of receipt, although they survive for up to five days in their shipping carton (at room temperature). An ideal bin is a smooth-sided plastic container at least 1 meter high, provided with cardboard tubes cut to rest on the bottom for the healthy distribution of crickets within the bin. The tubes also facilitate cricket capture for feeding: shake a hundred or so crickets from their tubes into a funnel into an Erlenmeyer flask for easy redistribution into individual cages. Never let a flask of cricket sit untended. The cricket bin must be cleaned at least every third shipment; after washing and drying, the sides are to be sprayed with food-grade silicon. Crickets survive well for at least two weeks without food or water. Further, provision of food or water would encourage cricket growth beyond the size best for their service as lizard food and also encourage invasion by pests, particularly hump-back gnats. They must be kept remote from insecticide.
6f. Water. Lizards prefer the naturally occurring “dew” that forms as temperatures cycle daily with the photoperiod. Water can, however, be sprayed to provide a naturalistic source and is strongly advised for newly received animals, although lizards will readily come to recognize and use standing water which should be provided in shallow dishes.
Provision should be made to allow crickets that might get in the water a way to escape. Dishes with rocks, perches, or sloping sides are generally satisfactory. Water should be generally present but an occasional day of “drying out” is valuable for cages that have become excessively humid. Lizards tolerate brief deprivation. Ordinarily, glass-topped cages show diurnally varying amounts of condensation on the inner surfaces, but if the substrate becomes uniformly and continually wet for more than two days the tank should be dried out.
Sustained humidity predisposes cages to infestation by fruit fly-like “hump-back gnats” (tentative identification). While these have not been seen to trouble healthy lizards, maggots may infest weak or wounded subjects.
7a. Handling must be minimized to eliminate this potential stressor which may manifest itself in subtle ways including supersensitivity to observer effects, altered fat storage and gonadotrophin release (Meier, et al. 1973).
7b. Tail autotomy. A. carolinensis are small (3-6 gram) but robust subjects. They should nevertheless be handled carefully and are best restrained by a firm fingertip grip on a portion of one of their limbs near the torso. A lizard may autotomize its tail if it is gripped and the subject’s body is free to tug or thrash against the restraint. While this is never fatal and tails regenerate eventually, it is stressful and is likely to compromise assessments of stress responsiveness in specific experiments; there is some indication that tailless lizards have altered social behavior.
7c. Caregiver’s precautions. After handling, washed hands thoroughly; A handling-stressed lizard may void its cloaca and there is the possibility that Salmonella may be transmitted by infected lizards.
7d Marking. A commercial permanent waterproof marking pen (Sanford’s “Sharpee”) is a proven effective marker to allow discrimination of similar individuals that must cohabit. Over several years, no skin irritations or toxic effects have been observed. For long term studies, use supplementary toe-clipping: healthy A. carolinensis shed quickly, and toe clipping will avoid a confounding muddle of who is who.
8. SURGERY, INJECTIONS, AND BLOOD SAMPLING
8a. Preparation for surgery. Animals must be prepared by having sites of invasive procedures swabbed with an antiseptic such as Novolsan/alcohol or Betadine/alcohol. The only infections attending castration or neurosurgical procedures in thousand of animals in at least four laboratories have been those associated with contamination from skin flora. Similarly, swabbing before injection is advised.
The most common surgical procedures are pellet implants, castration, and brain lesioning.
Subcutaneous implants: Pellets are generally 6 or 10 mm lengths of Dow-Corning Silastic tubing (0.047-in. o.d. x 0.025-in. i.d.), providing 12.6 to 20.6 mm2 of hormone surface area, respectively. Silastic tubing is packed with hormone and sealed at both ends with Dow-Corning silicone medical adhesive. Cured hormone pellets were soaked in 95% ethanol for 30 min to remove any hormone from the implant’s surface. Hormone pellets are implanted subcutaneously dorsolaterally in cold-immobilized subjects after the site is cleansed with Betadine or Novolsan – alcohol combination and locally anaesthetized.
Castrations: Castrations are conducted on cryoimmobilized subjects using crushed ice after cleaning skin with an antiseptic and providing isoflourine inhalant for general anaesthesia. Castrations are performed by making a 5mm dorsolateral incision, palping the testes through the opening, ligation, and removal. Fine surgical silk is used to close. Animals so treated are vigorous and spontaneously active within 1-3 hours.
Brain lesions: After immobilization with hypothermia, a lizard is placed in a modified stereotaxic device. Anesthesia by Ketamine+lidocaine or Telazol+lidocaine. The upper jaw is gently clamped against a rubber-coated bite plate that serves as the reference for the horizontal plane of a brain atlas. The head is centered by rubber tipped bars fixed in the external meati with dental impression material. The center of the surface of the parietal eye is the zero reference point for the stereotaxic atlas. After making a small opening in the skull and piercing the dura, a modified platinum microelectrode is lowered to a target site. Lesions are made by radio frequency coagulator with sustained current not exceeding 25 mA for 10 sec.
8b. Immobilization and Anaesthesia. The physiology and responses of many reptiles often differs significantly from that of mammals (se Bennett 1991, for a recent comprehensive review). Immobilization facilitates experimental procedures, minimizing the time and intensity of any intrusions, anaesthesia renders the animal “insensitive to pain,” although the term “pain” is thought by many to be an inappropriate term for non-human animalsKitchell, R.L. and Erickson, H.H. 1983. What is pain? In: Kitchell and Erickson (eds), Animal Pain: Perception and Alleviation, Waverly Press, Baltimore MD, pp. vii-viii. Cited in Stevens, Craig W. 1995. An amphibian model for pain research. Lab Animal 24(10):32-36; which see for a review of the idea of pain and its neural substrate in various taxa.
Hypothermia (crushed ice, cryo immobilization)appears an effective way of rendering subjects immobile and presumably insensible for surgical procedures (McDonald 1976) This technique cannot be generalized to other taxa without consideration of the habits of the species: in species that do not have the adaptive capacity to respond to environmentally-typical freezing by profoundly altering metabolism, the technique will immobilize but may not anaesthetize.. There is evidence that in those reptiles tested, EEG recordings flatten significantly under hypothermia. Hypothermia also has the advantage of hemostasis but can be inconveniently cold and wet for the researcher. No subject has ever been lost to this procedure as contrasted with other anaesthesia that occasionally induce complicating problems and fatalities; recovery is rapid and complete where the surgical procedure has been uncomplicated by a mechanical accident. Use local surface anaesthetic (such as Opthane, xylocaine, or lidocaine) at site of surgery. Clearly more work is necessary.
Ketamine (0.2 mg / 5g lizard, intramuscular) is an effective alternative or supplement to hypothermia and can similarly be supplemented in extended procedures (and see Arena and Richardson 1990). Its mechanism of action and the characteristic dissociative/cataleptoid state induced by Ketamine is poorly understood, but appears related to altered peripheral autonomic activity, possibly secondary to blocked amine reuptake (Varano, Laforgia, De Vivo 1985). Because its effect is mainly an altered state in which some reflexes to “painful” stimuli are occasionally retained, Ketamine may only be appropriate for relatively mild procedures (Throckmorton 1981). In any event, a contact anaesthetic such as xylocaine is a necessary precaution.
8c. Injections. Injections may be administered (depending on the drug and its pathways) subcutaneously (maximum .05ml per injection; larger quantities should be delivered in divided doses beneath the loose nuchal or axillary skin) or intraperitoneal (0.1 ml maximum dose). Needles should not be larger than 25 gauge, and 27-30 is preferred.
8d. Blood Sampling. Blood sampling for stress-sensitive circulating hormones must be rapid and stress-free. Only one method, rapid decapitation, is able to achieve this. When blood is needed for determinations of such hormones or their metabolites, only rapid decapitation is appropriate, and then only if the capture and procedure is in less than 15-20 seconds. In practice this procedure appears far less painful or stressful than alternative life-sparing methods mentioned below. Once experienced in the procedure, it takes less than 10 seconds. Subjects that are not decapitated in that time are returned to their vivarium unharmed for future or alternative use. The incision is made just above the position of the heart; blood (collected from the head first) is collected in heparinized capillary tubes. In a typical (5g) lizard, 50-100µl of blood are obtained. Well known and effective alternatives for A. carolinensis are tail incision through to the caudal artery and vein (Sellers, Trauth, Lawrence 1980), and tapping the postorbital sinus with a capillary tube (MacLean, Lee, Wilson 1973). Toe-clipping does not appear to provide enough blood. These measures, however, involve handling that will affect rapidly changing stress-sensitive hormones.
9a. Physical. Decapitate individuals that appear irreversibly morbid; immediately double pith and/or crush brain unless tissue or blood samples are needed. If blood is needed, drain the head immediately. These lizards have a relatively short life span and senescent individuals appear to be starving or dehydrated. While not aesthetic, decapitation is virtually instantaneous and requires less handling (and thus stress) than an injection of barbiturate.
9b Preservation of tissue. Use 500 mg/kg to euthanize (=2.5mg/5g lizard). Inject 0.05ml commercially prepared (50mg/ml) pentobarbital sodium (Nembutal) with saline intraperitoneal and wait until animal is immobile and unresponsive. For tissue to be fixed (particularly brain), perfuse through the heart with saline followed by fixative. After exposing the heart (remove the pericardium) place the cut-off tip of a 27 gauge needle connected by a good length of limp tubing (to avoid tremors disturbing placement of tip in heart) to a syringe filled with saline; open an atrium and apply gentle pressure to syringe. Heart will swell and blood will leave through nicked atrium; when it runs clear, switch to fixative. Muscle tremors indicate that perfusate has reached muscles.
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