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Management of Reptiles

By Stephen J. Divers, BVetMed, DZooMed, DACZM, DECZM (herpetology), FRCVS, University of Georgia

Veterinarians must not only be familiar with common reptile species and their management but also be able to extract clinically relevant husbandry information from the owner in a timely manner. The use of a history form can greatly facilitate this process and ensure that nothing is overlooked.

Species:

Different species from different locations must not be mixed. Ideally, only a single species should be kept in any enclosure, and care must be exercised to avoid competition for resources such as food, basking areas, and retreats. In general, the solitary reptile pet is often the healthiest. Most nonbreeding pet snakes and aquatic turtles are best maintained as a single pet, because trauma while feeding is common in groups. Some lizards, notably the chameleons (Chameleo spp), are so territorial that isolation is often essential for longterm survival in captivity.

Enclosure:

The size of the enclosure is important, and although many breeders and retailers may be able to intensively manage stock, pet owners should be advised on minimum enclosure sizes (see Table: Recommended Minimum Space Requirements for Reptiles), the importance of providing the largest enclosure possible, and correct cage furniture. The type of enclosure (arboreal, terrestrial, subterranean, or aquatic) should be appropriate for the species (see Table: Important  Husbandry  Requirements  for  Selected  Reptiles).

Glass aquaria are commonly used, but the greater visualization perceived as an advantage to the owner may be stressful to the reptile. Glass is also a poor insulator, and greater heat loss may lead to dramatic temperature fluctuations. Even if the entire top of the enclosure is covered by mesh, ventilation may be severely hampered. Plastic or fiberglass enclosures are more expensive but more versatile.

Newspaper, artificial turf, and organic particulates (eg, bark chips) are suitable materials to line cages and vivaria, but they must be completely replaced regularly. Soil, sand, and natural leaf litter can also be used, but oven baking is recommended to sterilize before use. Gravel and pebbles are not recommended for terrestrial species, because they are difficult to clean and often ingested by reptiles. Other essential items include a water bowl (large enough for the reptile to bathe) and various retreats (eg, cardboard boxes, cork bark, shredded paper). Clean, secure branches are required for arboreal species. Soap and water is generally all that is required to clean cages, but bleach can be used as long as rinsing is thorough. Some cleansers (eg, phenolic disinfectants) are toxic. In certain areas, the climate may permit keeping reptiles in outdoor enclosures, which is highly desirable, although theft, predators, and wildlife carriers of disease should be considered.

Recommended Minimum Space Requirements for Reptiles

Reptile Group

Minimum Space Requirement

Tortoises and terrapins

0.4 m2/0.1 m carapace length

Purely aquatic chelonians

0.25 m3/0.1 m carapace length

Terrestrial lizards

0.2 m2/0.1 m total length

Arboreal lizards

0.4 m3/0.1 m total length

Boas and pythons

0.6 m2/meter of snake

Kingsnakes and rat snakes

0.6 m2/meter of snake

Whipsnakes and racers

1.2 m2/meter of snake

Arboreal snakes

0.8 m3/meter of snake

Important  Husbandry  Requirements  for  Selected  Reptiles

Species

Habitat / Vivarium Type

POTZa (°C)

Humidity %

Lightingb

Hibernation

Dietc

Corn/rat snake

Terrestrial, scrubland

25–30

30–70d

NS

Yes

C

Boa constrictor

Terrestrial, rain forest (semi-arboreal/aquatic)

28–31

70–95

NS

No

C

Ball python

Terrestrial, scrubland

25–30

50–80

NS

No

C

Leopard gecko

Terrestrial, arid scrub

25–30

20–30d

NS

No

I, c

Green iguana

Arboreal, rain forest

29–33

60–85

BS

No

H

Bearded dragon

Terrestrial, desert

20–32

20–30d

BS

No

I, c, h

Water dragon

Arboreal, rain forest

24–30

80–90

BS

No

I, c, h

Savannah monitor

Terrestrial, arid scrub

25–32

20–40d

BS

No

C, i

Greek tortoise

Terrestrial, temperate to subtropical

20–26

30–50

BS

Yes

H, c

Box tortoise

Terrestrial, temperate to subtropical

22–28

50–80

BS

Yes

H, I, c

Leopard tortoise

Terrestrial, tropical

25–30

30–50

BS

No

H, c

Red-footed tortoise

Terrestrial, tropical

25–30

50–90

BS

No

H, c

Red-eared slider

Temperate to subtropical gravel bottom or bare, water depth 30 cm min, land area ⅓ of tank

24–28

Aquatic

BS

Possible

H, I, c

a POTZ = preferred optimal temperature zone. Temperature requirements shown are air temperature gradients. In general, basking temperatures should be 5°C warmer, whereas at night these temperatures should fall by 5°C.

b BS = broad-spectrum lighting (UVB 290–300 nm) essential, NS = no special lighting requirements

c I or i = insectivorous, H or h = herbivorous, C or c = carnivorous; upper case indicates major food preference, lower case minor food preference.

d Humidity requirements are significantly greater during ecdysis.

Heating:

A variety of heaters can be used, including incandescent bulbs, infrared ceramics, heating pads or mats, warming cables, tubular heaters, radiators, convector heaters, and natural sunlight radiation. Heaters of an appropriate size should be thermostat controlled, screened from the animals, and positioned toward one end of the enclosure to provide a thermal gradient. “Hot rocks” frequently result in burns in larger animals and should be avoided. Light bulbs cannot be used to provide nocturnal heat.

Environmental Lighting:

All reptiles benefit from broad-spectrum lighting. However, UVB light (290–300 nm) is especially important for most diurnal lizards and chelonians for vitamin D3 synthesis and calcium regulation. The best source of lighting is unfiltered sunlight. However, many artificial fluorescent strip-lights, compact fluorescent, and mercury halide bulbs are available. Almost every light marketed for reptiles has the term “broad" or "full" or "natural” on its packaging, creating confusion for veterinarians and owners alike. Appropriate lights must be labeled as providing UVB or, better still, be examined and tested using a spectrometer. Even suitable fluorescent lights must be placed relatively close to the reptile and replaced every 9–12 mo. Most transparent plastic and glass barriers filter out UVB wavelengths. A photoperiod of 12 hr/day is suitable for general maintenance.

Humidity:

Humidity that is too high or low can create serious problems. Humidity is seldom directly controlled, although the advent of dedicated humidifiers and sprinkler systems makes this practical. Decreasing ventilation to maintain temperature and humidity is ill advised and frequently causes skin and respiratory disease.

Quarantine and Record Keeping:

Although the incubation period of many reptile disorders is unknown, quarantine periods of 3–6 mo are recommended. Owners should also be encouraged to keep detailed records of any changes in husbandry or nutrition, breeding activity, in-contact animals, disease outbreaks, health issues, and previous treatments.

Nutrition:

Species identification is essential to critically appraise captive diets (see Table: Composition of Animal Foods that May Be Offered to Reptiles and see Table: Composition of Plant Foods that May Be Offered to Reptiles). Rodent-eating carnivores such as most snakes present few problems as long as the rodent is recognizable to the snake as food. Obese rodents that have been frozen for prolonged periods may contain fewer nutrients. Feeding live rodents is not advised (and is illegal in some countries) because of the dangers of prey-induced trauma to sedate reptiles and the welfare implications for the live prey. Insectivorous reptiles can be well catered for with a variety of commercially available crickets, waxworms, tebos, locusts, mealworms, cockroaches, and flies. Feeding insects a nutritional diet high in calcium and dusting the insects with a high-calcium reptile supplement immediately before feeding is required to prevent deficiencies.

Composition of Animal Foods that May Be Offered to Reptiles

Food Item

Dry Matter (%)

Protein (%)

Fat (%)

Energy (Kcal/g)

Calcium (%)

Phosphorus (%)

Ca:P Ratio

Mealworms (Tenibrio)

42.2

52.8

35

6.53

0.06

0.53

0.11

Superworms (Zophoba)

40.6

17.4

17.9

0.01

0.02

0.43

Locusts

31.2

61.7

19.4

0.1

0.75

0.13

Crickets

38.2

55.3

30.2

0.23

0.74

0.31

Earthworms

22

49.9

5.8

0.59

0.85

0.69

Waxworms

38.3

15.5

22.2

0.03

0.22

0.13

Chicken muscle

25.6

20.5

4.3

1.21

0.01

0.2

0.05

Egg, whole

25.2

12.3

10.9

1.47

0.05

0.22

0.02

Mice, 1–2 days old

1.6

1.8

0.88

Mice, adult

19.86

8.81

2.07

0.84

0.61

1.37

Providing herbivorous reptiles with a varied and nutritious diet can be difficult. Foods with a high calcium:phosphorus ratio should be selected with due regard given to the species-specific requirements for vegetables and fruits. Human nutrient databases can be useful (http://ndb.nal.usda.gov/). Calcium or vitamin D3 deficiency (generally due to poor quality lighting and low-calcium diets) leads to secondary nutritional hyperparathyroidism in insectivores and herbivores. A variety of commercially available foods are available in moist, canned, and dry pellet forms. These may help provide a balanced diet, but they have not been critically evaluated.

Composition of Plant Foods that May Be Offered to Reptiles

Food Item

Dry Matter (%)

Protein (%)

Fat (%)

Energy (Kcal/g)

Calcium (%)

Phosphorus (%)

Ca:P Ratio

Alfalfa

15.5

37.1

3.94

1.29

0.21

6.14

Applea

0.2

0.6

0.57

0

0

0.57

Bananaa

29.3

1.1

0.3

0.79

0

0.02

0.25

Black currants

0.06

0.04

1.4

Blackberry

0.06

0.02

2.62

Broccoli

3.6

0.3

0.1

0.06

1.49

Cabbage

1.3

0.2

0.04

0.03

1.22

Carrots

10.1

0.7

0.23

0.04

0.02

2.29

Clover hay

11

1.9

1

0.2

5.0

Collard

0.2

0.07

2.76

Cranberries

0.01

0.01

1.36

Damsons

0.02

0.01

1.5

Dandelions

0.18

0.07

2.4

Endive

1.7

0.1

0.08

0.03

2.67

Fennel

2.8

0.4

0.1

0.05

1.96

Fig

0.28

0.09

3.04

Grass (lawn)

33

2.4

1.2

1.58

0.1

0.09

1.1

Iceberg lettuce

1.2

2.5

0.14

0.03

0.02

1.34

Kale

0.17

0.06

2.9

Lemon

0.11

0.01

9.17

Lettucea

4.1

1

0.4

0.12

0.02

0.02

0.85

Mustard cress

0.06

0.06

1

Orange

13.9

0.8

0.35

0.04

0.02

1.71

Parsley

0.2

0.13

1.53

Radish

0.04

0.02

1.63

Raspberry

0.04

0.02

1.41

Red currants

0.03

0.03

1.2

Spinach

3

0.3

0.09

0.05

1.69

Tomatoa

6.6

0.9

trace

0.14

0.01

0.02

0.62

Turnip

1.1

0.9

0.05

0.01

2.89

Watercress

2.2

0.3

0.22

0.05

4.23

White grapea

20.7

0.6

trace

0.63

0.01

0.02

0.86

a The poor Ca:P ratio of these commonly used items may make them less suitable as a staple component of a reptile diet.

Although some species will drink from a water bowl, others will only imbibe water droplets on plants and décor. Poor water quality has been implicated as a cause of stomatitis in snakes. Lack of appropriate water delivery has been implicated as a predisposing cause of renal disease in green iguanas. The advent of timer-controlled sprinkler systems makes regular water provision possible for many of these more fastidious drinkers.

Physical Examination

The successful diagnosis and treatment of reptile diseases requires proper restraint and performance of a variety of clinical techniques. Although the principles are similar to those used for domestic animals, there are a number of reptile-specific peculiarities. It may be possible to observe calm specimens unrestrained, permitting an assessment of demeanor, locomotion, and obvious neurologic disorders such as lameness, paralysis, paresis, and head tilt. Observation of reptiles within their usual environment is particularly valuable and should be done whenever possible. Nervous or aggressive species are best restrained at all times using towels, snake hooks, clear plastic containers, and restraint tubes. Gauntlets severely reduce the clinician’s tactile sensation but may be required when dealing with large lizards or small to medium-sized aggressive crocodilians. Careful consideration should be given to the safety of veterinary staff, zoo keepers, and private owners when dealing with large or otherwise potentially dangerous reptiles. In many cases, chemical agents can expedite procedures and considerably reduce risks to both the reptile and human handlers. Given the improvements in reptile anesthesia, even manageable reptiles may be preferentially sedated or anesthetized for procedures that would otherwise take longer to accomplish and cause unnecessary stress or discomfort to the animal. It is possible that sedatives and anesthetics may affect clinical pathologic results, especially hematology.

The decision to examine a potentially dangerous reptile should be made with due regard to legislative and safety requirements. No species of Chelonia is legally considered dangerous, but several species (eg, snapping turtles, Chelydra spp) have a ferocious bite that makes them a formidable opponent. In addition, the Convention on the International Trade of Endangered Species of Fauna and Flora (CITES) may also have implications for Appendix 1 and 2 reptiles kept as pets. Even some common pet species (eg, corn snake) may be illegal in some endemic areas, whereas native venomous snakes may be freely collected because they are considered “vermin.”

The risks of reptile-borne zoonoses are probably no greater than for other animal groups, and basic personal hygiene after handling reptile patients will minimize these risks. The major zoonoses include Salmonella, Pseudomonas, Mycobacterium, Cryptosporidium, and Rickettsia spp and pentastomids (arachnid lung parasites). Major public concern centers on the commensal reptile Salmonella spp, and clinicians are advised to obtain a copy of the statement policy and client brochure on this subject produced by the Association of Reptilian and Amphibian Veterinarians (http://www.arav.org/special-topics/).

Every reptile must be accurately weighed; an accurate weight is important to avoid deaths associated with drug overdose, particularly anesthetics and aminoglycosides. In addition, serial weight measurements permit an appraisal of growth and captive management, response to treatment, and disease progression or resolution. Relating body weight to length and conformation gives an assessment of body condition. The snout-vent length of lizards and especially snakes is worth noting, because organ position and growth can be calculated as a result. Chelonian body condition relies on relating total weight to straight carapace length or body volume.

Transillumination of the coelom using a cold light source can be used to visualize the internal structures of small lizards and snakes and is particularly useful to confirm suspected impactions and foreign bodies. Care must be exercised if a hot light source (eg, incandescent spotlight) is used, because of the possibility of burns.

Auscultation of reptiles is difficult and often unrewarding. Electronic stethoscopes with moistened gauze between the shell or scales and the stethoscope diaphragm can be helpful. Doppler ultrasound is particularly useful to determine heart rates.

Snakes:

The head of an aggressive snake or a snake of unknown disposition should be identified and restrained before opening the transportation bag to remove the animal. In general, the head of the snake is held behind the occiput using the thumb and middle finger to support the lateral aspects of the cranium. The index finger is placed on top of the head. The other hand is used to support the body. Restraining the snake’s head in this manner supports the cranial-cervical junction, which, having only a single occiput, is prone to dislocation. When dealing with large boids, a second, third, or even fourth handler is required to support the body during the examination. It is usually safer and more convenient to sedate a large, pugnacious snake than to risk injury to the snake, owner, or staff.

Nonvenomous species should be supported using one or two hands, depending on size. Nervous or aggressive snakes can be restrained using plexiglass tubes or sedated before examination. The clinician should attempt to gauge muscle tone, proprioception, and mobility. Systemically ill serpents will often be limp, lack strength, and be less mobile. Head carriage, body posture, cloacal tone, proprioception, skin pinch, withdrawal, and papillary and righting reflexes can be used to assess neurologic function.

The entire integument, particularly the head and ventral scales, should be thoroughly examined for evidence of dysecdysis (poor shedding), trauma, parasitism (especially the common snake mite, Ophionyssus natricis, and ticks), and microbiologic infection. Any recently shed skin should also be examined, if available, for evidence of retained spectacles. Skin tenting and ridges may indicate cachexia (“poverty lines”) or dehydration; ticks and mites may congregate in skin folds, infraorbital pits, nostrils, and corneal rims. The infraorbital pits (where present) and the nostrils should be free from discharges or retained skin. The eyes should be clear, unless ecdysis is imminent. The spectacles covering the eyes should be smooth; any wrinkles usually indicate the presence of a retained spectacle. The spectacle represents the transparent fused eyelids, and therefore the cornea is not normally exposed. The subspectacular fluid drains through a duct to the cranial roof of the maxilla. When blocked, the buildup of fluid causes a subspectacular swelling that often becomes infected. Damage to the underlying cornea can result in panophthalmitis and ocular swelling. Retrobulbar abscessation results in protrusion of a normal-sized globe. Other ocular pathologies can include uveitis, corneal lipidosis, and spectacular foreign bodies, including slivers of wood or other vivarium materials.

Working from cranial to caudal, the head and body are palpated for swellings, wounds, and other abnormalities. The position of any internal anomalies, noted as a distance from the snout and interpreted as a percentage of snout-vent length, enables an assessment of possible organ involvement. Recently fed snakes have a midbody swelling associated with the prey within the stomach; handling such individuals may well lead to regurgitation. Preovulatory follicles, eggs, feces, enlarged organs, and masses may be palpable. The cloaca can be examined using a dedicated otoscope or by digital palpation.

Examination of the oral cavity is often left until last, because many snakes object to such manipulation. However, even before the mouth is opened, the tongue should be seen flicking in and out of the labial notch with regularity. The mouth can be gently opened using a plastic or wooden spatula to permit an assessment of mucous membrane color and the buccal cavity for evidence of mucosal edema, ptyalism, hemorrhage, necrosis, and inspissated exudates. White deposits may indicate uric acid deposition due to visceral gout. The pharynx and glottis should be examined for hemorrhage, foreign bodies, parasites, and discharges. Open-mouth breathing is a reliable indicator of severe respiratory compromise. The patency of the internal nares and the state of the polyphyodontic teeth should be noted.

Lizards:

Lizards vary considerably in size, strength, and temperament; therefore, a variety of handling techniques are required. The tegus and monitors are renowned for their powerful bites, whereas other species, particularly the green iguana, are much more likely to use their claws and tail. The main problem when handling small lizards is restraining them before they flee. The lizard should be transported in a securely tied cloth bag, so that the position of the lizard can be identified and the lizard held before the bag is opened. Large lizards are best restrained with the forelimbs held laterally against their coelom and the hindlimbs held laterally against the tail base. The limbs should never be held over the spine, because fractures and dislocations can occur. Nervous lizards can be wrapped in a towel to aid restraint. Smaller lizards can be restrained around the pectoral girdle, holding the forelimbs against the coelom, although care is required not to impair respiratory movements. A lizard should never be grasped by the tail, because many species can drop the tail (autotomy) in an attempt to evade capture. Restricting the vision of a lizard (eg, a towel placed over the head) is often the simplest way to facilitate handling and examination. A useful restraint technique for iguanid or monitor lizards uses the vasovagal response: gentle digital pressure applied to both orbits causes many lizards to enter a state of stupor for up to 45 min (or until a painful or noisy stimulus is applied). This technique enables the mouth to be gently opened without the need for excessive force. If possible, the lizard should be observed unrestrained to check for neurologic problems. Calm lizards may be permitted to walk around the examination table or on the floor. However, if in any doubt, the lizard should be placed in a large plastic enclosure to prevent escape during observation.

The integument should be examined for parasites (essentially mites and ticks) and trauma due to fighting, mating, and burns. Lizards tend to shed their skin in stages. Classically, dysecdysis and skin retention occurs around the digits and tail, causing ischemic necrosis. Extensive skin folding and tenting may indicate cachexia and dehydration.

The head should be examined for abnormal conformation. The mouth can be opened using a blunt spatula or, in iguanas, by gently applying pressure to the dewlap. The buccal cavity and glottis should be examined thoroughly for evidence of trauma, infection, neoplasia, and edema, especially pharyngeal edema. The internal extent of any rostral abrasions should be evaluated. The nostrils, eyes, and tympanic scales should be clean and free of discharges. The presence of dry, white material around the nostrils of some iguanid lizards is normal, because some species excrete salt through specialized nasal glands. The rostrum should be examined for trauma, often caused by repeated attempts to escape from a poorly designed vivarium or to evade cagemates. The head, body, and limbs should be palpated for masses or swellings, which can be abscesses or metabolic bone disorders. Lizards suffering from severe hypocalcemia and hyperphosphatemia may exhibit periodic tremors and muscle fasciculations. The coelomic body cavity of most lizards can be gently palpated. Food and fecal material within the GI tract, fat bodies, liver, ova, and eggs are usually appreciable. Cystic calculi, fecoliths, enlarged kidneys, impactions, retained eggs or ova, and unusual coelomic masses may also be noted.

The cloaca should be free from fecal staining, with visual and digital examination considered routine. In the green iguana, renomegaly can be appreciated by digital cloacal palpation. The high incidence of dystocia necessitates a need to identify gender during examination. Many species of lizards are sexually dimorphic, although sexing juveniles can be difficult.

Tortoises, Turtles, and Terrapins:

Small to medium-sized tortoises are not difficult to handle, although their strength and uncooperative nature can hinder examination. Patiently holding the tortoise with its head down will often persuade a shy individual to protrude the head from the shell. Placing the thumb and middle finger behind the occipital condyles prevents retraction of the head. However, with larger species, it may be physically impossible to prevent a strong individual from pulling free. In such cases, sedation or use of a neuromuscular blocking agent may be necessary. The more aggressive aquatic species should be held at the rear of the carapace. Some species (eg, snapping turtles) have long necks and an extremely powerful bite, necessitating great care. Certain species also have functional hinges at the front and/or back of the plastron, and care should be exercised not to trap a finger when the hinge closes.

Examination of the head should include the nostrils for any discharges and the beak for damage and overgrowth. The eyelids should be open and not obviously distended or inflamed, and the eyes should be clear and bright. Conjunctivitis, corneal ulceration, and opacities are frequent presentations. The tympanic scales should be examined for signs of swelling associated with aural abscessation. Applying steady distractive pressure to the maxilla and mandible can open the mouth, and a mouth gag can be inserted to prevent closure. Aggressive chelonians, generally aquatic species, often threaten by open-mouth displays, which provide a good opportunity to examine the buccal cavity with minimal handling. Mucus membrane coloration is normally pale; hyperemia may be associated with septicemia or toxemia. Icterus is rare but may occur with biliverdinemia due to severe liver disease. Pale deposits within the oral membranes may represent infection or urate tophi associated with visceral gout. The glottis is positioned at the back of the fleshy tongue and may be difficult to visualize; however, it is important to check for any inflammation and glottal discharges consistent with respiratory disease.

The integument should be free of damage. Subcutaneous swellings are usually abscesses. Aquatic species appear more susceptible to superficial and deep mycotic dermatitis, especially around the head, neck, and limbs. The withdrawn limbs can be extended from the shell of small to medium-sized chelonians by applying steady traction. Because the coelomic space within the shell is restricted, gently forcing the hindlimbs into the shell will often lead to partial protrusion of the forelimbs and head, and vice versa. A wedge or mouth gag can be used to prevent complete closure of a hinge. No chelonian will close a hinge on an extended limb. The integument should be examined for parasites (particularly ticks and flies), dysecdysis, trauma, and infection that may arise due to predator attacks. Aggressive conflicts and courting trauma must also be considered in the communal environment. Limb fractures are less common in chelonians than in other reptiles, but when they do occur they are often associated with rough handling and secondary nutritional hyperparathyroidism. Focal subcutaneous swellings are usually abscesses, but grossly swollen joints or limbs are more often cases of fracture, osteomyelitis, or septic arthritis.

The prefemoral fossae should be palpated with the chelonian held head-up. Gently rocking the animal may enable palpation of eggs, cystic calculi, or other coelomic masses. The shell should be examined for hardness, poor conformation, trauma, or infection. Soft, poorly mineralized shells are usually a result of secondary nutritional hyperparathyroidism resulting from dietary deficiencies of calcium, excess phosphorus, or a lack of full-spectrum lighting. Pyramiding of the shell appears to be more associated with inappropriate humidity than dietary imbalances. Shell infection may present as loosening and softening of the scutes with erythema, petechiae, purulent or caseous discharges, and a foul odor.

Prolapses through the vent are obvious, but it is necessary to determine the structure(s) involved. Prolapses may include cloacal tissue, shell gland, colon, bladder, or phallus. Internal examination using digital palpation and an endoscope is recommended.

Anesthesia and Analgesia

For some minor procedures (eg, blood sampling), simple restraint may be all that is required. This can be enhanced by temporary immobilization techniques such as dorsal recumbency, reduced light intensity, or gentle ocular pressure (vasovagal response). For more invasive and painful procedures, general anesthesia must be used. Although considerable anatomic, physiologic, and pharmacologic differences exist between reptiles, some general guidelines are applicable. The following is therefore intended as a practical approach, rather than an exhaustive review of reptile anesthesia.

Preanesthetic Assessment and Stabilization:

All reptiles should be hospitalized and maintained at their preferred optimal temperature zone at all times to minimize physiologic disturbance, facilitate recovery, and maintain immunocompetence. Although hypothermia will reduce movement, it does not provide analgesia and is therefore unacceptable as a means of anesthesia on welfare grounds. It can also dramatically affect the pharmacokinetics of any drugs administered and greatly prolong recovery.

A full clinical examination should be performed and the animal accurately weighed, although this may not be practical or possible in some cases. The hydration status of all reptiles should be assessed, especially if debilitated or post-hibernation. For elective procedures (eg, neutering), underweight, dehydrated, or debilitated animals should be nursed for days, weeks, or months until their condition improves. For nonelective surgery, dehydration should be corrected before anesthesia. Even the most moribund egg-bound reptile will usually benefit from stabilization for 24–48 hr before surgery is performed. Reptiles that have not been stabilized before surgery tend to succumb intra- or postoperatively. Although oral fluids are least invasive to administer and provide the most physiologically normal method of rehydration, they are sufficient only for mildly dehydrated animals and are contraindicated immediately before surgery because of the risks associated with regurgitation. Intracoelomic fluids are more suitable, but uptake can take many hours and their use is problematic if coeliotomy or coelioscopy is planned. For dehydrated surgical candidates, IV or intraosseous fluid therapy should be administered before, during, and after surgery as necessary.

Reptiles should be fasted before all elective surgery to avoid the compression of lung(s) associated with large meals and potential regurgitation. Fasting depends on the feeding regimen of the reptile, but in general one feeding cycle should be skipped before surgery. Premedication with sedatives such as acepromazine and atropine is generally rare in reptiles, although midazolam has been advocated in large chelonians. However, presurgical administration of analgesics should be considered routine (see Table: Analgesics, Sedatives, and Anesthetics Used in Reptiles).

Anesthetic Induction:

IV or intraosseous propofol provides a rapid, controlled mode of induction. It is relatively nontoxic, and there is reduced risk of thrombophlebitis if it is injected perivascularly. This is of particular concern, because IV access may be relatively difficult, especially in active animals undergoing elective procedures. Alfaxalone has been licensed in the USA; it is an excellent IV induction agent and is also effective when used IM.

If IV access is impractical or dangerous to attempt, IM agents can be used to induce sufficient chemical restraint for intubation. For IM injections in lizards and chelonians, the forelimb muscles are preferred, whereas for snakes, the epaxial muscles are used. An IM combination of ketamine, medetomidine (or dexmedetomidine), and morphine or hydromorphone has proved effective for a variety of chelonians; this can be readily reversed using atipamezole and, if necessary, naloxone or naltrexone.

Squamates can also be induced by inhalation agents in an induction chamber or by mask. However, breath holding tends to occur in turtles and crocodilians, which can respire anaerobically for prolonged periods. Induction may take 10–30 min in cooperative lizards and snakes. Intubation of conscious patients has been suggested after local lidocaine spray, but the adverse effects of increased stress and catecholamine release should always be considered. There is also the potential danger of being bitten.

Maintenance of Anesthesia:

Isoflurane or sevoflurane are the agents of choice for maintenance of anesthesia. These volatile inhalation agents have faster modes of action, are more controllable, and facilitate faster recoveries than most alternatives. Furthermore, their lack of reliance on hepatic metabolism or renal excretion reduces the anesthetic risk to debilitated reptiles or those with questionable renal or hepatic function.

Analgesics, Sedatives, and Anesthetics Used in Reptiles

Drug

Dose and Route

Comments

Morphine

1.5 mg/kg, IM, SC

10 mg/kg, IM, SC

Chelonians (red-eared sliders)

Lizards (bearded dragons)

Not analgesic for snakes. Causes pronounced respiratory depression in turtles.

Hydromorphone

0.5 mg/kg, IM, SC

Chelonians: appears to cause less respiratory depression than morphine

Tramadol

5 mg/kg/day, PO, SC;

10 mg/kg, PO, every 4 days

Chelonians (red-eared sliders); less respiratory depression than morphine

Butorphanol

20 mg/kg, SC

Snakes (cornsnakes) at high doses. Not analgesic for bearded dragons or red-eared sliders. May cause respiratory depression, and high-dose volume may be impractical. May be able to partially antagonize opiate agonists like morphine.

Meloxicam

0.2 mg/kg, IV, IM, SC, every 24–48 hr

Ketamine

10–40 mg/kg, IM

40–60 mg/kg, IM

Sedation, prolonged hangover effects

Deeper sedation but not sufficient for painful procedures and care in debilitated individuals

Ketamine 10 mg/kg, combined with medetomidine 0.1–0.2 mg/kg (or dexmedetomidine 0.05–0.1 mg/kg) and morphine 1.5 mg/kg or hydromorphone 0.5 mg/kg, IM (or 50% dose, IV)

Deep sedation/anesthesia in many chelonians. Reversed using atipamezole (0.5 mg/kg, IM) and, if necessary, naloxone (0.2 mg/kg, IM)

Midazolam

2 mg/kg, IM

Premedication, or can be given with ketamine to increase sedative effects or promote similar effects at lower ketamine doses.

Dexmedetomidine

0.05–0.1 mg/kg, IV or IM

Often used in combination with ketamine and opiates as a sedative to light anesthetic. Useful when IV access is difficult.

Atipamezole

10 × mg dose of dexmedetomidine, IM

Used to reverse dexmedetomidine

Tiletamine/zolazepam

3–12 mg/kg, IM

Tortoises, lizards, snakes. Low dose useful to facilitate intubation. Higher doses associated with prolonged recoveries.

Propofol

3–10 mg/kg, IV, intraosseous

Low dose rate for larger reptiles. Subanesthetic doses produce variable short-term sedation.

Alfaxalone

5–10 mg/kg, IV

10–20 mg/kg, IM

Similar effects to those of propofol IV, but higher doses effective IM. Larger IM dose volumes may warrant dividing into two or more injections.

Isoflurane

1%–5%

Routine gaseous agent; subanesthetic levels provide short-term sedation. Mask down or conscious (sedated) intubation possible in some species.

Sevoflurane

2%–7%

Very similar effects to those of isoflurane but recoveries appear to be faster. Preferred agent for critical or large reptiles.

Intubation of reptiles is relatively simple. Small-gauge endotracheal tubes or catheters are easily inserted through the glottis immediately caudal to the tongue; this may be aided by forcing the tongue up and forward by pressing a finger into the intermandibular space from under the jaw. The reptilian glottis is actively dilated, and therefore its movement will often be abolished in anesthetized animals; a guiding stylet can be useful to facilitate endotracheal tube placement. The bifurcation of the trachea may be far craniad in some chelonians, and gaseous exchange has also been reported within the tracheal lung of some snakes; care should be taken to use a short endotracheal tube that is securely taped in position.

Noncrocodilian reptiles lack a diaphragm; skeletal intercostal muscles (Squamata) or limb movements (Chelonia) control breathing. The action of these muscles is abolished at a surgical plane of anesthesia, and intermittent positive-pressure ventilation is required. Ventilation rates should initially mirror preanesthesia evaluations and then be adjusted to maintain end-tidal capnography readings of 15–25 mmHg. Electrical ventilators enable precise control of ventilation rates and pressures.

Monitoring anesthesia in reptiles differs considerably from doing so in mammals. Palpebral and corneal reflexes are reliable in those species in which they can be elicited (ie, all chelonians, all crocodilians, most lizards, but no snakes). However, corneal reflexes are abolished at toxic doses, and pupillary diameter may bear little relation to the depth of anesthesia (unless fixed and dilated, which indicates excessive anesthetic depth or brain anoxia and death). Jaw tone and withdrawal reflexes (tongue, limb, or tail) are abolished only at a surgical plane of anesthesia. This also correlates with full loss of the righting reflex, loss of spontaneous movement, and complete muscle relaxation.

Heart rate may be monitored by auscultation or by visualization or palpation of the heart beat in most snakes and some lizards. Pulse oximetry, using either an esophageal or cloacal reflectance probe, is useful to monitor pulse rate and strength. Although the blood oxygen saturation (SpO2) readings are often low and have not been validated for reptiles, monitoring the trend in SpO2 is often helpful. Doppler ultrasonography can also be used over peripheral arteries or the heart. Blood gas estimations are often affected by intracardiac or pulmonary shunts, especially in aquatic species. However, end-tidal capnography has proved effective.

Toward the end of surgery, the anesthetic gas should be discontinued while maintaining ventilation for another 5–10 min to facilitate excretion. At this point, oxygen should be discontinued in favor of room air delivered by bag-valve mask to encourage spontaneous respiration.

Postoperative Support:

Once it is breathing spontaneously, the reptile can be returned to an incubator or vivarium to fully recover. Continued monitoring is essential until righting reflexes return and the animal is ambulatory. Additional analgesia, fluid, and nutritional support should be provided as indicated.

Diagnostic Techniques

Radiology:

Several anatomic differences make it difficult to obtain quality radiographs in reptiles. The relatively small size of most pet reptiles and the lack of diffuse body fat often produce images of poor contrast. Thick, highly keratinized scales, osteoderms, or shells can severely hinder the x-ray beam, necessitating greater power and a subsequent loss of fine soft-tissue detail.

Despite these difficulties, most high-capacity units can be set to produce quality radiographs of reptiles. High-detailed screen/film combinations (eg, mammography film) are essential to obtain sufficient detail and contrast, especially in smaller animals. Various agents can be used to improve contrast. Barium sulfate (30%) can be used for GI studies. Water-soluble iodine compounds such as iohexol can be used for GI, urogenital, and IV techniques. The injection of air into the coelom of a lizard can greatly improve the appreciation of preovulatory follicles.

Snakes:

Snakes can be difficult to position and restrain for radiographic examinations unless anesthetized. If the purpose of the examination is simply to exclude radiodense foreign bodies, the snake may be allowed to coil in its natural position. If detailed examination of the skeletal, respiratory, and digestive system is desired, the snake must be extended. A plastic restraint tube can be used for this purpose; however, this may produce some radiographic artifact. In larger snakes, several films will be needed to radiograph the entire length of the body. Lateral views are best taken using horizontal beams to avoid displacement artifact of the viscera. However, standard laterals with the snake taped in lateral recumbency can be useful, especially when horizontal beams are not possible or safe to undertake. The interpretation of dorsoventral views are hindered by the spine and ribs but can still be useful when dealing with obvious lesions, including eggs and mineralized masses.

Lizards:

Small lizards can often be restrained by taping them to the radiography film or table for a dorsoventral view. Placing cotton balls over the eyes, and wrapping them with self-adhesive tape will often produce a calm, motionless lizard. A dorsoventral view can help identify foreign bodies, intestinal impaction, or coelomic masses. A horizontal x-ray beam provides the best lateral imaging in lizards, especially when evaluating the respiratory system. The positioning for this view involves rotating the x-ray tube 90° and placing the film vertically behind the lizard. Elevating the body of the lizard on rolled towels or foam pads helps prevent superimposition of the limbs with the coelomic cavity. The positioning for, and interpretation of, crocodilian radiographs are similar to those used for lizards.

Chelonians:

Most chelonians are fairly easy to position and restrain. For vertical beam dorsoventral radiographs, most conscious animals will remain motionless long enough to permit exposure. Ideally, the head and limbs should be extended from the shell to reduce superimposition of the limb musculature on the coelomic viscera. More active animals can be restrained by taping them to the cassette or by placing them in a radiolucent container, although this should be avoided with smaller specimens (and lower exposures), because material artifacts may appear. For lateral horizontal beam radiographs, the chelonian is best placed on a central plastron stand to encourage extension of the limbs and head while the tortoise remains immobile. Both left and right lateral projections should be taken with the lateral edge of the shell touching (or as close as possible to) the cassette. The third basic coelomic view is the horizontal craniocaudal (or anterior-posterior) view. The chelonian is positioned on a central plastron stand, with the caudal edge of the carapace touching (or as close as possible to) the cassette; the head should be facing the x-ray tube and the beam centered on the midline of the cranial rim of the carapace. Radiology of the head and limbs requires exteriorization from the shell, usually under general anesthesia. The use of sandbags, foam, and tape aid positioning. Standard interpretation requires that both true lateral and dorsoventral views should be taken—even slight rotation makes interpretation more difficult.

Ultrasonography:

A useful and often underrated technique, ultrasonography has gained popularity as a diagnostic technique for reptiles. It is particularly useful for examining tissue parenchyma, guiding biopsy needles, and, with color flow Doppler, investigating cardiac disease. Unfortunately, the equipment tends to be expensive, and several probes with small footprints are required, given the variation in reptile size and ultrasound applications. Ultrasound waves cannot penetrate through mineralized tissue or air and, therefore, ultrasonography has obvious limitations to investigate respiratory and GI diseases.

The giant species require a 5-MHz probe, whereas a 7.5-MHz probe suffices for most reptiles. When examining very small specimens (or for ultrasonography of eyes), a 10–20 MHz probe is more appropriate. Good contact and imaging generally require copious quantities of gel or a water bath. It is helpful to maintain the animal in a normal position or, failing that, at least appreciate the complications associated with organ displacement. Ultrasonography can be a useful adjunct to radiography, especially to assess the reproductive tract (evaluating ovarian activity and distinguishing between pre- and postovulatory egg stasis), liver and gallbladder, urinary system, soft-tissue masses, and heart. Ultrasonography has been used to guide liver biopsy, although iatrogenic trauma has been reported in snakes.

CT and MRI:

CT offers excellent, high-resolution, detailed images. Potentially, it would be the diagnostic imaging technique of choice for the respiratory system and skeleton of reptiles. MRI produces primarily high-detailed images of soft-tissue structures and is useful to diagnose neurologic, hepatic, renal, and reproductive diseases in reptiles. Disadvantages of these techniques are the need for general anesthesia to completely eliminate motion, equipment availability, and cost.

Endoscopy:

Endoscopy has proved to be a most useful diagnostic tool in reptile medicine, and given the small and delicate nature of many species, continued development of minimally invasive techniques is likely. Flexible endoscopes are useful for respiratory endoscopy in snakes or GI endoscopy in many species. The main disadvantage of the flexible, fiberoptic endoscope is the poorer image quality than for those obtained using rigid scopes of a similar diameter. However, the continued development of smaller videoscopes looks likely to redefine flexible endoscopy in reptiles. The compact body size of most pet reptiles, coupled with their coelomic body design, makes rigid endoscopy useful in many situations. Although equipment must be matched to size of the reptile, in general the 1.9-mm and 2.7-mm telescope and sheath systems work well for most pet species, enabling gas insufflation, fluid irrigation, and biopsy capability.

Insufflation is essential to provide the lens-organ distance required for visualization. For GI endoscopy, air and/or saline is used; for coelioscopy, medical grade CO2 or saline is preferred. Coelomic pressures seldom need to be >5 mmHg. When performing endoscopy in small neonates or within a hollow viscus (eg, bladder, oviduct, cloaca, stomach), warmed sterile saline irrigation often provides better clarity than gas insufflation.

General anesthesia is recommended for all endoscopy procedures. Certain examinations (eg, buccal cavity and cloaca) may be possible in a conscious or sedated animal using a mouth gag or other appropriate restraint, but complete immobilization is preferred to avoid risking damage to patient, staff, or equipment. Anesthesia is mandatory for coelioscopy.

Blood Collection:

Venipuncture is generally a blind technique in reptiles. Up to 0.5 mL/100 g may be safely collected from healthy reptiles, less in debilitated animals. There is a relative lack of hematologic or biochemical data for most reptiles. In addition, blood values can vary dramatically with species, environment, nutrition, age, breeding, and hibernation. Given this variability, published ranges may be of limited value. More reliance should be placed on establishing an individual’s observed range and using serial sampling to monitor the progress of hematologic and biochemical changes. The use of a toenail clip to obtain blood may result in fecal or urate contamination, increased tissue enzymes, and hemogram and electrolyte changes due to the peripheral nature of the sample and the crushing artifact of collection. Of even greater concern are the ethical and welfare issues associated with toenail clipping.

The two common sites for venipuncture in snakes are the caudal vein and the heart. The caudal vein is accessed caudal to the cloaca, between 25% and 50% down the tail, and avoiding the paired hemipenes of males. In lizards, the most clinically useful vessel is the ventral midline caudal vein, best accessed 20%–80% down the tail. The most clinically useful vessels in chelonians appear to be the jugulars, subcarapacial sinus, and dorsal coccygeal veins. The left and right jugular veins are preferred because of the reduced risk of lymphatic contamination.

Necropsy:

A detailed necropsy should be undertaken whenever possible, because it often provides a definitive diagnosis. When managing a disease outbreak in a population, elective euthanasia and necropsy of one or more individuals is often the most efficient and cost-effective means of diagnosis. Fresh necropsies can provide organ biopsies, blood, and other bodily fluids for laboratory examination. The submission of microbiology samples, especially bacteriology, from reptiles that have died and remained within a heated enclosure must, however, be interpreted with caution.

Surgery

In general, performing surgery on a reptile patient should be approached with the same principles as those used for domestic animals. However, there are some specific anatomic considerations, as well as unique aspects of patient preparation, positioning, and equipment. Only a basic discussion follows here, and consulting other sources on reptile anatomy, physiology, husbandry, anesthesia, and surgery, as well as domestic animal surgery literature, is essential before performing surgery on any reptile.

For truly giant reptiles, such as giant tortoises, the use of stronger, large animal instruments is recommended. For reptiles weighing 5–50 kg, most instruments used for small animals are appropriate. However, most pet reptiles weigh <1 kg, and microsurgical instruments are often required. These instruments are not miniaturized versions of standard instruments but rather balanced instruments with fine, small tips. Because microinstruments can be costly, ophthalmologic instruments can be useful alternatives. Plastic, self-retaining retractors can be adjusted to fit different sizes of incisions and do not compromise ventilation. Smaller versions of standard abdominal retractors can also be used but are significantly heavier. Eyelid retractors can be useful to retract coelomic incisions in small lizards and snakes. Epoxy resins or low-temperature veterinary acrylics are used for many chelonian plastron closures and shell repairs.

Rapidly absorbable suture materials are recommended for internal soft-tissue applications. For prolonged internal durability, polydioxanone or nylon are preferred. Poliglecaprone, monofilament nylon, and polydioxanone are favored for skin suturing, although wire may be necessary for giant tortoises or monitor lizards.

Because most reptile patients that require surgery are significantly smaller than mammalian patients, some degree of magnification is generally recommended. Headband or frame-mounted operating loupes (2–4 × magnification) with a dedicated halogen or xenon light source are affordable, versatile, comfortable, and simple to use. While operating, microscopy telescopes can provide magnification with selected rigid endoscopy systems.

A healthy reptile can generally tolerate 0.4–0.8 mL/100 g body wt of blood loss. Reptiles in need of surgery are often compromised, and diagnostic blood samples may have been collected before surgery. Therefore, the amount of blood a reptile can afford to lose during surgery may be considerably less. Careful consideration must be given to minimizing hemorrhage using cotton-tipped spears or applicators, vascular clips, and radiosurgery.

Patient positioning will depend on the species and the nature of the surgery. Consideration should be given to ensuring that head and neck position does not interfere with ventilation; avoiding excessive compression of the head, limbs, or coelom to prevent pressure necrosis, visceral rupture, or hypoventilation of the lungs; avoiding extreme and prolonged hyperextension or hyperflexion of any joint; and ensuring that the surgical site is easily accessible and does not require surgeon positioning that will result in fatigue. Sand-bags, bean-bags, foam supports, and adhesive tapes can be used to maintain patient position.

When reptile skin is incised, it tends to invert. Therefore, everting suture patterns (eg, horizontal mattress) are recommended to ensure opposition of tissue without future dysecdysis. The skin suture materials should be monofilament. Wire suture may be required for repairs involving shell or thickly keratinized skin containing osteoderms. Staples have also been advocated, because they cause mild eversion of the skin. Given the length of time it takes for reptile wounds to heal, sutures should not be removed until 6–8 wk after surgery.

The primary factors to consider postoperatively are analgesia and continued vigilance concerning hydration, temperature, nutrition, and hygiene. It is widely understood and accepted that reptiles can feel pain. Pain slows the healing process and depresses the normal function of the immune system in mammals. There is no evidence to suggest this process would not be similar in reptiles. Clinically, reptiles that receive postoperative analgesia appear to recover better than those that do not. The continuation of preemptive analgesia using opioids and/or NSAIDs should be a routine part of postoperative care.

Few drugs are approved by the FDA for use in reptiles. Medications can be given by a variety of routes, including PO, SC, IM, IV, intracardiac, intracoelomic, intraosseous, intrasynovial, or intratracheal injection. Certain drugs can be applied topically, given per cloaca, by inhalation (nebulization), or by direct intralesional administration. Because reptiles are ectotherms, temperatures outside the preferred optimal temperature zone (POTZ) can have profound influences on drug distribution, metabolism, excretion, and elimination half-life. Some therapeutic regimens state a fixed temperature at which the reptile should be held during treatment. If there are pharmacokinetic data on the drug, then the elimination of the drug will be known and constant. However, if this stated temperature is below or above the POTZ for the species being treated, then stress and debilitation may ensue. Even when the stated therapeutic temperature is within the POTZ for the species being treated, constant exposure to a fixed temperature is likely to be stressful.

Reptiles have a well-developed renal portal system; blood from the caudal half of the body passes through the kidneys before reaching the systemic venous circulation. Drugs injected into the caudal half of the body may have a significantly reduced half-life if excreted via tubular secretion. However, studies have demonstrated that these effects are unlikely to be clinically significant. Of potential concern is the caudal injection of nephrotoxic drugs that may reach renal tissue in high concentration. Also, the large bladder of chelonians may act as a drug reservoir and lead to a secondary therapeutic peak many hours after drug administration. The shell of tortoises, turtles, and terrapins is largely living tissue; therefore, all chelonian medication should be based on total body weight.

Drug Dosage and Allometric Scaling:

Numerous pharmacokinetic studies have been published for reptiles, and these should be considered the most reliable source for information on drug dosages. When species-specific information is not available, it is possible to extrapolate from closely related species. If there are no pharmacokinetic data or reliable clinical experience for a particular species, it may be necessary to extrapolate pharmacokinetic data from other animals. Allometric scaling calculates the drug dose and dosing frequency using metabolic rate rather than body weight. The basic allometric equations are shown below, in which W = body weight (kg) and K = energy constant, which is 10 for reptiles. These equations can be used to calculate a dose and dose frequency for a species for which no data are available, by using pharmacokinetic data from a known species (control), whether another reptile or mammal or bird:

Minimum energy cost = K (W0.75)

Specific minimum energy cost = K (W-0.25)

Antimicrobials:

Many bacterial infections in reptiles are caused by gram-negative bacteria, particularly Pseudomonas, Aeromonas, Citrobacter, Klebsiella, and Proteus spp. Bacterial resistance to many commonly used antibacterials is seen, and many gram-negative bacteria can have unexpected sensitivity to particular antibacterials; therefore, sampling for Gram stain, cytology, culture, and sensitivity testing should be done before starting therapy. Antibacterial therapy must usually be given while awaiting the results of bacterial sensitivity tests. In these circumstances, amikacin, ceftazidime, and enrofloxacin or ciprofloxacin are often preferred (see Table: Antimicrobial Drugs Used in Reptiles). In severe infections, amikacin may be combined with ampicillin or amoxicillin for respiratory tract infections or with ceftazidime for generalized or systemic infections. Chloramphenicol in combination with neomycin may be given for GI infections. Penicillin, metronidazole, lincomycin, or clindamycin can be used for anaerobic infections.

Antimicrobial Drugs Used in Reptiles

Drug

Dosage

Comments

Acyclovir

80 mg/kg/day, PO; topical cream 12 times daily

Antiviral

Amikacin

Corn snake: loading dose 1.7 mg/kg, IM, followed by 26 mcg/kg/hr via osmotic infusion-pump implant

Gopher snake: initial dose 5 mg/kg, IM, then 2.5 mg/kg, IM, every 3 days

Gopher tortoise: 5 mg/kg, IM, on alternate days

American alligator (juvenile): 2.25 mg/kg, IM, every 3–4 days

Ball python: 3.5 mg/kg, IM, every 4–5 days

50 mg/10 mL saline × 30 min nebulization bid

Maintain hydration

Amoxicillin

22 mg/kg, PO, bid

10 mg/kg/day, IM

Often ineffective unless given in combination with aminoglycosides

Amphotericin B

0.5–1 mg/kg, intracoelomic, IV, 1–3 days for 14–28 days

Aspergillosis; fluid therapy recommended

Tortoise: 0.1 mg/kg/day, intrapulmonary, for 28 days

5 mg/150 mL saline for 1 hr of nebulization, bid for 7 days

Pulmonary candidiasis

Ampicillin

Most species: 10–20 mg/kg, SC, IM, 1–2 times/day

Tortoises: 50 mg/kg, IM, bid

Azithromycin

Ball python: 10 mg/kg, PO, every 3–7 days (3 days for skin infections, 5 days for respiratory tract, 7 days for liver and kidneys)

Carbenicillin

200–400 mg/kg/day, IM

Cephalexin

20–40 mg/kg, PO, bid

Ceftazidime

20–40 mg/kg, SC, IM, IV, every 2–3 days

Ceftiofur

Tortoises: 2.2–4 mg/kg/day, IM

Snakes: 2.2 mg/kg, IM, on alternate days

Lizards: 5 mg/kg/day, IM, SC

Cefuroxime

100 mg/kg/day, IM, for 10 days at 30°C

Chloramphenicol

Indigo snake: 50 mg/kg, IM, bid

Midland water snakes: 50 mg/kg, IM, every 4 days

Most species: 20–40 mg/kg, IM, once to twice daily

Ciprofloxacin

10 mg/kg, PO, on alternate days

Clarithromycin

Desert tortoise: 15 mg/kg, PO, every 2–3 days

Mycoplasma

Clindamycin

5 mg/kg, PO, bid

Clotrimazole

Topical

Fungal dermatitis

Doxycycline

Most species: 5–10 mg/kg/day, PO

Hermann's tortoises: 50 mg/kg, IM, then 25 mg/kg, every 3 days

Enrofloxacin

Most species: 5-10 mg/kg/day, IM, PO

IM injection likely causes necrosis, and consideration should be given to single injection followed by oral administration

Nasal flush 50 mg/250 mL sterile water; 1–3 mL/nares daily to every other day

Burmese python (juvenile): 10 mg/kg, IM, initial dosage, then 5 mg/kg, IM, every 48 hr. Pseudomonas: 10 mg/kg, IM, every 48 hr

IM injection likely causes necrosis, and consideration should be given to single injection followed by oral administration

Hermann’s tortoise: 10 mg/kg/day, IM

IM injection likely causes necrosis, and consideration should be given to single injection followed by oral administration

Monitor lizards: 10 mg/kg, IM, every 5 days

Indian star tortoise: 5 mg/kg, IM, once to twice daily

IM injection likely causes necrosis, and consideration should be given to single injection followed by oral administration

Crocodilians: 5 mg/kg IV every 2–3 days

Fluconazole

Lizards: 5 mg/kg/day, PO

Sea turtles: 21 mg/kg, SC, once; then 10 mg/kg, SC, 5 days later

Gentamicin

American alligator: 1.75 mg/kg, IM, every 3–4 days at 22°C

Maintain hydration, nephrotoxicity reported

Painted turtle: 10 mg/kg, IM, on alternate days at 26°C

Maintain hydration, nephrotoxicity reported

Red-eared terrapin: 6 mg/kg, IM, every 25 days

Maintain hydration, nephrotoxicity reported

Gopher snake: 2.5 mg/kg, IM, every 3 days at 24°C

Maintain hydration, nephrotoxicity reported

Itraconazole

Chameleon: 5 mg/kg/day, PO

Fungal dermatitis

Spiny lizard: 23.5 mg/kg/day, PO, for 3 days, with persistent drug concentration for 6 days

Snakes: 10 mg/kg/day, PO

Sea turtles: 5 mg/kg/day, PO, or 15 mg/kg, PO, every 72 hr

Kanamycin

10 mg/kg/day, IM, at 24°C

Fluid therapy recommended

Ketoconazole

Crocodilians: 50 mg/kg/day, PO

Most species: 15–30 mg/kg/day, PO, for 14–28 days

Lincomycin

10 mg/kg/day, PO; 5 mg/kg, IM, once to twice daily

Marbofloxacin

Ball python: 10 mg/kg, PO, every 48 hr

Metronidazole

Bacterial infections, 20–50 mg/kg, PO, every 1–2 days

Maximal dose for tricolor snake, king snake, indigo snake, or Uracoan rattlesnake, is 40 mg/kg

Neomycin

10 mg/kg/day, PO

Oral only, not to be given systemically

Nystatin

Enteric fungal conditions in turtles: 100,000 U/kg/day, PO, for 10 days

Oxytetracycline

Most species: 5–10 mg/kg/day, IM, PO

Pain, irritation, and inflammation at injection site

American alligator: 10 mg/kg, IV, IM, every 4–5 days

Mycoplasmosis

Piperacillin

50–100 mg/kg, IM, every 12 days

Fluid therapy recommended

Polymixin B

Topical

Abrasions, wounds

Silver sulfadiazine

Topical every 24–72 hr

Broad spectrum for skin infections

Sulfamethoxydiazine

80 mg/kg, SC, IM, then 40 mg/kg/day, for 5–7 days

Coccidial infections

Tobramycin

Chelonians: 10 mg/kg, IM, every 1–2 days

Most species: 2.5 mg/kg, IM, every 1–3 days

Potentially nephrotoxic, fluid therapy recommended

Tylosin

5 mg/kg/day, IM

Mycoplasmosis

Voriconazole

10 mg/kg, PO

Fungal and yeast infections commonly occur in reptiles. GI and skin mycoses are particularly common in reptiles maintained on inappropriately longterm broad-spectrum antibacterials. Cutaneous mycoses can often be treated by debridement and topical application of antifungals. GI infections can be treated with nystatin, whereas systemic infections may require itraconazole or fluconazole. In cases of pulmonary mycoses, antifungal medication may be given by nebulization or intratracheal or intrapulmonary injection.

Herpesviruses can cause severe morbidity and mortality in chelonians. Acyclovir has been used with some success during the early stages.

Parasiticides:

Parasiticides used commonly in reptiles are listed in Parasiticides Used in Reptiles. Parasiticide overdosage may lead to drug toxicity, which may be seen as neurologic signs, including seizures. Ivermectin is contraindicated in chelonians, and adverse reactions have been reported in some iguanid lizards, skinks, and indigo snakes. Milbemycin has been successfully used in box tortoises and terrapins, but it is recommended that ivermectins and milbemycins be avoided in all chelonians because safer alternatives are available. Permethrin is licensed for use in reptiles and appears safe and effective against mites and ticks.

Parasiticides Used in Reptiles

Drug

Dosage

Parasite

Comments

Endoparasiticides

Fenbendazole

25–100 mg/kg, PO every 14 days for up to 4 treatments

50 mg/kg/day, PO, for 3–5 days

Roundworms, Hexamita

Can cause leukopenia

Ivermectin

200 mcg/kg, PO, IM, SC, repeat after 14 days

Not in chelonians; care in skinks and indigo snakes

Levamisole

5–10 mg/kg, SC, intracoelomic, repeat after 14 days

Lungworms and other nematodes

Snakes, lizards, care in tortoises (use 5 mg/kg)

Mebendazole

20–25 mg/kg, PO, repeat after 14 days

Strongyles and ascarids

Metronidazole

20–40 mg/kg, PO, every 1–2 days for 2–5 treatments

Protozoa

Oxfendazole

68 mg/kg, PO, as a single dose

Nematodes

Paromomycin

35–100 mg/kg/day, PO, for 28 days

Amoebas, cryptosporidia

Does not eliminate cryptosporidia

Ponazuril

Bearded dragons: 30 mg/kg, PO, every 2 days for two treatments

Coccidiosis

Praziquantel

8 mg/kg, PO, SC, IM, repeat after 14 days and 28 days

Tapeworms, flukes

Pyrantel

5 mg/kg, PO, repeat in 14 days

Nematodes

Spiramycin

160 mg/kg/day for 10 days, then twice weekly for 3 mo

Snakes with cryptosporidiosis

May reduce clinical signs but does not clear infection

Toltrazuril

5–15 mg/kg, PO, repeat in 14 days

15 mg/kg, every 48 hr for 10 days; discontinue for 2 wk; repeat every 48 hr for 10 days, and repeat as necessary

Bearded dragons, coccidiosis

Tortoises, intranuclear coccidiosis

Safety, efficacy, and pharmacokinetic data lacking

Trimethoprim-sulfa

30 mg/kg/day, PO, for 10–28 days

Coccidia

Ectoparasiticides

Dichlorvos-impregnated strip

1 cm2 of strip/30 cm3 vivarium for 28 days or 2.5 cm2 of strip/25 cm3 vivarium for 2–3 days every week

Toxic, vivarium should be emptied; keep out of direct contact of animals

Fipronil

By spraying, every 7–10 days

Mites and ticks

Ivermectin (10 mg/mL)

By spraying, 1–2 mL/L water every 7–10 days 200 mcg/kg, IM, every 7–14 days

Mites and ticks

Should not be used in chelonians; use care in skinks and indigo snakes

Permethrin (10%)

Topical spray

Mites and ticks

Licensed product available for reptiles in the USA

Other Medications:

Dosages for medications used for a variety of other disorders of reptiles are listed in Miscellaneous Drugs for Reptiles.

Miscellaneous Drugs for Reptiles

Drug

Dosage

Condition

Allopurinol

20–25 mg/kg/day, PO

Gout, reduces uric acid production

Aluminium hydroxide

100 mg/kg, PO, every 12–24 hr

Reduces blood phosphorus levels

Aminophylline

2–4 mg/kg, IM

Respiratory disease when bronchodilation required

Argipressin (vasopressin)

0.01–1 mcg/kg

Egg binding (more potent than oxytocin)

Ascorbic acid

10–200 mg/kg, IM, as required

Ulcerative stomatitis

Calcitonin

1.5 units/kg, SC, tid

50 units/kg, IM, repeat after 2 wk

Hypercalcemia, fluid therapy also recommended

Secondary hyperparathyroidism

Calcium gluconate (10 mg/mL)

100 mg/kg, IM, qid, or 400 mg/kg, IV, intraosseous, given over 24 hr

Hypocalcemia in iguanas; high phosphorus concentration may cause soft-tissue mineralization

Cimetidine

4 mg/kg, PO, tid-qid

Regurgitation, vomiting, gastritis, GI ulceration

Cisapride

0.5–2 mg/kg/day, PO

GI motility modification; not recommended to use with clarithromycin in tortoises

Cholecalciferol

100–1,000 units/kg, IM, as a single dose

Hypocalcemia, fibrous osteodystrophy in iguanas

Cyanocobalamin

50 mcg/kg, SC, IM

Appetite stimulation

Dexamethasone

30–150 mcg/kg, IM, IV, intraosseous

Inflammation, shock, beware of immunosuppression

Dinoprost (prostaglandin)

500 mcg/kg, IM, as a single dose

Egg binding in snakes

Doxapram

5–10 mg/kg, IV, intraosseous

Respiratory stimulation

Flunixin

100–500 mcg/kg, IM, IV, once to twice daily

Inflammation, pain

Furosemide

2–5 mg/kg, IM, IV, once to twice daily

Diuresis (effective despite lack of loop of Henle in reptiles)

Iodine

2–4 mg/kg, PO, every 7 days

Prophylaxis for goitrogenic diets

Iron

12 mg/kg, IM, every 7 days (alligators)

Anemia in alligators

Levothyroxine

20 mcg/kg, PO, on alternate days

Hypothyroidism in tortoises

Metoclopramide

60 mcg/kg/day, PO, for 7 days

Stimulation of gastric emptying in tortoises

Prednisolone

1–2 mg/kg, PO

Anti-inflammatory, reduction of nephrocalcinosis, beware of immunosuppression

Selenium

25–500 mcg/kg, IM

Deficiency in lizards

Sucralfate

0.5–1 g/kg, PO, tid-qid

Gastric irritation/ulceration

Thiamine

50–100 mg/kg, IM

Thiamine deficiency

Vitamin A

5,000 units/kg, PO, every 7 days

Hypovitaminosis A (iatrogenic hypervitaminosis A may result from repeated treatment)

Fluid Therapy:

Dehydration in reptiles is usually associated with prolonged anorexia and water loss, deprivation, or an inability to drink, rather than mixed electrolyte losses through frequent vomiting or diarrhea. Water balance in reptiles differs from that in mammals, because, per unit body weight, reptiles have a higher percentage of total body water (63.0%–74.4%) and a higher percentage of intracellular water (45.8%–58.0%). These values appear to be highest in freshwater species, lower in terrestrial reptiles, and lowest in marine reptiles, with the concentration of isotonic saline in nonmarine reptiles being 0.8%. This has led to the conclusion that normal 0.9% saline may be too concentrated for most reptiles. Balanced crystalloid fluids containing dextrose (260–290 mOsm/L) appear to be effective. As a general guide, maintenance requirements are about 5–15 mL/kg/day, and rehydration should proceed at 35–40 mL/kg/day, although in cases of shock, rates of 3–5 mL/kg/hr can be used for several hours.

In many instances, simply permitting a reptile to bathe in shallow, warm water within a vivarium maintained at the species-specific preferred optimal temperature zone will promote drinking. Such a method is acceptable when reptiles are able and willing to drink voluntarily. However, in many cases oral fluids must be delivered via a stomach tube. For oral rehydration, mammalian electrolyte solutions can be used but are best diluted by a further 10%–15% to produce a slightly hypotonic solution. Significant amounts of water may also be absorbed via the cloaca when chelonians (and possibly other species) bathe. Oral fluid therapy works well to provide maintenance requirements, rehydrate mildly dehydrated reptiles, and as a vehicle to give oral medications and food. For alert and active reptiles, this method is preferred because it facilitates GI activity, and fluids are rapidly absorbed when the reptile is maintained at correct temperatures. Repeated stomach tubing is stressful and can be difficult in strong chelonians; therefore, esophagostomy tubes are recommended for longterm oral therapy. Small fluid volumes can be given SC, but in moderate to severe cases, intracoelomic, IV, or intraosseous routes are favored.

Intracoelomic administration of fluids is faster, less stressful, and allows delivery of a larger volume than stomach tubing. Large volumes of intracoelomic fluid could compromise lung function and may be slowly absorbed; however, IV catheterization is not easy, and cut-down procedures are often required. In emergencies with larger snakes, it is possible to place an intracardiac catheter. Fluid infusion is best controlled by a syringe driver or infusion pump. If such devices are not available, the total daily fluid volume can be divided into three or four bolus injections, each given slowly over 10–20 min. IV catheters can usually be left in place for up to 72 hr; intracardiac catheters up to 24 hr. Intraosseous infusion is an easier technique in lizards, small crocodilians, and chelonians. The needle is directed into the medullary cavity of a long bone, and aspiration of marrow or radiography can verify correct positioning. Great care must be exercised when dealing with osteodystrophic lizards to avoid limb fractures. Infusion rates for IV and intraosseous administration are similar. As a general guide, 0.8–1.2 mL/kg/hr is suitable for rehydration purposes, but in cases of severe dehydration or shock or during surgery, 3–5 mL/kg/hr can be given for 2–3 hr.

Colloids are less frequently used in reptiles because much of the water loss is from the intracellular space rather than plasma, but they can be used in cases of acute hemorrhage. If severe hemorrhage occurs (ie, PCV <5%), whole blood may be given by IV and intraosseous routes. Cross-matching does not appear necessary, at least for a single transfusion. Ideally the donor and recipient are of the same species.

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