Reptile Anaesthesia

Reptile anaesthesia is very different to that of cats and dogs and many physiological systems are temperature dependant which is why they must be kept at their preffered optimal body temperature (POBT) during the perianesthetic period. Anaesthetic agents given parenterally are metabolised or transformed and then eliminated from the body through complex systems. Temperature will have a direct effect on enzyme activity and therefore elimination will be slowed with suboptimal temperatures. Inhalatory anaesthetic potency is also affected by temperature. Times of onset and elimination are slower than in mammals as circulation speed and respiratory efficiency are reduced in the reptile patient. An additional complication is that reptiles are able to shunt pulmonary blood which in turn also has a direct effect on onset and elimination times.

PRE-ANAESTHETIC CONSIDERATIONS

Any abnormality should be corrected if possible before anaesthesia induction and correct stabilisation should always be attempted. Food and water intake should be monitored and registered. Fasting is not a concern in reptile anaesthesia as long as there is no food in the esophagus, nor live food prey in the stomach. To ensure digestion and prevent any regurgitation (ie in snakes) a fasting period of 18h may be applied in chelonians and lizards or 72-96h in larger lizards or snakes.

POBT should be considered through the entire peri-anaesthetic period and the use of a esophageal or cloacal temperature probe may be useful. Even more care should be taken to ensure correct temperatures if injectable agents such as ketamine are used.

Premediaction is not considered necessary in reptiles although many reptile vets will use anti-inflammatory drugs before surgery and before any inflammation begins which certainly does seem to make sense. As similar regime may be used for opioids.

ANAESTHESIA

A similar approach is used as in mammals which means that anaesthesia should deliver analgesia, amnesia and immobilisation with the use of drug combinations delivered via the parenteral or inhalatory route.

Reptiles are extremely resistant to hipoxic situations, with aquatic turtles having survived in 100% nitrogen for over 6 hours (Wasser et al., 2001). They may switch to anaerobic metabolism if requiered and this may include using inhalatory gases where they readily breath hold for long periods of time if inhalatory induction is attempted. Snakes and lizards, on the other hand, may be induced with inhalatory anaesthetics deliverd by oxygen with the use of face masks or induction chambers.

Induction and Intubation

Long periods of apnoea may be present during inhalatory induction and therefore extreme care should be taken and will be dependant on the reptile group in question. Direct intubation in snakes, especially larger individuals is generally quick and induction is performed within minutes. Lizards may also be induced via facemask or induction chamber, especially smaller species. Injectable agents such as propofol or alfaxalone is preferred in larger lizards or tortoises, delivered via the ventral tail vein or dorsal tail vein (dorsal coccygeal vein), respectively.

Heart rate (HR) via Doppler, respiratory rate (RR), righting reflex and pinch reflex should be monitored during induction.

Intubation is certainly a good idea in all reptiles in the same way that it is in humans or small animals as it is generally an easy procedure to perform, will deliver oxygen (and an anesthetic agent if required) and is a precaution for if problems arise during anaesthesia. Different endotracheal tubes (ET tubes) can be used depending on the size of the individual; these may be modified intravenous (IV) or urinary catheters or even small animal (cat and dog) ET tubes for larger species. Remember that the glottis opens on inspiration and therefore we may have to wait as respiratory rate is reduced compared to mammals. Use of topical anaesthetics for intubation are innecessary.

Tracheal rings may be complete or incomplete and therefore, similar to avian anaesthesia, ET tubes should not be cuffed as a general rule. Tracheal bifurcation position varies with species and in tortoises a proximal bifurcation ocurrs just behind the mandible; this should be considered to prevent bronchial intubation. Tortoises have a fleshy tongue that takes up a major part of the oral cavity and therefore extending the head whilst applying submandibular pressure should expose the glottis. Marine turtles have pharyngeal spines. Intubation of lizards is straight forward although may prove challenging in smaller species and the glottis is positioned just behind the tongue. A cotton bud as a tongue depressor works well at the same time that pressure is applied in the submandibular region to elevate the glottis to improve visualisation. In snakes the glottis is situated above the lingual recessin the anterior region of the oral cavity. A conscious intubation technique may be performed, especially in larger species. In venemous species a intramuscular (IM) dose of ketamine (2-5mg/kg) is recommended. This will increase recovery time. Crocodilians posess a basihyoid valve that dificultates visualisation of the glottis.

A ventilator may be considered an essential tool to deliver intermittent positive pressure ventilation (IPPV) as apnoeas are frequent in reptiles. Although generally speaking they are very resistant to hipoxia in comparison to mammals or birds, we are able to modify oxygen partial pressure by increasing depth or frequency. A redily available ventilator is the Vetronics ventilator (SAV03) as it works well even with smaller patients ie 30g geckos.

Parenteral Anaesthesia

The advantages of this form of anaesthetic delivery are: easy administration, readliy available drugs and reduced necessity of specialised equipment. Disadvantages include the need for an exact body weight, possibility of overdosing and inability of reversal, challenging intravenous delivery, longer recovery periods depending on the agents used.

Ketamine is readliy available but has a disociative action affecting recovery times, especially with repeated dosing. Induction takes 10-20min and recovery may extend to 24-96h. Injection is painful and it is not recommended in debilitated animals where a prolonged recovery may prove detrimental. Low doses may be used in chelonia for oesophageal tube (O-Tube) placement or jugular blood sampling. A 3-4mg/kg IM dose has been suggested as efficacious in tortoises and snakes. In smaller snakes a subcutaneous (SC) injection may be administered.

Medetomidine/Ketamine combination can be used, especially to reduce recovery times with the use of atipamezole. Greer et al. (2001) suggested a dose 5 times that of medetomidine to enable complete recovery within 60min.

Propofol is a non-barbituric hypnotic sedative that enables quick induction and recovery. Administration may be IV or intraosseus (IO). Doses varie between 5-10mg/kg and 1-2mg/kg for large chelonia. Care must be taken not to administer to dose-effect as induction time is more prolonged than in dogs and cats. Metabolism is fast and it is not accumulative. cardiopulmonar depression and apnoea may be seen. An example dose: 10mg/kg slow IV administration in a Green Iguana will deliver 20min of light anesthesia and spontaneous breathing.

Alfaxalone

Muscle relaxants will not deliver analgesia and are not generally used in routine reptile anesthesia. A combination of midazolam and ketamine may be useful for procedures such as taking radiographs, clinical examination or ET tube placement.

Barbiturics are normally used for euthanasia and not for surgical anesthesia. They have been associated with elevated death rates in skinks.

Inhalatory Anaesthesia

Advantages include easier anaesthetic depth control, 100% oxygen delivery, possibility of IPPV, faster recovery times and weight measurement not as important (ideal for venemous or extremely heavy animals). Disadvantages include the need of specialised equipment and costs.

Isofluorane is the most commonly used gas.

Intermitent positive pressure ventilation (IPPV)

In reptiles it is very convenient and safer to use pressure ventilators rather than volume ventilators. If we think of a small animal for which we have calculated a tidal volume of 2ml and we deliver 3ml, that is 50% excess. A ventilator that works at low pressures is recommended.

Generally pressures are maintained between 5-12cm of water. Patients must be intubated as described previously. Smaller patients may benefit fro the use of a IV catheter as they have a Luer conection that fits on to a 2.3mm ET tube adaptor. Dead space will be reduced and if a port is provided CO2 may be monitored through capnography.

The Vetronic Small Animal Ventilator (SAV03) may ventilate animals up to 10kg with pressures between 1-20cm of H2O. It is an easy to use ventilator, readily available and a common piece of equipment in exotic animal practices.

IPPV Steps

1) Weigh the reptile

2) Volume per minute = Breaths (per minute) x Tidal Volume (10ml/kg)

3) Gas flow = 3 x Volume (per minute)*

*Inspiration/Espiration ratio is 1:2 which means that we have 1/3 of the time to deliver air/gas equivalent to 3 times the Volume per minute.

4) Pressure 3-4mm through Minilak**

** Dead space is an important consideration and low dead space circuits have been developed - photos to follow.

Isofluorane is the preferred inhalatory gas as the vast majority is eliminated through the lungs. Sevofluorane is now also being used in veterinary practices. There have been reports of certain reptile species being maintained with difficulty and therefore it may be a good idea to have access to both inhalatory anaesthetics. Induction is generally 4-5% at 1l/min O2 for 5-20min, maintenance 1-4% and recovery is generally 10-30min.

Respiratory rate needed to maintain anesthesia frequently is higher than physiological respiratory rate and this is the reason that a ventilator is essential to maintain anesthesia in reptiles. Remember that espiration is active in a conscious reptile and therefore pressure should be applied to empty the lungs every so often.

Anaesthestic Monitoring

Muscular relaxation starts from cranial and extends caudally with recovery the other way around - this is important in venemous species.

Anaesthetic planes are light (L), surgical (Sx) and too deep (DP)

Anaesthetic depth is measured through numerous reflexes:

Digit Pinch Abolished: Sx/DP  
Tail Pinch Abolished: Sx/DP  
Head retraction Abolished: Sx/DP Upon pinching digit. Specific to chelonia
Palpebral Abolished: L/Sx/DP  
Cloacal Tone Abolished: DP  
Righting reflex Abolished: Sx/DP Useful in snakes
Lingual reflex Abolished: DP Useful in snakes
Corneal reflex Abolished: DP  
Bauchstreich response Abolished: Sx/DP Ventral stroking in snakes from cranial to caudal
Mandibular tone Abolished: Sx/DP  

 

Remember that the glottis will be closed in a live snake apart from when taking breaths. If it is constantly open then it is most likely dead.

Useful equipment: ECG, Doppler, Thermometer (with rectal probe). Capnography is of little use in reptile anaesthesia (contrary to avian anaesthesia)

RECOVERY

Wheras in mammals increase of CO2 partial pressure (PCO2) stimulates spontaneous ventilation, in reptiles this is dependant on partial pressure of oxygen (PO2) and therefore recovery time is reduced by using an ambibag or by disconnecting 100% oxygen delivery after an anaesthesia maintained with IPPV. Stimulation of spontaneous ventilation and accelerated recovery after IPPV may be achieved by reducing partial pressure of oxygen (Diethelm, 2001).

Preffered optimal body temperature (POBT) should be maintained and hydration should be considered by using intracoelomic (IC) or intraperitoneal (IP) fluids. It is frequent for reptiles to become acive suddenly and then appear to become anaesthetised again and so care should be taken that the patient does not end up at a coler part of the hospital vivaria. Analgesia should be monitored closely and the use of non steroidal anti-inflammatories or opioids are readlily avaliable and should be considered.


REFERENCES AND FURTHER READING

BSAVA, Manual of Reptiles, pag 134

Bennet RA (1998) Pain and analgesia in reptiles and amphibians. Proceedings of the Association of Reptilian and Amphibian Veterinarians 1998, pp.29-31

Heard DJ (2001) Reptile Anaesthesia. Veterinary Clinics of North America: Exotic Animal Practice 4, 83-117

Bennett, R. A. 1991. A review of anesthesia and chemical restraint in reptiles. J. Zoo Wildl. Med. 22: 282– 303.

Bennett, R. A. 1996. Anesthesia. In: Mader, D. R. (ed.). Reptile Medicine and Surgery. W. B. Saunders Co., Philadelphia, Pennsylvania. Pp. 241–247

Diethelm G (2001) The effect of oxygen content of inspiratory air (FIO2) on recovery times in the Green Iguana (Iguana iguana), Doctoral thesis, Germany, Universitaet Zuerich

Heard, D. J. 1993. Principles and techniques of an- esthesia and analgesia for exotic practice. Vet. Clin. N. Am. Small Anim. Pract. 4: 1301–1327.

Heard, D. J. 2001. Reptile anesthesia. Vet. Clin. North Am. Small Anim. Pract. 4: 83–117.

Page, C. D. 1993. Current reptilian anesthesia pro- cedures. In: Fowler, M. E. (ed.). Zoo and Wild Animal Medicine Current Therapy 3. W. B. Saunders Co., Phila- delphia, Pennsylvania. Pp. 140–143.

Perry, S. F. 1998. Lungs: comparative anatomy, functional morphology, and evolution. In: Gans, C., and A. S. Gaunt (eds.). Biology of the Reptilia. Vol. 19, Mor- phology G Visceral Organs. Society for the Study of Am- phibians and Reptiles, St. Louis, Missouri. Pp. 1–92.

Schumacher, J. 1996. Reptiles and amphibians. In: Thurmon, J. C., Tranquilli, and G. J. Benson (eds.). Lumb and Jones’ Veterinary Anesthesia, 3rd ed. Williams & Wilkins, Baltimore, Maryland. Pp. 670–685.

Schumacher, J. 2003. Reptile respiratory medicine. Vet. Clin. N. Am. Exotic Anim. Pract. 6: 213–231.

Stoelting, R. K., and R. D. Miller. 2000. Basics of Anesthesia. Churchill Livingstone, Indianapolis, Indiana.

Wang, T., A. W. Smits, and W. W. Burggren. 1998. Pulmonary function in reptiles. In: Gans, C., and A. S. Gaunt (eds.). Biology of the Reptilia. Vol. 19, Morphol- ogy G Visceral Organs. Society for the Study of Amphib- ians and Reptiles, St. Louis, Missouri. Pp. 297–374.

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