ABSTRACT: Monitoring anaesthesia in reptiles can be daunting for the RVN. but with increasing numbers of reptiles being seen and treated in practice it is more important than ever for the RVN to understand how to monitor anaesthesia safely in these patients. The usual monitoring tools can be used in reptile anaesthesia, but there are some differences when compared with their use in mammals. Such differences are highlighted here, alongside useful practical information of what to expect during the induction, maintenance and recovery phases of anaesthesia, and how best to monitor an anaesthetised reptile throughoul each stage

The principles of general anaesthesia apply as much to exotics as they do to dogs and cats – requirements for oxygen delivery and elimination of waste carbon dioxide must be maintained via correct implementation and monitoring of tidal volumes, respiratory rates and body temperature.

Anaesthetic nursing of reptiles does, however, differ considerably from that of mammals and birds; but only because of their unique anatomy and physiology. Reptiles present a host of problems when it comes to anaesthesia and anaesthetic nursing. These problems can be overcome and, with a little background knowledge and preparation, reptiles can be nursed safely through an anaesthetic procedure that utilises suitable anaesthetic agents and monitoring equipment.

Preparation for anaesthesia

Many reptiles that are presented for anaesthesia will have subclinical or overt disease and this should be considered when monitoring any anaesthetic. Wherever possible, any disease should be diagnosed and treated beforehand – so at least the patient should be rehydrated and offered nutritional support for a period of days (where appropriate), and patients should always be at their preferred body temperature at the time of anaesthetic induction.

An anaesthetic chart should be used for every anaesthetic, with a note made of induction, intubation, surgery start, finish and extubation times. Preparing the anaesthetic chart in advance allows monitoring (and recording) to commence immediately upon induction of anaesthesia.

The nurse should always have an idea of the patient's resting heart and respiratory rates, as well as physiological characteristics (such as colour and demeanour) prior to anaesthetic induction. It is also helpful to have a separate area on the anaesthetic chart to record these parameters before the procedure for comparison throughout the anaesthetic.

Monitoring equipment

Anaesthetic agents produce depression of the cardiopulmonary system; therefore, during anaesthesia, cardiopulmonary performance must be closely monitored by means of the use of visual and mechanical monitoring techniques. Visualisation and manual counting of the respiratory and heart rates is possible in most reptiles, although it may not be easy, and subtle changes are certainly harder to ascertain. So it is recommended that specific monitoring equipment is utilised (if available) to provide a more accurate picture of cardio-respiratory function.

Pulse oximetry

Although pulse oximeters are regularly used in practice to detect arterial haemoglobin oxygen concentrations in mammals, they are unreliable in reptiles as the characteristics of such parameters vary markedly and so results should be interpreted with caution. If a pulse oximeter is to be used, more accurate readings may be obtained with a reflectance probe rather than a transflectance probe, which should be taped in place. 


Capnography allows for monitoring of respiratory rate and end tidal carbon dioxide (ETCO,). It offers an idea of how effective respiration is at the time, although its usefulness is limited by the fact that reptiles can develop cardiac shunts. Changes in ETC02 may give valuable information on existing complications, however, and an increase in levels may highlight airway leaks or obstruction, disconnection from the breathing system or ventilator malfunction.

Capnography is most useful and accurate when the patient is intubated, and a machine designed specifically for use in small or light patients – with a low tidal volume and a side-stream sampling rate of 50 ml per minute – is usually required.

Specific, narrow-bore endotracheal tube (ET) connectors reduce dead space and are recommended for use in small reptiles. A capnograph of this type also allows for the monitoring of ETC02 during the recovery phase, whilst the animal is wearing a face mask or directly following extubation.

When monitoring ETCO, in anaesthetised reptiles, levels should remain between 3 – 5% to ensure that over- or under-ventilation (respectively) do not occur. If the patient ‘breath-holds’, manual intermittent positive-pressure ventilation (IPPV) with a ]ackson-Rees modified Ayres T-piece, is recommended; or, alternatively, mechanical ventilation should be instigated. Using a capnograph alongside ventilation allows the nurse to monitor how effective it is and to adjust its management accordingly.

Blood gas analysis

Although available in most practice situations now, blood gas analysis is still a relatively expensive way to monitor an anaesthetised reptile and results are subject to misinterpretation. Because reptiles tend to be more tolerant of changes in pH, PCO, and P02, normal values (calibrated for mammals) may not be applicable.

Obtaining an arterial blood sample can also prove difficult – or impossible – in many species, making this monitoring tool relatively impractical.


It is difficult to hear the heart via auscultation with a stethoscope in reptiles, owing to the ‘interference’ from scales or the plastron and carapace of chelonia, so a Doppler probe is recommended for accurate and constant monitoring. Used with conduction gel, the Doppler should be secured in place with an adhesive bandage over a large blood vessel or the heart: tortoises – place the probe between the neck and the fore limb, close to the heart (located near the thoracic inlet) and major blood vessels lizards – the heart is usually positioned in the anterior thorax, so the probe can be placed on the left-hand side of the thorax, between the fore legs, along the midline over the ventral abdominal vein or on the tail over the caudal tail vein – along the ventral midline of the tail approximately one third of the distance from cloaca to tail tip (Figure 1) snakes – the Doppler can be held directly over the heart, which is located as the first solid palpable mass in the cranial third of the body as you work caudally from the head. The heart is relatively mobile, so marking the position –   once found – with tape, can be useful.

Figure 1: Doppler probe in place on the left thorax of a Bearded Dragon limage courtesy Marie Kubiak)

Doppler can also be used to monitor systolic blood pressure. However,
because of the lack of suitable equipment and scientific information relating to blood pressure in different reptiles, it is rarely used for this purpose.

Oesophageal stethoscopes

Inexpensive and easy to use, an oesophageal stethoscope can be a very useful piece of equipment when monitoring anaesthesia in reptiles. The author advocates always having one in place (in patients that are large enough) as a back-up measure in case of movement or malfunction of other equipment.

Advancement of the oesophageal stethoscope probe should be performed whilst listening until maximum sound intensity is reached.

Electrocardiogram (ECG)

An electrocardiogram can be used to monitor heart rate and rhythm. However, electrical activity of the heart can continue in reptiles despite a loss of muscular activity. It is best, therefore, not to rely totally on ECG as a means ot monitoring an anesthetised reptile.

Placement of ECG leads can also be problematic in reptilian patients, particularly chelonia and some lizards; hence it may be necessary to place needles through the skin at the point of attachment, with the alligator clips already attached to the needles.


Reptiles are exothermic – they derive almost all of their body heat from their environment. Keeping the reptile within its preferred body temperature range during anaesthesia, by means of circulating warm air or water, heat pads used with care or forced blowers (a hair dryer will do) will improve metabolic function and homeostasis (Figure 2).

Figure 2: Bearded Dragon on a protected heat pad with a plastic drape used to retain heat during surgery I image courtesy Marie Kubiakl

This, in turn, will decrease risks associated with anaesthesia outside the normal parameters. Hypothermia is a major factor associated with anaesthesia- related complications in reptiles, so maintenance of appropriate temperature will speed up/improve recovery.

Conversely the patient should never be overheated, as temperatures above 42°C usually prove fatal.  Rectal or oesophageal thermometers can both be used to monitor temperature in anaesthetised reptiles; although the latter is deemed to be more accurate. To avoid the risk of trauma to the delicate rectal mucosa, a thermometer should be secured to the tail with adhesive bandage –   but only once the patient is in position. Moving a patient with a secured thermometer increases the likelihood of rectal mucosal damage.


Induction phase

The patient should have been provided with appropriate pre-operative analgesia and should be at the preferred optimal body temperature. Where possible, the patient should be hydrated and provided with fluid therapy throughout any procedure.

Successful induction of anaesthesia will create a loss of the righting and palpebral reflexes. In snakes, the tongue withdrawal reflex is lost. There should be a reduced or absent response to pain.

The equipment described earlier should be deployed, when possible, as soon after induction as is feasible. Monitoring can then begin and the nurse can assess stability of anaesthesia before any procedures are performed (Figure 3).

Figure 3: Crested Gecko undergoing gaseous induction of anaesthesia via face mask, with monitoring equipment prepared and visible in the background (image courtesy Mane Kubiakl

Monitoring during maintenance

Achieving a surgical plane of anaesthesia will ensure that there is complete muscle relaxation and cessation of spontaneous voluntary movements.

Palpebral and corneal reflexes and sphincter and jaw tone should gradually become reduced; although a total loss of the corneal reflex indicates that anaesthesia is too deep (a cotton swab lubricated with ocular lubricant can be gently touched onto the cornea to promote a response). It is important to note that palpebral and corneal reflexes cannot be effectively monitored in reptiles – such as snakes and some lizards –   that have spectacles’.

In snakes, a total loss of the tongue withdrawal reflex indicates that the depth of anaesthesia is too great. In chelonia the tail, limb and head withdrawal reflexes should all be absent during a surgical plane of anaesthesia.

It is recommended that a small animal ventilator is used – or ready for use – to mechanically ventilate any reptile undergoing anaesthesia (Figure 4).

Figure 4: Anaesthetised Boa Constrictor on a ventilator with Doppler probe in place [image courtesy Marie Kubiak)

Breath-holding is common, and respiratory depression is a side effect of most anaesthetics. By manually or mechanically ventilating all patients that breath-hold or show increased levels of ETC02, optimal respiratory function is maintained throughout anaesthesia. Using a mechanical ventilator allows for better monitoring and recording of anaesthesia, as the nurse is not having to ‘bag’ the patient every few seconds.

When ventilating reptiles manually, the pop-off valve should never be fully closed, in order to avoid excessive pulmonary pressure and potential pulmonary rupture.

Completion and recovery

Reptiles may have a prolon
ged recovery phase when compared with mammals and birds. The recovery phase can be shortened by:

•  ensuring optimal temperature is maintained throughout the procedure reducing the depth of anaesthesia towards the end of the procedure – do not lighten anaesthesia whilst painful procedures are still underway, for example, during the placing of staples or sutures 

•  ventilating the patient with room air –   not 100% oxygen unless hypoxia is known or suspected – at the end of the anaesthetic procedure using an Ambu bag (or equivalent) at a rate of 1-4 breaths per minute.

Endotracheal tubes should be left in place until the reptile is making regular, spontaneous, adequate respiratory movements.

Small animal incubators or a pre-prepared ‘recovery’ vivarium are ideal ways of ensuring that the temperature and humidity of the patient’s immediate environment are controlled. Monitoring should continue (even if remotely) until the patient is awake and mobile. 


Emma Whitlock RVN BSc (Hons) VNS

Emma Whitlock RVN completed a BSc[Hons] degree in Veterinary Nursing Science in 2004 before working with wildlife, zoo and exotic species in various nursing roles. A stint in academia led to her current role as senior avian and exotic nurse at Great Western Exotics, heading up the exotic nursing team at the multidisciplinary Vets Now Referrals in Swindon.

To cite this article use either

DOI: 10.1111/j.2045-0648.2012.00181.x or Veterinary Nursing Journal Vol 27 pp 215-219

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Veterinary Nursing Journal • VOL 27 • June 2012 •