A full pre-anaesthetic examination should be performed in any animal undergoing anaesthesia. Any patient undergoing an elective procedure should be in good health with no obvious external abnormalities such as poor haircoat, ocular, aural or nasal discharge or abnormal ambulation. Auscultation of the heart and lungs should be performed and can be facilitated by the use of neonatal or paediatric stethoscopes in smaller patients. Normal physiological values can be found in Table 1. Rectal temperatures can be taken in larger patients and ‘well-behaved’ smaller patients. In non-elective procedures a thorough pre-anaesthetic examination can help identify comorbidities that may affect the anaes-thesia, and these can be planned for appropriately. During the physical examination it is essential that a bodyweight is obtained for the patient in order to calculate anaesthetic drugs as necessary as well as calculate cardiopulmonary resuscitation drugs prior to induction of anaesthesia.
Table 1. Normal resting biological values for common small mammal species
| Species | Normal HR | Normal RR | Normal rectal temp |
|---|---|---|---|
| Rabbits | 130–325 beats per minute | 32–60 breaths per minute | 37.8–39.4°C |
| Ferrets | 180–250 beats per minute | 33–36 breaths per minute | 37.7–40.0°C |
| Guinea Pigs | 226–300 beats per minute | 69–150 breaths per minute | 38.0–39.0°C |
| Chinchillas | ~100 beats per minute | 40–100 breaths per minute | 38.0–39.0°C |
| Rats | 250-450 beats per minute | 70–115 breaths per minute | 37.2–38.3°C |
| Mice | 325–780 beats per minute | 40–80 breaths per minute | 36.5–38.0°C |
| Degus | 100–150 beats per minute | ~75 breaths per minute | 37.9–38.3°C |
| Hamsters (all species) | 310–471 beats per minute | 38–110 breaths per minute | 37.0–38.0°C |
| Gerbils | 260–600 beats per minute | 85–160 breaths per minute | 38.0–39.0°C |
| African Pygmy Hedgehogs | 180–280 beats per minute | 25–50 breaths per minute | 36.1–37.2°C |
| Sugar Gliders | 200–300 beats per minute | 16–40 breaths per minute | 35.8–36.6°C (cloacal temperature) |
Compiled from Brust and Pye (2013) Keeble (2011), Tully (2009), Wolf (2009), Vennen and Mitchel (2009), Heatley and Harris (2009) and Heatley (2009)
Most pet mammal species are prey animals and so the smell, sight and sound of predators can cause significant stress. Stress can result in stimulation of the adrenergic system which could result in tachycardia, hypertension, reduced renal perfusion and reduced gut perfusion (Longley, 2008a). Small mammal patients should be housed in appropriate kennels or cages for their size in a quiet, darkened location away from loud noises and predators to help reduce stress.
Small mammal patients can be assessed for a modified American Society of Anaesthesiologists (ASA) grade, similar to that in dogs and cats. The ASA grade in veterinary medicine is assigned out of five, with each grade described in Table 2. ASA grades 1–2 are generally considered healthy and ASA grades ≥3 are generally considered sick (Portier and Ida, 2018). This classification system can give a good idea of prognosis and allows for the staff performing the anaesthesia and procedure to discuss comorbidities and extra precautions should they need to be taken.
Table 2. American Society of Anesthesiologists physical status classification system (2019)
| ASA Grade | Definition |
|---|---|
| ASA I | Normal, healthy animal, no underlying disease (elective procedure) |
| ASA II | Slight risk, minor disease present (healthy patient that needs a procedure) |
| ASA III | Moderate risk, obvious disease present (moderate systemic disease) |
| ASA IV | High risk, significantly compromised by disease (pre-existing disease or severe disturbances) |
| ASA V | Extreme risk, moribund (life-threatening systemic disease) |
Fasting is often not required or recommended for small exotic mammals. Fasting is, however, recommended for ferrets as they will readily vomit on induction. Fasting times of 3–4 hours are recommended to allow gastric clearance prior to induction, however any longer than this can predispose to severe hypoglycaemia (Matchett et al, 2012). Fasting in rabbits and small herbivores may cause gastrointestinal ileus and does not significantly reduce their gastric volume (Girling, 2013). Small rodents cannot vomit, and so fasting them is unnecessary and can cause them to develop hypoglycaemia (Allweier, 2016).
Induction
Premedication can be a useful tool in small mammal medicine. The use of a premedicant can sedate the patient, reducing stress on induction, and some drugs may have an anaesthetic-sparing effect, allowing for lower concentrations of volatile anaesthesia during the proposed procedure (Longley, 2008b). Opioid only or opioid combination premedication can provide analgesia before the procedure even begins, resulting in patients that are well-analgised throughout the procedure and into the recovery.
Induction can be performed with injectable agents or via inhalational anaesthesia. In smaller species where intravenous access is not readily available, inhalational anaesthesia is usually the preferred mechanism of induction. Sevoflurane is the preferred inhalational agent as it is less irritating to mucous membranes and respiratory passages compared with isoflurane (Thompson and Bament, 2012) and reduces incidences of breath holding in rabbits during induction at levels of 4% or lower (Girling, 2013). Guinea pigs often hypersalivate when a gaseous anaesthesia is introduced which can result in aspiration of saliva if it is not suitably removed; however hypersalivation is lessened with sevoflurane compared with isoflurane (Longley, 2009), and can be further reduced with the administration of glycopyrrolate or atropine with the premedication (Girling, 2013).
Administration of inhalational anaesthesia can be performed in an induction chamber (Figure 1) or via a face mask (Figure 2). Induction chambers are often made from modified Tupperware or plastic food containers that can provide an air-tight seal on the lid of the box and around the anaesthetic ports. The induction chamber should be appropriately sized for the patient, to allow for less gas waste and shorter induction times. During induction the gas flow rate can be calculated as f=V/t ln (S/S-C) where ‘f’ is the oxygen flow rate, ‘V’ is the chamber volume, ‘t’ is the time to reach the desired concentration of induction agent, ‘S’ is the concentration that the vaporiser is set to and ‘C’ is the desired concentration of inhalational agent (Hawkins and Pascoe, 2012). Induction chambers can also be useful to pre-oxygenate a patient prior to induction, which can be useful in cases with cardiovascular compromise or respiratory disease (Longley, 2008b). Pre-oxygenation benefits are often achieved in less than a minute in healthy animals (Hawkins and Pascoe, 2012).
While induction is being performed in an induction chamber most monitoring is relatively hands-off, due to the airtight seal required. It is important to constantly monitor the patient's demeanour, movements and respiratory rate on induction. Some animals can thrash in the induction chamber and injure themselves, others may become recumbent and place their ocular surface on the inside of the chamber, predisposing to corneal ulceration, so it is imperative they are constantly watched on induction. If there are any concerns about apnoea or a patient's demeanour during the induction process it is recommended to remove the patient from the chamber, assess its vital parameters and once satisfied continue the induction, either within an induction chamber or via face mask.
Larger mammals such as rabbits, ferrets and guinea pigs can be induced via face mask, however the restraint required may be stressful for them. Due to this the recommendation is to induce via intravenous agents wherever possible (Grint, 2013). Rabbits and most rodents are obligate nasal breathers, so if mask induction is utilised it is important not to obstruct the nares as this can prevent the patient from breathing and cause them to panic (Thompson and Bament, 2012). This is also important during maintenance of anaesthesia if the patient is maintained on a face mask (Figure 3), as obstruction of the nares and therefore the airways can lead to hypercapnoea, hypoxia, bradycardia and death. During induction with gaseous anaesthesia, rabbits are prone to breath-holding, especially if no premedication has been administered (Foote, 2018).
Once the patient is anaesthetised it is important to ensure that they are properly prepared for the procedure ahead. The oral cavity of rodents should be checked for any food that may obstruct the airways during anaesthesia, especially with hamsters that have large cheek pouches or guinea pig patients that routinely store food within the cheeks. When a patient with large abdominal visceral volume (e.g. rabbits, guinea pigs) is placed in dorsal recumbency it is important to elevate their chest. This is because the weight of the abdominal organs can press on the diaphragm and decrease the ventilation volume of the lungs (Allweier, 2016). The chest can be elevated by using a folded towel, foam wedge or sandbag under the patient's chest, neck and head area.
Intubation is possible in larger mammal species, however this depends on the patient's size and the anatomical considerations of the species. In ferrets intubation can be performed following application of topical lidocaine, using a similar process to intubating feline patients (Allweier, 2016). Rabbits can be intubated endoscopically, using an otoscope or via blind technique (Eatwell, 2014), however the authors would recommend first visualising the glottis to ensure no food material is obstructing the glottis prior to intubation (Figure 4). Maintenance of rabbit patients on a face mask has been shown to cause hypercapnia and hypoxia (Bateman et al, 2005), however significant trauma can occur with poor intubation technique in rabbits (Phaneuf, 2006). In cases where the operator is not confident or experienced to perform endotracheal intubation a laryngeal mask, such as a V-gel® (Millpledge Veterinary, United Kingdom) can be considered. V-gels® are a species-specific supraglottic airway device designed for the rabbit glottis and are shaped to fit in place over the glottis. Capnography is essential when using supraglottic devices as capnography provides the only safe way of evaluating whether the device is providing a patent airway (Richardson, 2015).
Monitoring anaesthesia
Patients can be maintained on inhalational anaesthesia such as isoflurane or sevoflurane, or on total intravenous anaesthesia. Any changes to the anaesthetic agent administration should be discussed with the veterinary surgeon as, under the Veterinary Surgeons Act 1966, Veterinary Nurses are only permitted to administer anaesthetic agents under direct instruction of the veterinary surgeon (RCVS, 2017).
Patient monitoring should start from the point of induction until full recovery. Patient compliance may inhibit monitoring of vital signs pre-induction and post-anaesthesia. Some multiparameter machines are fine enough to be able to monitor exotic patients, and the specifications of the machine should be checked prior to requirement. However, the primary monitoring method should be manual measurements taken by the veterinary nurse.
Cardiovascular monitoring
Cardiovascular monitoring allows assessment of heart rate, audible problems such as arrhythmias and murmurs, and pulse strengths. Neonatal or paediatric stethoscopes are recommended when monitoring small mammals due to their size. Oesophageal stethoscopes can be used in ferrets, rabbits and in some larger guinea pigs. Care should be taken when placing oesophageal stethoscopes to ensure excessive pressure is not placed on the tongue, impeding blood flow and causing cyanosis of the tongue.
When using an oesophageal stethoscope, it is important to pre-measure the length to reach the approximate heart position, then adjust to locate the point of optimal audibility. In most small mammals the heart is easily found approximately in line with the elbow of the patient. However, ferrets have a longer chest size in relation to the body size and the heart is located more caudally at the 6th–8th rib (Keeble and Heggie, 2012).
In small rodents where the thorax is not accessible due to the type of surgery (e.g. mammary mass removals), placement of a Doppler probe over the femoral artery, located on the medial thigh, is suggested. It can be beneficial to tape Doppler probes onto a tongue depressor to allow position alterations to be made from a distance. Evaluation of the femoral pulse, either digitally or by Doppler, can help assess pulse strength and regularity in correlation to heart auscultation. Weak or strong pulses could be a sign of hypotension or hypertension respectively and should be correlated with the remainder of the patient's assessment (Ateca et al, 2018). Arrythmias can be linked to the anaesthesia, for example lightening of an anaesthetic plane, hypovolaemia, or due to concurrent patient disease (Thompson and Bament, 2012).
When using oesophageal stethoscopes or a Doppler to monitor the cardiovascular system, it is important to have a normal stethoscope available in case of emergencies. Any movement of the patient will require rechecking the placement of the Doppler or oesophageal stethoscope to ensure optimum placement.
Respiration
Respiratory system monitoring should include evaluation of the rate, effort, depth and presence of any abnormal respiratory noise. Identifying problems can be difficult in rodents due to patient size.
Increased effort can indicate several potential problems such as endotracheal (ET) tube blockage or kinking, poor patient positioning or concurrent disease. Blockages of the ET tubes by respiratory secretions can cause increased respiratory effort and sometimes abnormal respiratory noise. Face masks can become dislodged, causing occlusion of the nares and a resultant increased respiratory effort.
Depth of respiration should be observed prior to anaes-thesia, and the respiration depth under anaesthesia compared with the initial observations. Shallow respiration can indicate the patient's anaesthetic plane is becoming light or is responding to stimulus from surgery. Deeper respirations can indicate the patient's anaesthetic plane is deeper. An assessment of anaesthetic planes in small mammals can be seen in Table 3. Increased depth of respiration can eventually lead to apnoea.
Table 3. Table showing planes of anaesthesia (Thompson and Bament, 2012)
| Plane 1 | Voluntary excitement | Increased heart rate, all reflexes present, increased respiration rate, potential for shallow breaths |
| Plane 2 | Involuntary excitement | Increased respiration rate, potential for shallow breaths, jaw tone present, slow palpebral reflex, deep pain present |
| Plane 3 | Surgical anaesthesia | Reflexes absent, some may remain with slight jaw tone or anal tone. Stable respirations and normal depth of respirations. Stable heart rate |
| Plane 4 | Medullary paralysis prior to death | All reflexes lost, decreased heart rate, reduced blood pressure, reduced respiration rate |
Reflexes
Reflexes should be monitored to aid assessment of the surgical plane of anaesthesia. When at a surgical plane, all reflexes should be absent from stimulus (Thompson and Bament, 2012). If only a light anaesthetic is required for minor procedures, for example blood sampling, jaw tone and palpebral reflexes may be present. In ferrets and rabbits, it can take quite a deep anaesthetic plane to lose the jaw tone.
Simple reflexes to monitor include pedal reflex, palpebral reflex, jaw tone, anal tone, muscle tone and deep pain. Each of these reflexes should be checked and recorded every time vital signs are recorded. Returning reflexes are indicative of lightening of the anaesthetic plane.
Testing of reflexes can be harder in smaller patients and care must be taken not to cause injury. Palpebral reflexes in mice, rats and similar sized rodents can be tested using a damp cotton bud compared with a finger. Table 4 shows at which plane of anaesthesia each reflex is expected to be lost.
Table 4. Table highlighting reflex testing and stages expected response. (Stanway and Morgan, 2003)
| Reflex | How to check | Anaesthesia plane reflex becomes absent |
|---|---|---|
| Palpebral | Touch medial canthus of eye with wet cotton bud. Assessing for any blinking reflex | Plane 2 into plane 3 |
| Pedal | Pinch between toes on the skin. Assessing for any response | Plane 2 into plane 3 |
| Anal | Apply small amount of pressure to the anal sphincter. Assessing for any response or twitching from sphincter | Plane 3 |
| Muscle Tone | Movement of limb. Assessing for how relaxed and easy to move limb is | Plane 2 |
| Jaw Tone | Attempt opening jaw by applying gentle pressure on the mandible and then releasing to relaxed state again. Assessing for any resistance on opening. | Plane 3 |
Temperatures
Temperatures should be closely monitored under anaesthesia as small mammals are more prone to hypothermia due to their large surface area in body size ratio in comparison to larger companion mammals (Thompson and Bament, 2012). Rectal temperatures can be used for all small mammals under anaesthetic and during recovery. Considerations should be made to patient size and thermometer probe size. Oesophageal thermometers can be used for rabbits, ferrets and some larger guinea pigs. As with oesophageal stethoscopes, it is important to monitor the tongue colour and vascularisation to ensure there is no vascular outflow compromise. Care should be taken when placing multiple oesophageal probes to not cause oesophageal trauma due to the number of monitoring probes placed.
Multi-parameter monitoring
Automatic monitoring can be useful, but it is important to be aware of the machine's specifications to ensure it is reliable when used to monitor smaller mammals. There are specific models of multi-parameter machines that are designed for small mammal monitoring if the machine in use is incompatible (Figure 5). Multi-parameter machines should always be used with caution as they can give falsely high or low readings, so all results should be correlated to the patient and manual assessments (Figures 6 and 7).
Rectal temperature can be constantly monitored via multi-parameter machines. Some machines have the option of oesophageal probe placement which can provide a more accurate core temperature than rectal readings.
Capnography is important for assessment of the gaseous exchange occurring in the lungs. If maintained with an ET tube the expired end tidal carbon dioxide should remaining between 35 mmHg and 45 mmHg (Thompson and Bament, 2012). Wave forms should be assessed to identify rebreathing, respiration depth and respiration rate. Mainstream capnography systems will evaluate gas exchange between the breathing system and ET tube, however dead space should be minimised in exotic patients by using side stream capnography (Grint, 2013). This system aspirates a small amount of gas for analysis and therefore may not be as accurate as mainstream capnography, but trends in end tidal CO2 can be monitored.
Capnography should be utilised on all intubated patients or those with a laryngeal mask. Where no connection port exists on the ET tube connector, a 25G needle can be inserted into the tube lumen and the capnograph attached to the needle. Capnography can also be utilised to monitor for partial or full blockage of the ET tube, which is identified as a prolonged expiratory phase (Hawkins and Pascoe, 2012). This is reported more commonly with smaller species due to the reduced diameter of the smaller ET tubes required, however the risk can be reduced slightly by using an ET tube with a Murphy's eye, if possible (Hawkins and Pascoe, 2012). Capnography can also be used in obligate nasal breathers maintained via tight-fitting face mask by placing the sampling line just rostral the nares, ensuring that this does not cause nare obstruction.
Blood pressure should be taken, if possible, at a minimum of 5-minute intervals. Hypotension can be seen due to hypothermia, hypovolaemia and as a side effect of some medications. Troubleshooting can include reducing the anaesthetic agent, administering fluid boluses, or providing vasoconstriction agents. Hypertension could suggest a lightening plane of anaesthesia or a response to surgical stimulus. When measuring blood pressure in small mammals the absolute numbers may not be accurate due to the small size of the patient, however following blood pressure trends and interpreting the values alongside other monitoring values is of use to evaluate the patient's plane of anaesthesia. In general, a decrease in blood pressure acutely or over time indicates either a deepening plane of anaes-thetic or a circulating volume loss, and a trend of increasing blood pressure indicates a lightening plane of anaesthesia (Thompson and Bament, 2012).
Measuring blood pressure can be difficult in smaller rodents due to the size of their limbs and available cuff sizes. Monitoring is possible in guinea pigs, ferrets and rabbits and similar sized small mammals. Correct cuff size should be determined by ensuring the width of cuff is approximately 30–40% limb circumference and when applied it stays within the range markings displayed (Thompson and Bament, 2012). When using a Doppler probe to measure systolic blood pressure the probe can be held in place with tape to ensure it does not move during the procedure. Clipping of fur to enable contact with a Doppler probe should not be performed in rabbits due to the risk of pododermatitis.
Electrocardiography (ECG) can be performed on all patients under anaesthetic, however machines that are not calibrated for high heart rates can struggle with the trace reading. Low pressure ECG clips without crocodile teeth or ECG pads should be used to minimise injury to the patient. In some situations, 25G needles can be passed through the skin and connected to the ECG leads (Thompson and Bament, 2012). Alcohol spray can be used to improve patient contact, however excessive use can lead to hypothermia.
Pulse oximetry is often only possible in rabbits and ferrets, with placement on the tongue being easily accessible and highly vascular. Placement of large clips on small tongues can reduce blood flow via compression. Non-haired feet can be used for probe placement in smaller rodents but is not always reliable.
Pain
Any patient undergoing a painful procedure should have analgesia administered as part of their premedication. Indications of patient pain while under anaesthesia include tachycardia, deeper respirations or panting, hypertension, and some return of reflexes. When patients show these signs it is important to administer appropriate analgesia if necessary, rather than increasing the anaesthetic agent. Constant monitoring can help identify pain at the earliest opportunity so that action can be taken to prevent anaes-thetic complications.
Recovery
Recovery of patients starts as soon as anaesthetic administration is ceased. Patients should be maintained on oxygen until they are extubated, ambulatory and responsive. Recovery from anaesthesia is associated with a high risk of complications (Geoffrey, 2019) and so during recovery patients should be monitored at the same level as under anaesthesia. The recovery area should be pre-warmed and draught free. If there are concerns about respiratory compromise, then pre-oxygenating the kennel is advised. The recovery kennel should be in a quiet area and ideally positioned to allow patient observations with minimal disturbance.
Patients should be positioned in sternal recumbency to prevent atelectasis, unless this is contraindicated by the surgical procedure. Elevating the chest is beneficial to prevent pressure on the diaphragm by visceral organs (Figure 8). Patients may be ataxic on recovery, so it is important to maintain them in sternal recumbency and ensure they do not injure themselves in the kennel during the recovery process. Rolled towels or blankets either side can assist with this.
Food should be offered once the patient is ambulating and swallowing. It is important to encourage feeding as soon as patients can tolerate it (Figure 9), and if a patient is not voluntarily eating then syringed nutrition can be administered (Figure 10). Prolonged recovery times in ferrets can be caused by hypoglycaemia, so blood glucose monitoring should be performed on ferrets with protracted recoveries. Gastrointestinal mobility of small herbivores can be decreased following anaesthesia or sedation. Syringe feeding high fibre food supplements can aid in restoring gastrointestinal mobility.
Bonded companions should be placed back in with the patient as soon as they have recovered from anaesthesia. Ideally, the patient should be offered food and water before the companion is returned to encourage eating without any competition. Any active warming should be removed prior to re-introducing bonded animals to prevent hyperthermia of the companion. Conspecific interactions should be closely monitored as there is a small risk of aggression following anaesthesia due to stress and abnormal smell.
Special considerations
There are a number of special concerns when anaesthetising small companion mammals compared with dogs and cats. Most of these considerations can be easily prepared for by modifying procedures for cats and dogs, however it is always important to be prepared.
Paperwork
All small mammal anaesthesia should have the appropriate paperwork completed, including a signed consent form and signed off-licence medications form. Anaesthetic charts may need to be modified to allow for recording of higher average respiratory rates and heart rates of smaller mammals under anaesthesia. Some practices choose to have separate companion animal and exotic mammal anaesthetic sheets to allow accurate recording of these parameters.
Appropriately sized equipment
As described above, there are a number of devices that can be used to monitor patients' vital parameters under anaes-thesia, however it is important that the monitoring equipment and anaesthesia equipment is appropriately sized for the patient. Figure 11 shows a patient wearing a rabbit specific medical pet shirt, these can be adapted to suit other small mammals as well for wound protection.
Hypothermia
Small exotic mammals have a higher surface area to body mass compared with larger companion mammals and therefore are more likely to lose heat under anaes-thesia when they are unable to regulate their body temperatures (Longley, 2008b). Supplemental heat should be provided, in the form of either a thermostatically controlled heat mat, a Hotdog®, warm air blankets (e.g. Bair Hugger®), overhead heat lamp, hot water bottle or ‘hot hands’. Warming devices should never be in direct contact with the patient as this can predispose to thermal burns, as an anaesthetised patient is unable to react to stimuli. Other warming methods can be used to maintain the patient's temperature, including providing a warm ambient temperature, wrapping extremities such as ears, feet and tails with insulating material such as bubble wrap or towels (Figure 12), covering the body with plastic surgical drapes rather than cloth and clipping as minimal an area as possible to allow the fur to naturally insulate the patient (Thompson and Bament, 2012). A heat and moisture exchanger can be added to the anaesthetic circuit to combat heat loss through the respiratory system, however these can increase dead space and create increased resistance for the patient to breathe against (Lamb, 2009).
Ocular complications
Due to the inhibition of blink reflexes while under anaesthesia, small exotic mammals are more prone to ocular desiccation while anaesthetised (Thompson and Bamnet, 2012). This can lead to painful corneal ulceration and associated complications. Patients should undergo liberal ocular lubrication prior to induction and during anaesthesia (Figure 13). Taping the eyes closed can also help to avoid this complication, however it should be ensured that the tape does not come in to contact with the cornea, as any rubbing can also result in corneal ulceration.
Gastrointestinal considerations
Gastrointestinal stasis (ileus) is a consideration in small herbivorous mammals such as rabbits, guinea pigs, chinchillas and degus. Close attention must be paid to the food intake and faecal output of these patients. Decreased appetite and smaller or absent faeces can be indicative of ileus. Adrenergic stress, such as that encountered during hospitalisation and anaesthesia, will reduce the mobility of the gastrointestinal tract and predispose to ileus (Varga, 2014). Steps must be taken at every opportunity to reduce stress in these patients, as described previously. Pre-emptive prokinetic drugs can help combat the onset of ileus and can be used as part of the premedication or immediately post operatively, including metoclopramide, cisapride and ranitidine (Longley, 2008b).
Inadequate analgesia following surgery is a major cause for ileus in patients (Allweiler, 2016). As such it is important that all patients receive adequate analgesia, which initially post operatively is likely to involve a combination of opioids and non-steroidal anti-inflammatory drugs. Pain should be assessed based on the behaviours and appearance of the individual patient, but generally signs of pain include a hunched posture, reluctance to move and vocalisations (Allweiler, 2016). Grimace scales have been developed to help identify signs of pain in small exotic mammals, and currently are available for rabbits (Keating et al, 2012), mice (Langford et al, 2010) and rats (Sotocinal et al, 2011) based on evaluation of facial expressions.
Conclusion
Monitoring the anaesthesia of exotic small mammal patients should not be daunting to veterinary nurses, as the principles are very similar to that of the canine and feline patients anaesthetised daily. However, there are a few factors to consider relating to their anatomy and physiology such as fasting patients, raising chests and heat loss in small mammals. It is important to know the species to know how to approach these in the best way for the patient. A whole-team pre-anaesthetic discussion should occur prior to pre-medication and induction, in order to allow the anaesthetic to become much safer and less stressful for both the veterinary team and the patient.
KEY POINTS
- Suitable multi parameter machines can be used on small mammals in the same way as canine and feline patients under anaesthetic, with understanding of their species-specific values.
- Stress should be kept to a minimum to help reduce complications during induction and maintenance of anaesthesia.
- Raising the thorax in most small mammals can have a large benefit to their thoracic volume and expansion, and therefore aids their breathing pre-, peri- and post-anaesthesia.
- Nutrition pre- and post-anaesthesia is an important part of reducing complications in recovery.
- Constant monitoring should occur from the administration of a premedication through until the patient is fully recovered, in order to identify and resolve any problems as soon as possible.