The immediate recovery period
The swallowing and cough reflexes are usually suppressed during anaesthesia, and these gradually return as the animal recovers consciousness. If an endotracheal tube is present, it should be removed when the animal begins to swallow spontaneously or attempts to cough. If the tube has been tied in position, the ties should be loosened in anticipation of the need to remove the tube. Care must be taken that the tube is not pulled out too soon, for example, when the animal is repositioned as surgical drapes are removed.
In this immediate recovery period, use of a pulse oximeter to assess respiratory function is extremely useful, particularly in larger species. Changes in oxygen saturation after disconnection from the breathing system should be assessed. An observation period of approximately 2 minutes is usually sufficient to assess the fall in oxygen saturation which occurs when changing to breathing room air rather than the higher oxygen concentration in anaesthetic gases. If saturation falls below 85%, then oxygen should be administered and ventilation supported or stimulated. If anendotracheal tube is in use, if it is a cuffed tube, deflate the cuff and gently remove the tube as the protective pharyngeal and laryngeal reflexes return. After removal of the endotracheal tube (if one has been used), maintain the animal on a face mask and administer oxygen. If respiratory movements and oxygen saturations are judged to be adequate, remove the mask and continue to monitor the animal.
Removal of the endotracheal tube often causes a fall in oxygen saturation as the airway is not as well-maintained. Before removal of the tube, or if one is not used, before recovery of swallowing and cough reflexes, the mouth should be inspected and any secretions removed using suction. A soft tipped catheter (e.g. a feeding tube) connected to a large syringe allows this to be carried out in small rodents. Monitoring of adequacy of ventilation can also be assessed using a capnograph, and this is particularly useful following prolonged periods of anaesthesia, especially if the animal has been mechanically ventilated.
During this initial recovery period, drapes and surgical equipment should be removed, together with any non-essential monitoring devices. This will allow the animal to be moved rapidly to a more comfortable environment if it regained consciousness more rapidly than expected.
The respiratory depression produced by most anaesthetic agents often persists into the post-operative period. The degree of depression may also increase post-operatively, and this may go unnoticed until severe hypercapnia and hypoxia have developed. For this reason, it may be advisable to continue monitoring the respiratory system, and the use of a pulse oximeter is ideal for this purpose, particularly in larger species. If not already in use, the probe can be attached to the animal in the operating theatre and a battery-operated instrument used to monitor the animal during movement to the recovery area. The probe can be left taped in place on the tail or on a digit until the animal has regained its righting reflex.
If a pulse oximeter is unavailable, then other forms of respiratory monitor can be used, for example, positioning a sensor close to the animal’s nose. At the very least, regular clinical observation of the animal should be made and the respiratory rate recorded. If respiratory depression is noted, it should be treated using a respiratory stimulant such as doxapram and by the administration of oxygen. Since doxapram has a relatively short duration of action (10–15 minutes), it may be necessary to administer repeated doses or to establish a continuous infusion of the drug.
Many animals appear to benefit if oxygen administration is continued into the immediate post-operative period. This is best achieved in small animals by piping the gas into an animal incubator, but in large animals, it is often more practicable to tape a small, soft-ended catheter at the external nares and use this to administer the gas. If oxygen is not to be provided, it is reassuring if oxygen saturation can be monitored for several minutes after the animal commences breathing room air, to ensure that saturations above 85–90% are maintained.
Non-ruminant species should be placed on their sides, with head and neck extended, to try to minimize the risk of airway obstruction. If the animal is recumbent for more than 4 hours, then it should be repositioned to lie on its other side, to prevent passive congestion of the lungs and the development of hypostatic pneumonia. In large animals such as dogs and farm animals, it may be necessary to massage areas such as the elbow and hock, to prevent pressure sores developing. If prolonged recumbency is anticipated, it may be advisable to protect these areas with padded bandages.
If the animal begins to vomit, it should be positioned so that its head is below the level of the thorax and abdomen, to try to prevent aspiration of the vomit. If practicable, the mouth and pharynx should be cleared using a vacuum suction, or a piece of suitably sized tubing attached to a 50-ml syringe. Oxygen should be administered, and if inhalation of vomit could have occurred, corticosteroids should be administered (30mg/kg iv of methyl prednisone), together with a broad-spectrum antibiotic.
Ruminants (sheep, goats and cattle) can present particular problems during recovery from anaesthesia. They should be propped up on their sternums to minimize the risk of over-distension of the rumen with gas (rumenal tympany) and to reduce the risk of inhalation ofregurgitated rumen contents. If rumenal tympany develops, it should be relieved immediately either by passing a stomach tube or by puncturing the rumen through the left abdominal wall with a large-bore trochar. If a trochar is not available, the largest possible needle (preferably 12 SWG or larger) should be used. If the member of staff involved is not familiar with this technique, veterinary advice should be sought immediately.
Fluid therapy
The voluntary water intake of all animals should be recorded post-operatively, even if this consists simply of making a rough estimate based on the level in a water bottle. Fluid intake is frequently reduced post-operatively, and if dehydration is allowed to develop, it can seriously compromise the recovery of the animal. Fluid requirements of most species are approximately 40–80 ml/kg/24 h, but the presence of vomiting or diarrhoea or other abnormal losses will increase this requirement.
If the animal is fully conscious, supplemental fluid is best given by the oral route. If the animal is unable or unwilling to accept oral administration, then dextrose–saline (4% Dextrose, 0.18% Saline) or saline (0.9%) can be given quickly and easily by the subcutaneous or intraperitoneal routes. Severe dehydration causes loss of skin tone that causes it to tent and tend to remain elevated when a fold is twisted between the fingers. In larger animals, dehydration will result in the mucous membranes becoming dry to the touch. If this degree of dehydration has inadvertently been allowed to develop, fluids must be administered intravenously.
The monitoring of body weight in the pre- and post-operative periods can provide a good indication of the adequacy of fluid intake. Although a small fall in body weight (<1-3% reduction) will be recorded because of the almost inevitable reduction in food intake that occurs post-operatively, most weight loss usually represents a fluid deficit.
Besides assessing food and water intake, the urinary and faecal output of the animal should be recorded and any abnormalities investigated. As with most of these variables, a meaningful judgement can only be made if the animal has been observed in the pre-operative period. A reduction in urine output may be the result of dehydration, urinary tract injury, or the animal suffering pain. If the bladder is full, it may require catheterization to empty it, although in some instances gentlepressure through the abdominal wall will trigger urination. Catherterisation is a relatively simple technique, but requires some degree of expertise, and it will usually be preferable to consult a veterinary surgeon or experienced animal technician. If catheterization is not possible, it may prove necessary to drain the bladder by direct puncture through the body wall. This procedure should only be attempted by individuals who have undergone training in the technique. Catheterization of the bladder of most laboratory species requires induction of a brief period of general anaesthesia, or heavy sedation.
Food intake and bowel function
If the animal fails to pass faeces, this may be due simply to an absence of faecal material because of pre-operative fasting. It may also be caused by a loss of normal peristalsis (ileus, see below), or the animal may be constipated and require administration of an enema (e.g. Microlax, SmithKline Beecham). Defaecation may also be suppressed if the animal is in pain, particularly following a laparotomy.
Ileus (gut stasis) can be a serious post-operative complication, and can be life-threatening in rabbits and guinea pigs. Ileus is particularly common after laparotomy, but can also occur after any surgical procedure. The incidence of ileus following abdominal surgery can be reduced by minimising handling of the bowel. When displacing and handling the viscera is unavoidable, ensure they are kept moist and handled gently.
If ileus is suspected, then motility stimulants (metaclopramide and cisapride) can be administered to stimulate gut function. In rabbits, ranitidine (2–5 mg/kg by mouth, daily) has been used for managing post-operative inappetance and gut stasis as it promotes gut motility. Pain must be controlled, since this can increase the severity of ileus. The surgical notes should be reviewed to check that a swab was not inadvertently left in the abdomen.
In some species (e.g. rabbits and guinea pigs), inappetance due to other causes can lead to the development of ileus, since normal gut function appears to depend to some extent on regular intake of fibre.
Supplemental feeding, using nasogastric tubes if necessary, may be beneficial. A range of specialist dietary preparations are now available for veterinary use in “exotic” companion species, and these can be of considerable benefit in laboratory animals. It is important that food and water intake and the other observations described above are recorded carefully. It is helpful to provide a standardised paper or electronic record for each animal, which will encourage nursing staff to complete the required observations. It will also allow easy and rapid reference by staff who may be called in to deal with any problems that might arise. It is always preferable to obtain measures of pre-operative body weight and, if possible, of food and water consumption, so that the progress of an animal can be assessed accurately in the post-operative period. Obtaining daily weights for 3–4 days pre-operatively both familiarises the animal with handling, and provides a base-line growth curve to allow interpretation of post-operative changes.
Prevention of wound infection
Provided careful aseptic surgical techniques have been employed, it may be considered unnecessary to administer antibiotics routinely to animals in the post-operative period. In addition, some species appear to show a remarkable resistance to the development of wound sepsis and appear to tolerate standards of cleanliness that would be totally unacceptable in human medical practice. This apparent resistance to infection must not be used as an excuse for poor surgical standards, and every effort should be made to adopt aseptic techniques for all animal surgical procedures (www.procedureswithcare.org.uk). It has been demonstrated, for example, that rats are not only susceptible to infection but also show behavioural changes following the establishment of wound infections. It is therefore important that all animal species should be monitored carefully for any signs of infection.
Since animals will almost inevitably soil their wounds with faeces and urine, administration of prophylactic antibiotics may be useful in minimizing the risk of infection. One problem of providing peri-operative treatment with antibacterials is the risk of inducing enterotoxaemia in some species, particularly the guinea pig, hamster and rabbit. The use of antibacterial agents in rodents and rabbits has been reviewed by, and provided care is taken in the choice of agent, such problems can be avoided. For advice on the use of antibiotics, contact your facility veterinarian.
Nursing care
The response to human contact varies considerably among different animal species and is influenced by previous experiences. Excessive contact may have adverse, stressful effects in some small rodents and rabbits, but other species will benefit from some degree of nursing care carried out in a calm and reassuring manner. The degree of alarm caused to the animal can be reduced if it has been gradually familiarized to regular handling in the pre-operative period. This process forms an important part of pre-operative acclimatization in all species (see above) and should be considered essential when planning any series of experiments.
All species, including rodents and rabbits, should be checked at least once a day. In the immediate post-operative period, constant attention may be needed, followed by observation every 1–4 hours for the first 8–12 hours. Particular attention should be given to cleaning the eyes, nose and mouth, which can become clogged with dried mucus or other debris. Monitoring of body weight and checking of wounds and surgical implants are also an important part of post-operative care. Rodents may be offered food pellets softened with warm water in bowls placed on the cage bottom, as many may be reluctant to reach up to food hoppers at this time.
It is important that a daily routine is followed as far as possible. It will be an advantage if some staff are assigned specifically to the care of post-surgical animals throughout the peri-operative period, as they are more likely to notice subtle changes that may take place on a day-to-day basis. Careful record keeping is essential, so that other staff attending the animal, for example during week-ends and out-of-hours, will be aware of all treatments given and the animal’s progress. It is important to record not only all active interventions, but also that the animal has been examined and found to be progressing satisfactorily. Records of clinical observations and treatments must be readily available, and use of electronic record systems can be of great assistance in providing legible, easily accessible and up-to-date information.