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Treatment and Care (LARGE PAGE - wait to load)

Bears: Click here for full-page view with caption Click here for full-page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption Rabbits:Click here for full page view with caption. Ovariohysterectomy of a rabbit doe with hydrometra. Sternal recumbency over the edge of a table for jugular access. Click here for full page view with caption. Rabbit. Surgical area prepared for castration. Click here for full page view with caption. Rabbit draped for castration. Click here for full page view with caption. Rabbit caudal ear vein cannulation. Click here for full page view with caption. Rabbit cephalic vein cannulation. Click here for full page view with caption. IM injection longissimus dorsi. Click here for full page view with caption. IM injection quadriceps femoris. Click here for full page view with caption. Rabbit prepared for ovariohysterectomy (spay). Click here for full page view with caption. Subcutaneous injection. Click here for full page view with caption. Cranes:Induction inhalational anaesthesia, crane. Click here for full-page view with caption. Maintenance inhalational anaesthesia, crane.  Click here for full-page view with caption. Anaesthetic induction, crane wearing hood. Click here for full-page view with caption. Anaesthetic recovery, crane being held. Click here for full-page view with caption.

Introduction and General Information

Depending on the disease and the number of individuals involved, the treatment and/or control of disease may require management and manipulation of the population and/or its environment, or treatment and care of an individual or group of animals.

For information on manipulation of the population and environment see Environmental and Population Management

Waterfowl Consideration

  • Where disease in wild waterfowl is concerned, manipulation of the population and the environment are most commonly employed to reduce the effect of disease outbreaks. However, treatment and care of individual waterfowl or groups of waterfowl may also be appropriate, for example in the cleaning of birds affected by Oiling, Hook and Line Injuries or Avian Botulism.
  • In captive waterfowl, the treatment and care of individual birds is employed more commonly, although consideration must also be given as to the role of the population and the environment in the development of the disease, and the possible effect on disease incidence of manipulating these.

For information on manipulation of the population and environment see Environmental and Population Management

Crane Consideration
  • While preventative treatment is generally conducted on a flock basis (e.g. administration of anthelmintics, use of coccidiostats in feed), treatment is mainly on an individual basis. There are exceptions such as in rehabilitation settings when a group of cranes have been affected by a single incident.

For information on manipulation of the population and environment see Environmental and Population Management

Bear Consideration
  • In zoo and wildlife rehabilitation settings, usually treatment will involve a single bear. However, group control of disease may be required for infectious diseases, including parasite infestations, and environmental management may also be required, particularly for parasite control. 
  • In rescue situations, large numbers of bears may require individual treatment to repair damage caused by years of abuse. (P503.1.w7)

For information on manipulation of the population and environment see Environmental and Population Management

Lagomorph Consideration

  • For pet rabbits, generally the treatment and care of individual rabbits is most important. However, in multi-rabbit households, including rabbit rescue centres and rehabilitation centres, consideration must also be given to population management, particularly for infectious diseases, including parasite infestations, and environmental management may also be required, particularly for parasite control.
  • In zoo situations, both individual and colony care is important.
  • In rabbit breeding establishments, and commercial and laboratory rabbits, population and environmental management is extremely important.
  • Note: The majority of the information on this page is based on the treatment and care of domestic rabbits. In general, procedures used should be the same for wild lagomorphs. Additional information relevant for wild individuals will be presented where available.

For information on manipulation of the population and environment see Environmental and Population Management

Ferret Consideration
  • For pet ferrets, the treatment and care of the individual ferret is likely to be most important. In multi-ferret households, population management may also be important.
  • Staff who might be infected with influenza virus should be kept away from ferrets as they are susceptible to these viruses and serious illness can result in immunocompromised ferrets. (J29.6.w3)
Bonobo Consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.

Care and treatment of bonobos is particularly aimed at the individual. For conditions such as respiratory infections, consideration of group care is also important.

  • NOTE: Positive reinforcement training facilitates individual medical treatment during serious illness. Additionally, by increasing trust between keepers and bonobos, it can enable temporary removal of infants for medical care. (P1.2002.w10)

Medication

  • Drug doses for adult Pan troglodytes - Chimpanzee are often on a "per individual" basis using human dose and drug recommendations. (D409.6.w6) For paediatric patients, a mg/kg dose rate may be more appropriate. (D409.6.w6)
  • Use of all medications will be off-label. (D409.6.w6)
  • Antibiotics should be used only after evaluation of an individual case indicates there is a real need for their use (e.g. secondary bacterial infection confirmed in an individual with viral respiratory infection), and should be chosen on the basis of culture and sensitivity testing whenever possible. (D409.6.w6, B10.44.w44g)
    • When it is not possible to base antibiotic treatment of primates on cuture and sensitivity testing then a broad-spectrum antibiotic should be used. (B10.44.w44g)
    • Antibiotic treatment should continue for at least two full days after clinical signs of disease have stopped. (B10.44.w44g)
  • A list of drugs is provided (Drugs used in the treatment of Bonobos).Dose rates which may be/have been used in bonobos/other great apes are given on the individual drug pages.

For information on manipulation of the population and environment see Environmental and Population Management

Published Guidelines linked in Wildpro

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Injection and Medication Techniques

  • The appropriate route for administration of medication will vary depending on factors including the severity and site of disease, type, volume and formulation of medication, and the size, number, feeding habits and temperament of the animals to be treated. Economic factors, requirements for dosing by the owner (rather than by a veterinarian) and legal registration requirements must also be considered.
  • Topical medication is appropriate for use on the skin, and for ocular treatment.
  • Most medications are given orally or by parenteral routes - mainly by subcutaneous, intramuscular or intravenous injection. Other routes such as intraosseous, intraperitoneal, intratracheal or intracardiac are used less commonly. Nebulisation is used when medication is to reach the lower respiratory tract.

Further information about the uses and limitations of the main routes of drug administration is provided in Routes of Drug Administration in Ruminants (Techniques Overview)

Waterfowl Consideration

  • Oral medication is indicated when there is normal gastro-intestinal tract motility, and offers the widest range of options with regard to administration and formulation. Medicated feed and water (Food / Water Medication for Birds) have their main advantages in treating diseases on a flock basis. Capsules and tablets (Capsule / Tablet Administration in Birds) can be easily administered to handleable birds and can be used to deliver precise amounts of medication. Administration by gavage (Gavage / Tubing of Birds) is an excellent way to initially improve gastro-intestinal tract motility in a debilitated animal, and provide longer term hydration and nutrition.
  • Parenteral administration is any mode other than topical or through the gastrointestinal tract. This usually means delivering medication through a needle to a specific anatomic site. Action at site, or circulating blood levels of drug are achieved more rapidly than oral medications. The potential for introducing septic agents is higher, so cleanliness is much more important. Tissues to which agents are delivered may be damaged. Needle sizes and volumes delivered must be chosen carefully, according to the size of the bird. (B11.5.w18, V.w5, V.w7).
Crane Consideration
  • Oral medication can generally be achieved by placing tablets or capsules inside a treat such as a dead mouse or fish then feeding this to the crane.  Oral medication can also be given by gavage or as a bolus. (B12.56.w14)
    • Oral medication is preferred for medications which may cause muscle necrosis if the intramuscular route is used. (B336.20.w20)
  • Medicated food is used for whole-flock treatments, such as a coccidiostat for control of coccidiosis. (B336.20.w20)
  • Medicated water is also used sometimes for prophylactic administration of coccidiostats. (B336.20.w20)
    • Vitamin D and some other medications can be administered in drinking water if the crane is drinking. (B12.56.w14)
  • Parenteral routes - intramuscular, intravenous and subcutaneous -  are used as appropriate. (B336.20.w20)
    • For intramuscular injections the pectoral muscles are recommended as the site of injection. (B703.10.w10)
    • For subcutaneous injections, several sites can be used; commonly such injections are given over the lateral thighs. (B703.10.w10)
  • Nebulisation is used when it is important for medication to reach the lungs and air sacs. This is used alongside parenteral medication. (B336.20.w20)
    • Both antifungal agents and antibiotics cane be given by nebulisation.(B12.56.w14)
    • Appropriate antibiotics include gentamicin (1 mL) or tylosin (2 mL) in combination with 1 mL acetyle cysteine in 15 mL normal saline, administered using a 1-2 litre flow of oxygen for 20 minutes three times a day. (B12.56.w14)
  • Note: administration of antibiotics in food or water is not ideal due to the inability to control the dose given (variation in consumption). (B12.56.w14)
Bear Consideration

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Oral, parenteral and topical medications may be used.
Oral medication
  • Oral medication is advantageous where possible, to remove the stress and risks associated with physical restraint (e.g. in a squeeze cage, see: Mammal Handling & Movement) for hand or pole injection, or with remote injection (darting). However, this can only be used when both (a) the bear will consume the required medication (usually hidden inside a food treat) and (b) it is possible to ensure that the correct individual will eat the medicated food.
  • Oral medications can be mixed into foods such as milk, honey, cod liver oil, meat or fish, or placed inside a small treat which will be eaten whole. Liquids or powders can be placed inside a gelatin capsule which is then hidden in the treat. (P6.6.w3, B16.9.w9)
  • Bears have an excellent sense of smell and may detect medicines and refuse to eat food containing medication. (D247.8.w8)
  • Individual bears vary greatly in what they will be willing to eat. (D247.8.w8)
  • In general, sweet foods can be used to camouflage drugs for bears. Powders or ground up pills may be given inside thick honey sandwiches; similarly liquid medications may be mixed with honey. (D247.8.w8)
  • It may be possible to conceal medication inside the carcass of a rabbit or chicken. (D247.8.w8)
  • When a group of bears is to be medicated orally, the bears need to be separated into individual dens to ensure that all animals can consume the medicated food, rather than dominant animals being overdosed and subordinate animals not getting any of the medication. (B16.9.w9)
Parenteral medication
  • Parenteral administration by remote injection (darting) is required in many circumstances, including for medicating free-living bears.
    • For intramuscular injections it is important to be aware of the anatomy of bears, in particular the large quantities of subcutaneous fat which may be present over the rump and hind legs of hibernating species in late summer and winter, and polar bears at any time of year. Therefore injecting into the shoulder or neck muscles is preferable. (B16.9.w9, D156.w2)
      • When delivering drugs by remote injection (darting), the rifle must be powered correctly; if a charged dart hits with too low velocity, the cushioning effect of the bear's thick fur and fat may be such that the firing cap fails to fire due to insufficient impact. (B16.9.w9)
      • Needle lengths of at least 7.5 cm (3 inches) are required for intramuscular injections into adult bears. (B10.48.w43, B16.9.w9)
      • See: Intramuscular Injection of Bears
      • Note: bears can be trained to allow injections by hand or using a pole syringe, without physical restraint. See: Mammal Handling & Movement - Husbandry Training
    • For intravenous access (in anaesthetised bears), the sublingual vein is most accessible; fat layers make the jugular and femoral veins more difficult to access. (B10.48.w43) Various veins can be used and different individuals prefer different veins for intravenous access.
  • See below, anaesthesia and chemical restraint, for more information on the use of anaesthetic agents.
Topical medication
  • Topical medication is needed for treatment of a number of skin diseases. Depending on the bear's cooperation, the precision needed in application and the volume of medication, it may be necessary to treat the bear while it is physically or chemically restrained, or it may be possible to apply topical medication to an unrestrained individual.

Lagomorph Consideration

Both oral and parenteral routes of delivering medication are used in lagomorphs.

Oral/enteral medication
  • Oral medication may be given in food or more commonly in water. However, it is difficult to control the amount taken, and many medications have an unpleasant taste. (B600.3.w3)
  • It is usually easier to give a rabbit a liquid medicine; tablets can be crushed and suspended in water (preferably flavoured).
  • It may be possible to give tablets crushed and mixed with jam or another favoured treat such as banana. (B602.14.w14, B606.17.w17, J213.9.w1)
  • Alternative methods include feeding liquid medications via a syringe, using a nasogastric tube, using a pharyngeal or oesophageal tube or (for a single administration) via an orogastric tube. 
  • Alternative routes which may be considered for repeated administration of medication include a pharyngostomy tube or a gastrostomy tube. Techniques for placing these in rabbits have been described, but various complications have been reported associated with their use. (J213.9.w1)
  • Details of different oral medication routes are provided in:
Parenteral medication
Topical medication
  • Topical medications are used in the treatment of wounds.
  • Care must be taken if applying a topical preparation which is then not covered by a dressing, that there will be no harmful effects if the preparation is ingested by the rabbit. 
  • A special use of topical medication is EMLA cream (ASTRA Pharmaceuticals Limited, King's Langley, UK). This local anaesthetic cream (Eutectic Mixture of Local Anaesthetics) contains lidocaine (2.5%) and prilocaine (2.5%). It is used on the ears of rabbits to allow pain-free blood sampling form or placement of catheters into the marginal ear vein. The cream is placed over the ear vein, covered with clingfilm or a dressing and left for 45 - 60 minutes before taking a blood sample or inserting a catheter. (B600.3.w3, J213.9.w1) It can also be used over the lateral saphenous vein. (B601.2.w2) Prior to venepuncture, remove the cream and wipe the site with 70% isopropyl alcohol. (B601.2.w2)
Ferret Consideration
Parenteral
  • Many medications can be given by Subcutaneous Injection in Ferrets or Intramuscular Injection in Ferrets. (P120.2006.w6)
    • To distract a scruffed ferret while injecting it, offer a treat (e.g. a fatty acid supplemment, cat hairball laxative paste or a sweet nutritional supplement) on the end of a tongue depressor (use of the tongue depressor avoids the risk of injury to the person offering the treat). (J29.6.w1)
    • Generally, the subcutaneous route is preferred over the intramuscular route, as muscle masses on a ferret are small, particularly in cachectic ferrets. (B339.9.w9, B602.2.w2, P120.2006.w6) 
  • If an intravenous catheter is in place (Venipuncture / Blood sampling / Intravenous Catheterization in Ferrets ), antibiotics can be given intravenously; this is preferable in sick ferrets. (B602.2.w2)
Oral
  • Ferrets dislike pills; whenever possible, oral medication should be given in a liquid form, by syringe. (P120.2006.w6)
    • Pills and capsules are unlikely to provide the correct dose rate for ferrets. (B339.9.w9)
    • Cat hairball laxative, fatty acid supplements, strained meat baby food, molasses, honey, cherry-flavoured syrup, banana-flavoured syrup, chocolate-flavoured syrup (not with real chocolate) or sweet nutritional supplements can all be used. (J29.6.w3)
      • Avoid using sugary flavourings if possible. (P120.2006.w6)
      • Avoid using a high-sugar treat if taking a blood sample to test glucose levels. P120.2006.w7)
      • Avoid using fish-flavoured bases for medication; these are not palatable for ferrets. (B602.2.w2)
    • To give pills, preferably crush the pill and mix it with one of these substances. (J29.6.w3)
    • See: Oral Medication and Syringe Feeding of Ferrets
Bonobo Consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
  • Preferred drinks can be used for administration of oral medication. (B437.w24)

Great ape information:

  • In primates, the femoral vein is readily accessible at the femoral triangle; the cephalic vein and (posterior surface of the lower leg) saphenous vein also usually are accessible, particularly in larger individuals. These veins can be used for intravenous injections. (B10.44.w44e, B538.33.w33)
    • In great apes, the brachial vein (accessible at or distal to the antecubical fossa), median antebrachial, posterior tibial and femoral veins are accessible. (B336.39.w39)
      • The cephalic vein is the preferred site in great apes. (B214.3.1.w18)
    • Note: in an individual with low blood pressure, for example due to dehydration, it may be hard to catheterise a vein. A sterile cut-down is preferable rather than prolonged attempts to access the vein. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
    • In infants, skill is required to insert an intravenous catheter into e.g. the saphenous vein. If giving fluids, close monitoring and often physical restraint of the infant is required to keep the catheter in place while fluids are given. For an acutely ill infant, it is preferable to carry out a cut-down onto the saphenous vein, place an indwelling catheter and secure this in place, providing constant intravenous access for fluids and other drugs (and allowing repeated blood sampling, if needed). (B678.w8)
      • A wide-luen needle (0.0 mm) is recommended, plus regular flushing with heparin to maintain patency. (P3.2005b.w2)
    • Intraosseous administration of fluids may be required if intravenous access cannot be obtained. Note: in humans, intraosseous administration of fluids is known to be extremely painful. This route should be used only in an anaethetised individual. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
  • Note: In primates in general, paediatric forms of human drugs are useful in smaller individuals, to allow accurate doses to be administered appropriate for body weight. (B10.44.w44e)
  • For arterial samples the femoral artery can be used, or the radial artery on the ventromedial aspect of the carpus. (B336.39.w39)
  • Intramuscular injections can be given into the thigh, upper arm or chest muscle of great apes. (B214.3.1.w18)
  • Subcutaneous injections can be given over the thorax or back with the skin lifted slightly. (B214.3.1.w18)
    • Note: This is not a good site for giving large volumes, e.g. for administration of fluids. Apes have little subcuticular space in which to place fluids, and individuals with low circulating blood volume or acidosis will have constriction of blood vessels supplying the skin, therefore fluids given by this route will not be absorbed. Note that e.g. 5% dextrose is contraindicated because it will actually cause vasoconstriction and draw fluid out of the vessels into the subcuticular space. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
Associated techniques linked from Wildpro Birds

Cranes

Mammals

Bears

Rabbits (Lagomorphs)

Ferrets

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Supportive Care and Nursing

NOTE: This section should be read in conjunction with the information given in:
GENERAL
  • Supportive care is frequently a vital part of the treatment of individuals and groups and may vary from simple provision of a safe quiet place for recuperation to force feeding and parenteral fluid administration.
  • In all dealings with sick or injured wild animals minimizing stress is very important. It is important to remember that the proximity of humans and human-associated noises are likely to be extremely stressful to the animal. Every effort should be made to minimize this stress.
  • Provision of appropriate accommodation is an important part of supportive care. See:
  • Wild animals should be handled only when necessary for their treatment (B11.4.w17, V.w5). Sick dogs, cats and other pets frequently benefit from being fussed over and patted or stroked: wild animals generally do not. For most wild animals the best "tender loving care" which can be given is to be left alone. Handling should also be minimized to reduce the chance of a wild animal which is to be released back into the wild becoming unwary of humans, which could be dangerous both to the animal and to the people it may approach after release. (P19.1.w10)
WARMTH AND TEMPERATURE REGULATION
  • Provision of a warm environment will decrease the energy expenditure required for the animal to maintain its body temperature, avoiding hypothermia.
  • This is particularly important for:
    • Small animals, which have a large surface area to volume ratio and a high metabolic rate;
    • Neonates, especially of altricial species;
    • Individuals unable to behaviourally regulate temperature due to illness, or during anaesthesia and recovery from anaesthesia.
  • Note: It is also important to ensure that animals are not becoming hyperthermic. Hyperthermia is more likely to occur in:
    • Individuals which have lost some of their normal insulation (fur or subcutaneous fat) or in which fur is contaminated;
    • Individuals unable to behaviourally regulate temperature due to illness or during anaesthesia and recovery from anaesthesia.
    • Individuals being kept in a confined space.
    • See: Hyperthermia - Sunstroke - Heatstroke
FOOD AND WATER
  • Intake of sufficient food and water is vital. Within a given class of animals, smaller animals have a higher metabolic rate than larger animals. Birds have a higher metabolic rate than similarly sized mammals. In calculating nutritional requirements, whether for voluntary food intake or force-feeding, it is important to remember the requirements for healing wounds, in the case of injured animals, and for regaining weight which may have been lost in the period prior to admission. Monitoring of weight and body condition is essential to ensure that adequate nutrients are being ingested and absorbed. See: 
  • Fresh water should always be available, with due consideration for the risks involved when an animal is recovering from anaesthetic or is unable to lift its head properly due to illness.
  • Fluid therapy should be given to dehydrated individuals and those unable to drink voluntarily. Depending on the degree of dehydration and the general state of the animal, fluids may be given by the oral, subcutaneous, intravenous or intraosseous routes. See section below on this page: Fluid Therapy
  • Appropriate food should be provided. 
    • It is important to remember when dealing with wild animal casualties that the food available for feeding animals during hospitalization is frequently very different from what they would eat in the wild and may not be recognized as food, at least initially.
    • Force-feeding may be required if an animal cannot or will not eat voluntarily. This procedure is likely to be stressful to the animal and should be discontinued as soon as adequate voluntary intake develops. 

Waterfowl Consideration

This section should be read in conjunction with the information given in: "Accommodation Design for Birds: Temporary / Hospital Accommodation".
  • N.B. Waterfowl, particularly wild waterfowl, hospitalized for treatment are susceptible to the development of diseases such as Aspergillosis and Bumblefoot. Prevention of the development of such diseases in hospitalized waterfowl includes reducing stress and designing hospital accommodation to avoid or minimize known factors associated with their development. Particular care should be taken with species known to have high susceptibility to these diseases.
  • Species with strong pair-bonds (e.g. swans) may be stressed and pine if separated from their mate during treatment. If possible, it may be best to hospitalize both birds, keeping the pair together.
  • N.B. It is very important to maintain and if necessary restore the waterproofing of waterfowl during treatment. Provision of water for bathing and swimming is important in maintaining waterproofing.
  • Providing the opportunity for hospitalized waterfowl to spend much of their time on water is particularly important in species which normally spend most of their time afloat, such as swans, seaducks and stifftails, and for any individual recovering from a leg or foot problem, as it minimized the amount of time spent weight-bearing on the affected limb.
  • For most hospitalized wild waterfowl, food such as wheat, brown/wholemeal bread and green food should be provided, both dry and in a bowl covered with water. Consideration should be given as to the natural diet and nutritional requirements of the individual species and information from the "Natural Diet" "Feeding Behaviour" and "Aviculture" sections of the relevant species page should be consulted. (see also: Food and Feeding for Birds).
  • Tube feeding may be required if hospitalized waterfowl are not taking food voluntarily (see: Gavage / Tubing of Birds).
  • It is important to remember that waterfowl which have lost the normal insulation properties of their feathers, such as may be seen with Oiling, will have greatly increased calorific (energy) requirements.

(B11.4.w17)

Crane Consideration Supportive care and treatment should aim to minimise stress, prevent further injury to the crane, and minimise the risks of infection.
Hospitalisation
  • Hospitalisation reduced the risks of intraspecific aggression on the sick crane, and reduces the change for disease transmission. (B12.56.w14)
  • A sick crane may need to be removed from a group, due to the risk of it being attacked by other members of the group. (B12.56.w14)
  • With an established pair of cranes, it may be less stressful and therefore beneficial if the pain can be kept together. (B12.56.w14)
  • Supplemental heat may be needed to keep a sick crane at an environmental temperature of 29 - 30 C. (B12.56.w14) 30-32 C (B115.8.w4) This may be provided using a heater or heat lamp.
    • Chicks may be placed into an incubator for warmth. (B12.56.w14)s
    • For older cranes, it is good to provide a background temperature of 21 C (70 F)  then use a raised heat lamp to give one area of up to 30 C (85 F).  (B115.8.w4)
    • If a crane is recumbent it should be placed under the heat lamp but continuously monitored for signs of overheating until it is mobile and able to move away from the lamp if it wishes to do so. (B115.8.w4)
Slings
  • In the event of serious leg injury or severe debility preventing the crane from standing, use of a sling may be required. See Medical Management of Cranes in Slings
    • While maintenance of a crane in a sling produces its own problems, it is preferable to leaving the crane recumbent. (P138.18.w1)
      • Cranes in sternal recumbency are unlikely to eat and may flail around and further injure themselves. Prolonged recumbency also leads to circulatory problems(P138.18.w1)
      • Being in a more normal physical posture (in a sling rather than recumbent) may assist the crane's mental outlook and improve the cranes desire to recover. (P138.18.w1)
Nutritional support
  • High quality nutritional support is essential.  (B12.56.w14)
    • Many ill/hospitalised cranes may be suffering from nutritional deficiency and/or may eat less than normal, therefore need to be tube fed by gavage (Gavage / Tubing of Birds). This is essential in anorectic or starving cranes. (B12.56.w14) Cranes can lose weight very rapidly when ill if not eating enough. (B115.8.w4)
      • Note: Wild cranes entering rehabilitation are highly likely to be in poor condition, having been ill or injured for several days before being captured. Additionally, they may not eat when first in captivity, therefore nutritional support is essential. (J311.21.w1)
    • The initial food given to a severely emaciated crane should be a high carbohydrate, low protein formula, such as Emeraid I (usually given mixed 1:1 with water). (B115.8.w4)
    • Once the crane has taken several feeds and normal faecal production has developed, diets which are more complex and contain protein, fat and fibre can be given, e.g. Emeraid II mixed 1:1 with warm water, liquid human enteral products, or the crane's normal pelleted diet mixed with water and other nutritional supplements. (B115.8.w4)
      •  Appropriate food mixtures to use for gavage are provided in Food and Feeding for Birds - Convalescent diets / Nutritional support
    • Details of how to carry out tube feeding are given in: Gavage / Tubing of Birds
    • For adult cranes, a flexible feeding tube and a 50-60 mL catheter-tip syringe are needed, with up to 12 mL per kg crane bodyweight given, slowly, at one feed. (B12.56.w14)
    • For chicks, 2.5 - 6 mL syringes and smaller feeding tubes are needed.
      • Note: given the limitations of the internal diameter of the syringe tip and the feeding tube, large particles need to be removed from the food mixture using a fine sieve, to avoid blockage of the tube. (B12.56.w14)
  • NOTE: It is essential to calculate the nutritional requirements of the crane and the energy density of the diet being given, to ensure that the crane is receiving sufficient nutrients and to continue supplementary feeding until there is objective evidence (quantity of food eaten, weight gain by the crane) that it is eating enough by itself. (V.w5)
  • Lactobacillus products have been given to both adult cranes and chicks (7g or 1.4 teaspoons per kg body weight), aiming to restore normal gastrointestinal flora and improve digestion; it is not known whether this is effective. (B115.5.w6)
  • For fluid therapy see section below: Fluid therapy
Initial treatment of acute trauma/shock
  • Dexamethasone (2 - 8 mg/kg intramuscularly, intravenously or subcutaneously, once or twice daily) or Prednisolone (10-20 mg/kg) are indicated if the crane has suffered acute trauma and/or is in shock. (B115.8.w4)
  • Consider whether antibiotics should be given, ideally based on culture and sensitivity testing, but initially before information is available on the causative organism, initial treatment must be chosen empirically based on previous experience/expected bacterial culture results. (B115.8.w4)
Vitamin/mineral injections
  • Consider whether vitamins and/or minerals should be given as part of the supportive care. (B115.8.w4)
  • Vitamin A should be given in any case of bumblefoot or if a specific vitamin A deficiency is suspected. (B115.8.w4)
  • Iron in the form of iron dextran should be given in cranes with anaemia (pale mucous membranes, low haematocrit) or after haemorrhage. (B115.8.w4)
Bear Consideration This section should be read in conjunction with the information given in:
Fluids
  • Bears may require fluids as supportive treatment for bears which have diarrhoea and vomiting and/or are refusing food and water, (J3.145.w4, J11.83.w1, J142.19.w1), when obviously dehydrated (B16.9.w9, B64.26.w5, P62.18.w1),  or while undergoing prolonged surgical procedures. (P3.2006a.w1, P62.18.w1)
Temperature regulation
Food and water
  • Water should be accessible except in the eight hours prior to anaesthesia or if the bear may be in danger of drowning (e.g. during seizures).
  • For wild bears undergoing temporary captivity for rehabilitation, high-quality named-brand complete dry dog food or omnivore food may be used. (B468.8.w8p)
  • See: Food and Feeding for Mammals - Convalescent diets / Nutritional Support

Lagomorph Consideration

This section should be read in conjunction with the information given in: 
General
  • Housing

    • Rabbits are prey animals and their accommodation should be chosen with this in mind, well away from the site and sound of predators including cats, dogs, ferrets, birds of prey etc.

    • A quite place away from the sounds and scents of predators is particularly important for critically ill rabbits. (J213.1.w1)

    • For wild lagomorphs, accommodation should also be away from normal human activities.

    • Non-slip flooring is important.

    • Hay is a familiar bedding in which rabbits can burrow to feel secure.

    • For recumbent rabbits, synthetic fleece and incontinence pads are useful and give some protection against urinary and faecal soiling. (B601.3.w3)

    • If possible, have the owner bring a familiar toy, piece of clothing etc. to leave with the rabbit. (B601.3.w3)

    • See: Accommodation Design for Mammals - Temporary / Hospital Accommodation

  • Hospitalised rabbits should be weighed on arrival; this allows monitoring of weight loss or gain. 

  • Rabbits should be assessed for for pain - see Physical Examination of Mammals - Observation
    • Reluctance to move, hunched posture, anorexia, teeth grinding, self-trauma, raised body temperature, increased respiratory rate, excessive or lack of drinking, head elevation or extension, pushing the abdomen onto the floor, unusual aggression, an anxious facial expression, reduced faecal output and occasionally vocalisation may all be signs of pain. (B601.3.w3, B534.43.w43f)
    • Careful observation is needed to detect subtle signs of pain. (B600.5.w5)
  • Monitor and record food and water intake, urination, faecal output and general observations. (B601.3.w3)

  • Check the perineal area of rabbits which are recumbent or unable to groom properly. Gentle washing and careful drying is needed if the rabbit is soiled with urine or faeces. (B601.3.w3) See: Bathing Rabbits

  • If possible, allow daily exercise; this is generally beneficial and assists in gut motility. (B601.3.w3, P113.2005.w3)

    • Keep the rabbit under observation when it is out of its cage, and make sure no electrical wires are accessible. (B601.3.w3)

Temperature regulation
  • Rabbits are generally more prone to overheating than to getting chilled. However, this may change if, for example, a large area has been clipped and prepared for surgery.
  • Rabbits are particularly prone to hypothermia in the perioperative period.
  • Heat lamps can be used, with care. (J15.23.w6)
  • Warmed intravenous fluids can be used. (J15.23.w6)
  • Temperature-controlled incubators can be used. (J15.23.w6)
  • For shock-induced hypothermia:
    • Place the rabbit on a water-circulating heat pad and cover the rabbit with a towel to reduce heat loss. 
    • Avoid heat lamps: rabbits have thin skin which easily burns. (J213.1.w1)
  • Hyperthermic rabbits pant rapidly; they have a body temperature over 41 C (106 F) and are hypovolaemic. (J213.1.w1)
Food and water
  • Fluid and electrolyte imbalances should be corrected first; nutritional support should then be given. (J213.10.w1)
  • Water should be accessible at all times unless there is risk of the rabbit drowning in it.
    • It is important to provide a hospitalised rabbit with the same type of water source as it has at home, where that is a bowl or a bottle, as otherwise it may not drink.
    • If using a bowl, a heavy (e.g. earthenware) bowl is recommended to reduce the risk of the rabbit tipping it up.
  • Food should always be available. Provide food which the rabbit is familiar with while it is hospitalised - the same brand, even in the same bowl. (J213.1.w1, J213.10.w1)
  • Good quality hay (e.g. Timothy hay) should always be available. (J213.10.w1)
  • Green foods may encourage eating. Offer meadow hay, fresh grass, dandelion leaves, broccoli, dark green lettuce, watercress, parsley, green cabbage, kale etc. (B601.3.w3, J213.10.w1)
  • Consider use of probiotics and multivitamins, particularly B vitamins. (B601.3.w3)
  • Encourage exercise to stimulate gastro-intestinal motility. (P113.2005.w3)
Anorexia in Rabbits - causes, effects and treatment
Causes of anorexia

Anorexia is common in rabbits, associated with:

  • Stress and anxiety (e.g. loss of companion, use of an Elizabethan collar);
  • Pain including dental and oral pain which make eating painful, but also any injury, arthritis or surgery;
  • Management (e.g. poor food quality, change in food)
  • Fractured jaw or other oral pr0blem physically preventing the rabbit from eating
  • Systemic disease
    • Renal failure
    • Liver disease
    • Neoplasia
    • Myxomatosis
    • Viral haemorrhagic disease
    • Pasteurellosis
    • Enterotoxaemia
    • Lead poisoning
  • Dental and oral disease
  • Gastro-intestinal diseases
Effects of anorexia
  • Effects of anorexia are varied and show complex interactions including: (B600.3.w3, J15.24.w3)
    • Negative energy balance, leading to:
      • hepatic lipidosis (often fatal) (Hepatic Lipidosis in Rabbits);
        • Obese rabbits are at greater risk.
      • ketosis, which leads to dehydration and electrolyte imbalances, which in themselves can cause anorexia, and may be fatal.
    • Reduced intake of dietary fibre, which:
      • reduces substrate available for caecal fermentation (causing negative energy balance) and 
      • reduces gut motility. 
    • Reduced gut motility which:
      • slows gastric emptying (which can cause anorexia) (Gastric Stasis Syndrome in Rabbits);
      • reduces substrate for caecal fermentation (causing negative energy balance);
      • leads to accumulation of gas in the GIT;
      • may promote gastric ulceration (Gastric Ulceration in Lagomorphs ) (which may be painful).
      • leads to dehydration and electrolyte imbalances (which both cause anorexia and may be fatal).
    • Gas accumulation (associated with decreased gut motility); this causes GIT distension, which is painful, resulting in anorexia and gastric ulceration (which may be painful).

    (J15.24.w3)

Treatment of anorexia

If the rabbit does not eat voluntarily then nutritional support is essential both to provide for the rabbit's protein and energy needs and to maintain (or promote) normal gastro-intestinal function.

Rehydration

  • Often the anorectic rabbit will be dehydrated and rehydration is needed. See: Fluid therapy section below

Assisted feeding

  • Various foods can be given in assisted (syringe) feeding, including pelleted complete rabbit diets ground up and mixed to a slurry with water, liquidised vegetables (J15.13.w7) and proprietary products designed for assisted feeding of rabbits or other small herbivores.
  • Ideally, the rabbit should be given a food which includes the indigestible fibre needed for normal gut function. Unfortunately, this may not be possible when feeding through an endotracheal tube, and may be difficult when feeding through a syringe with a standard tip; a syringe with a wide-bore tip should be used if available. (B601.3.w3, J213.10.w1)
  • Note: many rabbits resist syringe feeding. If owners are to carry out syringe feeding it is important to teach them proper rabbit restraint. (P113.2005.w2)
  • Note: Tempting foods should always be available while a rabbit is on assisted feeding. (J213.10.w1)
  • Probiotics, vitamins (especially B-vitamins) and even transfaunation - by feeding a rabbit with caecoliths from a healthy rabbit - may be useful. (B601.3.w3)
  • Avoid high carbohydrate or high fat nutritional supplements. (B609.2.w2)

Calculating food requirements

  • It is important to calculate the rabbit's energy requirements to determine the amount to be fed daily:
  • The rabbit's approximate basal energy requirements (BER) or Basal Metabolic Rate (BMR) can be calculated as follows:
    • 70 x (bodyweight in kg)0.75 = Basal Metabolic Rate (BMR) kcal/day. For a 2.0 kg rabbit, BMR = 70 x (2.0)0.75 = 117.7 kcal/day. (B192, J213.10.w1)
    • The actual amount required will vary depending on the rabbit's state of health and may be from 1.2 x BMR to as high as 2.0 x BMR. During healing, the rabbit should be hypermetabolic and need more food. However a rabbit which has been starved or stopped eating due to gastrointestinal problems will be hypometamolic and have reduced energy requirement. (J213.10.w1)
    • Equations are available which indicate the energy requirement for animals in different circumstances; in most circumstances the amount of energy required per day is greater than the basal energy requirement (BER): (B192)
        • Growth: 1.5-2.0 x BER 
        • Enclosure rest: 1.25 x BER
        • Following starvation: 1.25 x BER
        • Post-surgery: 1.25 x BER
        • Severe burns: 1.5 - 2.0 x BER
        • Sepsis: 1.5-2.0 x BER
        • Trauma: 1.5 x BER
        • Neoplasia (cancer) 1.5 x BER
        • Hepatic (liver) disease: 1.25 x BER
        • Severe renal (kidney) disease: 1.25 x BER
      • (B192)
    • For a debilitated rabbit, start feeding at 40 - 70% of calculated daily energy requirement and increase to 100% over three to five days. (J213.10.w1)
    • For a rabbit in good condition, start at 75 - 100% of calculated daily energy requirement. (J213.10.w1)
    • Note: Once the caloric requirement has been calculated (kcal/day), it is necessary to calculate the required amount of the food which is to be given. (J213.10.w1) This will vary depending on the food used. Use of a proprietary formula is recommended. Formulae designed for assisted feeding have a known caloric content.
    • When an enteral formula is provided as a dry powder, the amount needed should be calculated for the powder, before adding water, since the amount of water added may vary. (J213.10.w1)
    • The required amount should be given divided into several feeds over the day (24 hours). (J213.10.w1)
    • e.g. 10-15 mL/kg of Critical Care for Herbivores orally every 6 to 8 hours. Larger and more frequent feeds may be accepted by the patient- feed as much as the patient will accept. (B609.2.w2)

Methods of feeding

Medication

  • Consider using gut motility enhancers.
  • Give appropriate pain relief. (P113.2005.w4)
  • Cholestyramine may be used to treat/prevent enterotoxaemia, by binding with bacterial endotoxins. (P113.2005.w4)
  • Probiotics may be useful. (P113.2005.w4)
  • Antibiotics may be needed in the treatment of gastrointestinal infections. (P113.2005.w4) See:
  • Nandralone 2 mg/kg subcutaneously or intramuscularly may be given to stimulate appetite. (J15.24.w4, B539.1.w1)

Additional care

  • Make sure the rabbit is away from stressors such as loud noises, predator species (cats, dogs etc.). (P113.2005.w4)
  • Provide the opportunity for exercise, which can help stimulate gut motility. (P113.2005.w4)
  • Monitor food intake (assisted, voluntary), production of faecal pellets, weight and general health. (P113.2005.w4)
Ferret Consideration This section should be read in conjunction with the information given in: 

In general, ferrets should be hospitalised for as short a time as possible. Although relatively resilient, they may become stressed by barking dogs etc. (J29.6.w3)

Ferrets should be housed away from rabbits, rodents etc. as their presence may be stressful to the prey species. (B631.18.w18)

Feeding

  • Ferrets should be provided with the same diet they are used to, while they are hospitalised, as they may refuse different food. (B602.2.w2)
  • If necessary, a highly palatable ferret diet, or a premium-quality cat/kitten food can be given. (B602.2.w2)
  • If changes are to be made to the diet, these should preferably be started after the ferret has returned home, and should be made gradually. (B602.2.w2)

Assisted feeding

  • An anorectic ferret can be syringe-fed with a palatable, liquid nutritional product suitable for carnivores. (B602.2.w2, J29.19.w1) e.g. Hill's canine/feline a/d, or meat-based baby food can be used. (J15.24.w5)
  • Feed three to four times a day. (J29.19.w1) Give 12 mL at a time, 3-4 times a day. (J15.24.w5)
  • Use of a nasogastric tube is not recommended. The small nostrils make placement of a tube difficult, and the smalll tube diameter would prevent most diets passing through the tube. (J29.19.w1)
  • Pharyngostomy tubes are useful for ferrets which cannot swallow or will not accept syringe feeding. (J29.6.w3, P120.2006.w6)
    • See: Pharyngostomy Tube Placement in Ferrets
    • Consider the additional physiological stress associated with general anaesthesia to place the tube. Additionally, it is not easy to keep the tube properly bandaged on the neck. (J29.19.w1)
  • If a ferret cannot tolerate any food by the gastrointestinal route (e.g. with a malabsorptive diarrhoea), partial parenteral nutrition can be given intravenously. This supplies lipids, amino acids, dextrose, electrolytes, vitamins and minerals as well as fluid. Give using an infusion pump through a polyurathane or silicone elastomer intravenous catheter into the jugular vein. Maintenance of sterility is problematic. (B602.2.w2, P120.2006.w6)
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
Associated techniques linked from Wildpro

Rabbits

Ferrets

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Wound Management

  • Get all necessary equipment ready before starting treatment. (P62.23.w1)
  • When treating wild animals, maintain a warm, quiet environment. (P62.23.w1)
  • Ensure sufficient personnel are available for appropriate restraint and treatment, (P62.23.w1) or use chemical restraint. (V.w5)
  • In general, treatment of wounds is likely to require sedation or general anaesthesia of the animal.
    • This is particularly true if extensive cleaning and debridement (surgical removal of dead and severely damaged tissue) is necessary.
  • Note: the stress and pain involved in wound management must be remembered: just because it is possible to hold a conscious animal of a particular species sufficiently immobile for wound management to take place does not mean that treatment of the conscious animal without sedation and analgesia is appropriate.
  • Treat shock and dehydration first, before treating any wounds.

(J213.9.w4, P62.23.w1, V.w5, V.w6, V.w26)

Initial inspection and cleaning
  • Careful inspection should be carried out for the presence of fly eggs or maggots (which may not be superficially visible) and action taken to remove these. See: Myiasis.
  • Maintain sterile technique while cleaning and managing the wound, even if it is already infected. (P62.23.w1)
    • This reduces exposure of the wound to additional and nosocomial (hospital-associated) pathogens. (P62.23.w1)
  • If wound are superficial and mild, with minimal contamination and no infection, minimise clipping of hair (or feathers, in birds) to preserve insulation, and manage by simple first aid. (P62.23.w1)
  • Control any bleeding. (J213.9.w4)
  • Take samples for culture, sensitivity and a Gram stain. (J213.9.w4)
Hair clipping and skin preparation
  • Clip the area around the wound to allow full evaluation of the area affected, and to prevent additional contamination of the wound. (P62.23.w1)
  • Clipping of hair around the wound should be carried out using curved, blunt-ended scissors. If these are damped or dipped in mineral oil, cut hare will stick to the blades rather than fall into the wound. (J15.18.w3)
  • Further from the wound, clippers can be used; these must be sharp and whole, without any missing teeth, to minimise further trauma to the skin. (J15.18.w3)
  • Sterile swabs, a water-soluble jelly (e.g. K-Y Jelly, Johnson & Johnson) (J15.18.w3) or moist cotton wool may be placed in/over/along the edge of the wound to minimise clipped hair contaminating the wound by falling into it. (P19.2.w5)
    • Large wounds might be temporarily closed with towel clips or a continuous suture while the hair is being clipped. (J15.18.w3)
  • Note: The area clipped should not be excessive, as hair normally provides the animal with protection from cold, some trauma etc. Loss of hair from a large area will increase the risk of the animal becoming chilled, particularly in small animals. (P19.2.w5, V.w5)
  • Surgically prepared the skin around the wound using povidone-iodine or chlorhexidine diacetate. (P62.23.w1)
Lavage
  • For contaminated wounds, thorough flushing with an isotonic solution such as sterile normal (0.9%) saline or lactated Ringer's solution is recommended. (J15.18.w3, J213.9.w4, P19.2.w5, P62.23.w1,)
    • Large volumes of lavage/irrigation solution help to dilute contaminating bacteria in the wound. (P62.23.w1)
    • A substitute saline solution [not a precise substitute] may be produced if necessary by dissolving one teaspoon of salt in a pint of water (preferably boiled and cooled). (B337.A6.w12, P19.2.w5)
    • For grossly contaminated wounds, tap water directed using a hand shower head can be used for initial lavage, but it should be followed by use of physiological saline to give an appropriate environment for healthy tissue. (J15.18.w3, P62.10.w2, P62.23.w1)
    • Optimum pressure for lavage of wounds is 8 psi. This can be carried out by using an 18-gauge needle attached to a 20 or 35 mL syringe. (J15.18.w3, J213.9.w4, P62.23.w1)
      • Higher pressures should not be used as they may cause deeper contamination, or oedema in adjacent, undamaged, tissues. (J15.18.w3)
    • During lavage, make sure the wound can drain freely and that the patient is protected from the fluids, not becoming soaked. (J15.18.w3)
  • Contaminated or infected wounds should be cleaned using a non-irritant antiseptic solution. 
    • If an antiseptic solution is used, it is necessary to balance the beneficial anti-bacterial effects with the potential toxic effects of the antiseptic on tissue. (J15.18.w3, P62.23.w1)
    • Chlorhexidine can be used at a 0.05% final solution. This has good residual activity, but Staphylococcus aureus is often resistant. It is toxic to fibroblasts. (J15.18.w3, P62.23.w1)
    • Povidone iodine (Iodophors) can be used as a 1% solution; this is broad spectrum, but inactivated by debris, pus or blood. It does not have as good residual activity as chlorhexidine and is toxic to host cells. (J15.18.w3, P62.23.w1)
    • Hydrogen peroxide, cetrimide/chlorhexidine (Savlon Veterinary Concentrate, Mallinckrodt) and hypochlorite (Dakin's solution) have all been shown to be irritant and highly toxic to host cells and should not be used. (J15.18.w3)
    • Note: use of products such as Dettol and TCP should be avoided; they are irritant and sting severely on open wounds. (P19.2.w5)
  • Do NOT cleanse deep wounds with hydrogen peroxide or alcohol, as this may cause tissue injury and increase the chance of infection developing. (D249.w13)
    • Hydrogen peroxide does not have any antibacterial properties, and it injures capillary beds. It should not be used for cleaning wounds. (P62.23.w1)
    • Avoid using alcohol; this may cause pain and also cool the animal excessively. (P62.23.w1)
Surgical debridement
  • This aims to remove foreign material and devitalised, contaminated and infected tissue, reducing the need for debridement of the wound by macrophages and thereby allowing rapid onset of the proliferative phase of wound healing, as well as helping to control infection. (J15.18.w3)
  • Considerable debridement of wounds may be necessary to remove contaminated and devitalised tissue. Anaesthesia will often be necessary for this process as it will often be appropriate to remove the damaged tissue as far back as to where there is an effective blood supply (and thereby usually pain sensors) to encourage healing.
  • If the wound is contaminated but not infected (in the first six hours, the "golden period"), it can be debrided once to give a surgically clean wound; infected wounds require further stages of debridement. (J15.18.w3)
  • During surgical debridement, the wound should be draped and prepared as for surgery. Viable tissue must be handled gently; handle skin edges with skin hooks (bent 20-gauge needles can substitute), not retractors. (J15.18.w3)
  • Ensure haemostasis to avoid development of a haematoma, but note that multiple ligatures of necrotic tissue plus from electrocautery can delay healing. (J15.18.w3)
  • Explore the wound carefully; check there are no deep punctures complicating an apparently simple surface laceration. (J15.18.w3)
  • Handle different tissue types appropriately: (J15.18.w3)
    • Trim the skin edges; note that initial vascular spasm may cause mistakes in assessment of skin viability. Preserve as much skin as possible.
    • Debride muscle which is dark, friable, or does not contract.
    • Debride exposed fatty tissue back to a clean plane.
    • Conserve and protect nerves whenever possible.
    • Preserve tendons as much as possible. Note that anastomosis will fail if infection is present, and that strength will begin to develop only after three to five days.
    • Exposed joints should be lavaged thoroughly, repaired and immobilised.

    (J15.18.w3)

(B13.16.w11, B14, J15.18.w3, P19.2.w5, P62.23.w1, V.w5, V.w26, V.w6)

Suturing
  • Puncture wounds should never be sutured.
  • Suturing (primary closure) may be appropriate with fresh lacerations or with older lacerations if the tissue deficit following debridement is not too extensive.
    • In wild animals, absorbable sutures should be used for closure of the skin as well as deeper tissues, so that there is no need for additional handling to remove the sutures. 
      • It is particularly important to use absorbable sutures in field situations when the animal will be released immediately. (B345.4.w4)
      • Use a tapered needle to suture internal muscle layers on a deep wound. (B345.4.w4)
      • Use a cutting needle to suture the skin. (B345.4.w4)
    • Consideration should be given to wound drainage; the placement of a drain may be required (not in the field).
    • Care must be taken to avoid attempting to suture wounds with a large tissue deficit which would place excessive pressure on the wound.
  • Tissue glue or bandage strips can also be used to close clean fresh wounds. (P62.23.w1)
Delayed primary closure
  • Wounds which are grossly contaminated, infected (contain pus), contain necrotic tissue or involve defects which wound produce excessive tension at the wound edges cannot be closed.
  • Control of infection and management of the wound with bandaging may allow surgical debridement and delayed primary closure three to five days later. (P62.23.w1)
Encouraging healing by secondary intention
  • In many cases (e.g. old, contaminated wounds) it may be necessary to leave the wound to close by secondary intention.
  • These wounds are kept open and managed to promote the establishment of a healthy bed of granulation tissue. (P62.23.w1)
  • Granulation tissue should develop at the wound edges and progress across the wound, with contracture and epithelialisation. (P62.23.w1)
  • The application of topical preparations that encourage epitheliogenesis (stimulate healing) may be useful, e.g. Intrasite Gel (Smith and Nephew).
  • Note: healing by secondary intention is slower, often with more scarring and loss of skin pliability. (P62.23.w1)
  • Where possible, the use of dressings which promote healing may be used.
    • Note: Many wild animal casualties, particularly adult mammals, may not tolerate dressings and bandages.
Honey
  • Honey is recognised to be beneficial applied topically to infected wounds:
    •  The high osmolarity of honey and its consequent and ability to minimise water available to bacteria gives an antibacterial action.
    • Some types of honey such as manuka honey, have a slow, sustained production of hydrogen peroxide at very low concentration, producing an antimicrobial effect.
    • Some types of honey also have other components with demonstrable antimicrobial effects.
    • The low pH and high glucose content also may be stimulatory to macrophages.

    (J128.14.w1)

Antibiotics
  • All wounds in wild animals should be considered to be contaminated and appropriate antibiotic treatment instigated.
    • In the field, commonly, penicillins are given, since these are effective against many of the microbes found on skin (and likely to contaminate wounds) and are available in long-acting preparations. (B345.4.w4)
    • When giving a single dose of procaine penicillin/benzathine penicillin, give 22,000 IU/kg of the benzathine penicillin G to ensure an adequate repository effect giving antibiotic cover for 5-7 days. (B345.4.w4)
    • Give no more than 5 mL at any one site, subcutaneously or into the large muscle masses of the hind legs. (B345.4.w4)
  • With "cat-caught" puncture wounds it is particularly important to ensure that antibiotics are likely to be effective against Pasteurella multocida.
  • Once culture and sensitivity results are available, appropriate antibiotics should be given.

(B345.4.w4, J15.18.w3, J213.9.w4, V.w5, V.w6, V.w26)

Bandaging
  • If properly applied, bandages work "to provide an optimal environment for epithelialization and wound contraction with the fewest complications."  (P4.1990.w2). The functions of bandaging are to provide:
  • Pressure to reduce dead space, swelling, oedema and haemorrhage.
  • Protection from pathogenic microorganisms.
  • Immobilization of the wound.
  • Protection from desiccation.
  • Protection from self mutilation or abrasion.
  • Absorption of exudate.
  • Debridement of the wound surface.
  • Comfort for the animal.
  • Three layers of bandaging are usually used, the different layers having different functions.

PRIMARY / CONTACT LAYER:

This is the most important layer for wound healing and may be adherent or non-adherent. The contact layer should be sterile, remain in place despite patient movement, provide a moist environment for the wound, be comfortable, and assist in debridement.

a) Adherent dressings, used in the inflammatory stage of wound healing

  • Dry-to-dry dressings (open weave or fine mesh gauze pads) may be useful for the initial phase of treatment where excessive amounts of necrotic debris need to be removed by a process other than surgical debridement, or for wounds in which there is excessive exudate production. 
  • Wet -to-dry dressings, using sterile gauze pads soaked in saline or dilute disinfectant (e.g. 1:40 chlorhexidine diacetate), may be used to mechanically remove exudate and necrotic tissue.
    • Application is more comfortable if the solution used to wet the dressing is warmed first.
    • These are useful for wounds with loose necrotic matter, containing foreign material and producing viscous exudate.
  • Adherent dressings disrupt the surface of the healing wound at each dressing change.
  • The very moist environment produced by wet-to-dry dressings may lead to tissue maceration.
  • Felt-to-gel dressings - calcium alginate pad
    • This is strongly hydrophilic and useful for wounds with moderate to heavy exudate production.
    • This is a nonwoven, felt-like pad when applied. As it absorbs fluid, calcium in the dressing is exchanged for sodium in the wound fluid; calcium alginate changes to a sodium alginate gel. 
    • The gel traps bacteria and these are then lavaged away with the gel at bandage change.
    • Formation of granulation tissue may be enhanced.
    • Not suitable for use on wounds with exposed muscle, tendon or bone.
    • If the wound is not producing enough fluid to convert the calcium alginate to sodium alginate, it can form a hard, difficult to remove, eschar over the wound. 

b) Non-adherent dressings used in the granulation and epithelialization phase of wound healing.

  • Traditional non-adherent dressings - cotton dressings with a non-adherent film, and petroleum-impregnated fine-mesh gauze dressings:
  • These are widely available and inexpensive.
  • Allow excess fluid to be absorbed into the secondary layer.
  • Cotton non-adherent film dressings in practice do stick to wounds if left in place for more than 2-3 days; as a result when the dressing is removed disruption of the healing surface and bleeding may occur.
  • Petroleum-impregnated fine-mesh gauze dressings can slow the rate of wound epithelialization.
  • Fine-mesh gauzes can also be impregnated with polyethylene glycol. This is bland, non-toxic, non-irritating, watersoluble and hydrophilic, and provides a nonadherent bandage which does not interfere with epithelialization. (B534.3.w3a)
    • Being hydrophilic, this assists in drawing fluid up through the contact layer to the secondary layer, for absorption. (B534.3.w3a)
  • These dressings all may slip from the wound despite careful bandaging.
  • Modern non-adherent dressings have been developed which keep the wound surface moist, prevent scab formation and increase the rate of re-epithelialization (in comparison to traditional dressings):

i) Occlusive hydrocolloid or hydroactive dressings.

  • These dressings adhere to skin but not to wounds, are semi-flexible, opaque, and impermeable to moisture vapour and oxygen.
  • They absorb fluid and exudate, producing a moist gelatinous cover over the surface of the wound. 
  • Despite their adhesive qualities, additional bandaging is usually required to hold these dressings in place.
  • May leave a slightly sticky residue on skin and feathers.
  • Some hydrophilic dressings may be sufficiently rigid to be sutured lightly into position, and may assist granulation of quite large areas.
  • Dressing should be changed every 2-3 days initially, once a week when a healthy granulation bed has been established.
  • Dressing must be changed if quantity of exudate results in leakage around the edge of the dressing (to prevent bacterial invasion).
  • Useful for e.g. slow-healing, granulating wounds over the keel and carpal joints, and for granulating bumblefoot lesions, also for extensive wounds with considerable exudate production and wounds requiring debridement.

ii) Semi-occlusive moisture-vapour permeable dressings.

  • Thin, transparent flexible polyurethane membrane.
  • Stick to clean, dry, detergent-free skin but not to the wound.
  • May be conformed to even difficult-to-bandage areas.
  • Permeable to oxygen and moisture vapour.
  • Impermeable to water and bacteria.
  • Allow fluid and exudate to accumulate under the dressing.
  • Maintain a moist, aerobic environment
  • Maintain a sterile surface (if wound aseptically cleaned beforehand) while margins remain sticking to surrounding skin.
  • Prevent desiccation and scab formation, reduce pain associated with the desiccation of nerve endings
  • Promote leukocyte debridement of the wound surface, and migration of epithelial cells from the wound edges (epithelialization), and thereby speed healing.
  • Allow visual monitoring of the wound and both qualitative and quantitative assessment of the production of exudate.
  • Dressing should be changed every 2-3 days initially, once a week when a healthy granulation bed has been established.
  • Dressing must be changed if quantity of exudate results in leakage around the edge of the dressing (to prevent bacterial invasion).
  • Use should be discontinued if excessive redness, swelling and/or odour indicates gross infection.

SECONDARY / MIDDLE LAYER:

Functions:

  • absorption of fluids and wound exudate
  • protection of the wound from additional trauma 
  • immobilization of the wound while healing occurs

Conforming gauze bandages are usually used as the secondary bandage layer. Other materials include e.g. Sof-Band Bulky Bandage, Johnson & Johnson. Joint damage, vascular compromise and delayed healing may all be seen if bandages are improperly applied.

  • The secondary bandage layer should have a random fibre pattern to capillary action and absorption is maximised.
  • It is important to change the bandage before this layer becomes completely saturated. (B534.3.w3a)
    • If moisture saturates this layer and penetrates to the outer layer, this allows contamination by exogenous bacteria. (B534.3.w3a)

TERTIARY / OUTER LAYER:

Function: to hold the bandages in place and to immobilise the wounded area. May also be used to protect the bandages from the attentions of the patient 

  • Surgical adhesive tape is commonly used, or self-adhesive bandage.
  • Porous tape allows evaporation of fluid, but if the bandage gets wet from the outside, bacteria can move inward by capillary action and contaminate the wound.
  • Waterproof tape provides protection for external fluid, but make the bandage occlusive and may lead t tissue maceration. 
  • Elastic adhesive tapes (e.g. Vetwrap bandaging tape, 3M Co, or Conform stretch tape, KenVet Animal CareGroup) provide pressure and conformation as well as immobilization.
  • When maximum absorption is needed (for a wound which is draining considerable amounts of fluid), it is important that the tertiary layer is only tight enough to hold the other layers in place and keep the layers in contact with each other.
  • A tertiary layer which is too tight can compress the intermediate layers, which may reduce absorption, impede blood supply to the tissues and impair contraction of the wound.
  • A tertiary layer which is too loose leads to insufficient contact between the primary and secondary layers, so that fluid can accumulate over the wound, which can lead to tissue maceration.
  • Placing a final piece of adhesive tape partly on the bandage and partly on the patient's skin prevents bandage slippage.

(P4.1990.w2, B11.3.w10, B13.16.w1, B14B116.30.w3, B534.3.w3a)

Waterfowl Consideration

  • There may be a conflict between the indications for bandaging a wound and the requirement of the birds for access to water for swimming.
  • If wound management precludes constant access to water, the possibility of daily access to clean water prior to application of a new bandage should be considered.

(B11.36.w4, B13.19.w12, V.w5)

WOUND MANAGEMENT FOR BIRDS

GENERAL PRINCIPLES

It is important when managing wounds to recognize that a variety of factors may impede wound healing including:

  • Severe protein deficiency.
  • Chronic anaemia.
  • Dehydration.
  • Poor nutritional status.
  • Presence of necrotic tissue (may physically impede migration of epithelial cells, and may harbour bacteria).
  • Presence of blood clots (may physically impede migration of epithelial cells, and may harbour bacteria).
  • Presence of foreign bodies, including e.g. non-viable bone as well as dirt etc.
  • Sutures causing foreign body reaction (minimized by selection of appropriate suture material).
  • Tissue destruction (due to desiccation, severe trauma or poor surgical technique).
  • Poor vascular supply.
  • Lack of immobilization of wounds over joint surfaces.
  • Continual abrasion.
  • Infection by pathogenic bacteria.
INITIAL ASSESSMENT OF SOFT TISSUE WOUNDS
  • Take the history and perform a general physical examination.
  • Take care to locate wounds, e.g. by parting the feathers.
  • Note the location and extent of the injury.
  • Estimate / record the age of the injury (e.g. skin discolouration due to bruising develops after 2-3 days and may persist for a week or longer.
  • Note any associated fractures or luxations.
  • Check blood supply and nervous supply to the affected area, particularly for wounds of the limbs.
TREATMENT OF FRESH UNCOMPLICATED WOUNDS
  • May be treated by primary closure to produce first intention healing. (B13.40.w13, B14, P4.1990.w2)
  • Haemorrhage must be controlled, e.g. by direct pressure. (B14)
  • Primary closure should not be attempted on open contaminated wounds (B13.40.w13).
  • N.B. Puncture wounds should not be sutured due to the risk of bacterial contamination.
TREATMENT OF OLDER AND/OR UNCONTAMINATED WOUNDS

These should be managed to allow secondary intention healing. Once infection has been controlled and a healthy granulation bed established it may be appropriate to suture the wound in some cases.

1) WOUND PREPARATION: Aim is to remove foreign material, devitalized tissues and potentially pathogenic microorganisms.

  • Carefully pluck feathers, or trim feathers with fine sharp scissors to avoid tearing skin, to produce a 2-4 cm healthy feather-free area of skin around the wound.
    • Plucking will encourage regrowth of feathers; if feathers are cut they will not regrow until the next normal moult. The minimum area should be plucked and great care is required to avoid tearing the skin.
    • N.B. plucking of feathers is painful; this may be best carried out on an anaesthetised bird if more than a few feathers are to be plucked.
    • N.B. Care should be taken not to damage the feather follicles and thereby prevent proper regrowth of feathers. This is imperative for the flight and tail feathers of birds of prey, and any other species with a high dependency on flight such as swifts and swallows. If there is any doubt, such important feathers should not be plucked until absolutely necessary (which could be due to damage to blood feathers or the proximity of physical damage). (V.w6)

    (B13.16.w11, B14, P19.2.w5, V.w5, V.w26)

  • Gently irrigate with warm water or sterile isotonic saline to remove debris, blood clots and gross contaminants.
  • N.B. in warm months check carefully for fly eggs/maggots: Myiasis may lead to large quantities of soft tissue being consumed in just a few hours.
  • Take samples for bacterial culture after removal of surface contaminants but before application of any antiseptics if bacterial infection is suspected.
  • Lavage with 0.05% chlorhexidine diacetate solution or 0.5-1.0% povidone iodine solution, will provide antibacterial activity
  • Hydrogen peroxide may be used for initial cleaning of dirty wounds, or as a sporicide if clostridial infection is suspected.
  • Surgically debride non-viable and necrotic tissue, until viable, vascularized tissue is visible. (N.B. may have to debride several times over a period of days with old or complicated wounds).
  • Achieve haemostasis.
  • It is important to minimise the area of feathers removed when treating birds as these provide the bird with its protection against weather and water and loss of feathers may delay release until the feathers regrow. (P19.2.w5, V.w5)

2) TOPICAL MEDICATION:

3) BANDAGING:

  • If properly applied, bandages work "to provide an optimal environment for epithelialization and wound contraction with the fewest complications."  (P4.1990.w2). The functions of bandaging are to provide:
  • Pressure to reduce dead space, swelling, oedema, haemorrhage.
  • Protection from pathogenic microorganisms.
  • Immobilization of the wound.
  • Protection from desiccation.
  • Protection from self mutilation or abrasion.
  • Absorption of exudate.
  • Debridement of the wound surface.
  • Comfort for the bird.
  • Three layers of bandaging are usually used, the different layers having different functions.

PRIMARY / CONTACT LAYER:

This is the most important layer for wound healing and may be adherent or non-adherent. The contact layer should be sterile, remain in place despite patient movement, provide a moist environment for the wound, be comfortable and assist in debridement.

a) Adherent dressings

  • open weave or fine mesh gauze pads may be useful for the initial phase of treatment where excessive amounts of necrotic debris need to be removed by a process other than surgical debridement, or for wounds in which there is excessive exudate production. Wet -to-dry dressings, using sterile, saline-soaked gauze pads may be used to mechanically remove exudate and necrotic tissue.
  • Adherent dressings disrupt the surface of the healing wound at each dressing change, and the very moist environment produced by wet-to-dry dressings may lead to tissue maceration.

b) Non-adherent dressings are used in the granulation and epithelialization phase of wound healing.

  • Traditional non-adherent dressings - cotton dressings with a non-adherent film, and petroleum-impregnated fine-mesh gauze dressings:
  • These are widely available and inexpensive.
  • Allow excess fluid to be absorbed into the secondary layer.
  • Cotton non-adherent film dressings in practice do stick to wounds if left in place for more than 2-3 days; as a result when the dressing is removed disruption of the healing surface and bleeding may occur.
  • Petroleum-impregnated fine-mesh gauze dressings are not ideal for use in birds as they soil the feathers; work in dogs also indicates they can slow the rate of wound epithelialization.
  • Both types may slip from the wound despite careful bandaging.
  • Modern non-adherent dressings have been developed which keep the wound surface moist, prevent scab formation and increase the rate of re-epithelialization (in comparison to traditional dressings):

i) Occlusive hydrocolloid or hydroactive dressings.

  • These dressings adhere to skin but not to wounds, are semi-flexible, opaque, and impermeable to moisture vapour and oxygen.
  • They absorb fluid and exudate, producing a moist gelatinous cover over the surface of the wound. 
  • Despite their adhesive qualities, additional bandaging is usually required to hold these dressings in place.
  • May leave a slightly sticky residue on skin and feathers.
  • Some hydrophilic dressings may be sufficiently rigid to be sutured lightly into position, and may assist granulation of quite large areas.
  • Dressing should be changed every 2-3 days initially, once a week when a healthy granulation bed has been established.
  • Dressing must be changed if quantity of exudate results in leakage around the edge of the dressing (to prevent bacterial invasion).
  • Useful for e.g. slow-healing, granulating wounds over the keel and carpal joints, and for granulating bumblefoot lesions, also for extensive wounds with considerable exudate production and wounds requiring debridement.

ii) Semi-occlusive moisture-vapour permeable dressings.

  • Thin, transparent flexible polyurethane membrane.
  • Stick to clean, dry, detergent-free skin but not to the wound.
  • May be conformed to even difficult-to-bandage areas (e.g. head).
  • Permeable to oxygen and moisture vapour.
  • Impermeable to water and bacteria.
  • Allow fluid and exudate to accumulate under the dressing.
  • Maintain a moist, aerobic environment
  • Maintain a sterile surface (if wound aseptically cleaned beforehand) while margins remain sticking to surrounding skin.
  • Prevent desiccation and scab formation, reduce pain associated with the desiccation of nerve endings
  • Promote leukocyte debridement of the wound surface, and migration of epithelial cells from the wound edges (epithelialization), and thereby speed healing.
  • Allow visual monitoring of the wound and both qualitative and quantitative assessment of the production of exudate.
  • Dressing should be changed every 2-3 days initially, once a week when a healthy granulation bed has been established.
  • Dressing must be changed if quantity of exudate results in leakage around the edge of the dressing (to prevent bacterial invasion).
  • Use should be discontinued if excessive redness, swelling and/or odour indicates gross infection.

SECONDARY / MIDDLE LAYER:

Functions:

  • absorption of fluids and wound exudate
  • protection of the wound from additional trauma
  • immobilization of the wound while healing occurs

Conforming gauze bandages are usually used as the secondary bandage layer. N.B. figure - of -eight wing bandages should provide padding and immobilization due to their bulk, not by using a very tight bandage. Joint damage, vascular compromise and delayed healing may all be seen if bandages are improperly applied.

TERTIARY / OUTER LAYER:

Function: to hold the bandages in place. May also be used to protect the bandages from the attentions of the patient.

  • Self-adhesive bandages are commonly used. These are light-weight, breathable, conform well to avian anatomy e.g. limbs, and are generally well tolerated. White adhesive tape may be used in patients with a tendency to remove their bandages.

(P4.1990.w2, B11.3.w10, B13.16.w1, B14B116.30.w3)

Crane Consideration
  • Minor abrasions can be simply treated with a topical ointment.(D441)

Larger/more extensive wounds

  • First provide treatment for shock if required (see section above: Supportive Care and Nursing) giving fluids, corticosteroids and antibiotics; this is particularly important following intraspecific attack. (B115.8.w4)
  • Control haemorrhage as required, initially by direct compression of the wound site. (B115.8.w4)
    • Haemorrhage of small wounds can be controlled using ferric chloride, ferric sulphate or Monsell's solution. (B115.8.w4)
  • Pluck body feathers (NOT major wing or tail feathers) from around the wound with care not to restart the wound bleeding. (B115.8.w4)
  • Clean the wound(s) using warm saline solution, dilute (1%) povidone iodine solution or dilute chlorhexidine. (B115.8.w4)
  • Debride the edges back to fresh, clean tissue if these are not fresh or are grossly contaminated. (B115.8.w4)
  • Suture using 3-0 or 4-0 nylon or a similar diameter polyglycolic acid absorbable suture material. Use simple interrupted, simple continuous or horizontal mattress sutures as appropriate for the nature and extent of the laceration. (B115.8.w4)
    • If using nylon (non-absorbable) sutures, remove these after 10-14 days. (B115.8.w4)
  • Bandage as appropriate for protection. (B115.8.w4)
    • Consider that cranes may try to remove bandages using their bill. (B115.8.w4)
    • In general, use self-adhesive rather than adhesive bandaging when appling bandages to feathers; normal adhesive tape may be used (and prefered) for the featherless areas of the leg. (J29.3.w1)
  • If there are head injuries (e.g. following intraspecific aggression), once the crane is stabilised, radiograph the head to check for any damage to the skull. (B115.8.w4)
  • Lacerations should be treated as required depending on the degree of injury, from cleaning only with minor wounds to surgical repair of more extensive wounds. (B12.56.w14)
  • In the event of subcutaneous emphysema due to air sac damage, it may be necessary to draw air off and/or surgically insert Penrose drains and leave them in place for several days. (B12.56.w14, B115.8.w4)
Bear Consideration
  • If the wound is still bleeding, control bleeding by applying direct pressure to the wound or to the appropriate pressure point. (D249.w13)
  • Clip the hair around the wound. (B64.26.w5, D249.w13)
  • Flush the wound thoroughly with a weak solution of povidone iodine or chlorhexidine. (D249.w13)
    • It is particularly important to ensure that any pus (in an infected wound) or maggots (in a wound with myiasis) are flushed out of the wound. (D249.w13)
    • Do NOT cleanse deep wounds with hydrogen peroxide or alcohol, as this may cause tissue injury and increase the chance of infection developing. (D249.w13)
  • Debride any dead tissue. (B64.26.w5, D249.w13)
  • Apply topical antibiotics. (B64.26.w5)
  • Parental or oral antibiotics are recommended for five to seven days. (B16.9.w9)
  • If necessary, suture the wound using an absorbable suture material (B64.26.w5, D249.w13)
    • Absorbable sutures should be used for closure of the skin as well as deeper tissues, so that there is no need to remove the sutures. (B345.4.w4)
    • It is particularly important to use absorbable sutures in field situations when the animal will be released immediately. (B345.4.w4)
    • Use a tapered needle to suture internal muscle layers on a deep wound. (B345.4.w4)
    • Use a cutting needle to suture the skin. (B345.4.w4)
  • Note: 
    • Even quite large and infected wounds in adult polar bears (e.g. a suppurating wound more than 30 cm and another 18 cm open wound on one male, and a 50 cm long 6 cm deep wound on the upper thigh of another male, may heal well with minimal scarring. (J30.64.w1)

  • Note:

    • Note: Many wild animal casualties, particularly adult mammals, may not tolerate dressings and bandages.
    • Bears are strong and may interfere with external devices. (B64.26.w5)
    • Cubs may be prevented from interfering with casts etc. by fitting an Elizabethan collar. (B16.9.w9)

Lagomorph Consideration

  • It is important to remember that rabbits are prey species; care is needed to minimise stress levels during wound management, to avoid fatal catecholamine release. (J213.7.w2)
    • This is even more important with wild lagomorphs than with domestic rabbits which are used to people.
  • General systemic support is important for good wound healing. (J213.7.w2)
  • Evaluate the whole rabbit before addressing specific wound care. (J213.7.w2)
  • Avoid the use of Elizabethan collars on rabbits; these generally cause stress to the rabbit. (B600.15.w15)
  • An alternative cervical collar, named a "scratch guard" has been developed which is softer and apparently better tolerated. (B534.43.w43f)
    • A circular ring, about 14 inches diameter, is made from flexible tubing or roll gauze.
    • Four by four gauze sponges are wrapped around the tubing, padding the collar and increasing its external circumference; this gives a ring about 5 cm thick. (J501.33.w1)
    • Surgical adhesive tape, followed by self-adhesive tape (Vetwrap) is used to secure the gauze, provide a consistent collar diameter, and make it water-resistant.
    • The collar is placed over the rabbit's head.
    • Slack in the collar is compressed to give a snug fit, and maintained by holding the excess into a yolk with adhesive tape.
    • Rabbits wearing the collar are able to eat, drink and move normally, but without accessing an abdominal surgical site. (B534.43.w43f, J501.33.w1)
    • [Note: this was developed for use in laboratory rabbits undergoing surgery. There is a lack of published information on whether it has been used successfully in clinical situations for pet rabbits.]
Initial care and cleaning
  • The wound needs to be thoroughly cleansed. (J213.7.w2)
Clipping
  • Placing sterile lubricant (e.g. K-Y Jelly, Johnson & Johnson) in the wound before clipping the fur helps avoid further contamination. (B601.3.w3, J213.7.w2)
  • Gently shave the fur surrounding the wound, taking care to avoiding damage to the surrounding skin. (J213.7.w2)
    • Rabbits have thin, delicate skin which is easily damaged. (B600.15.w15, J213.7.w2)
    • The fine, dense fur easily clogs clipper blades. (B600.15.w15)
    • Use good-quality, robust clippers, and clip slowly to prevent fur catching. (B600.15.w15)
    • Avoid clipping the soles of the feet, as loss of the fur removes the protection from this area and the rabbit is then likely to develop pododermatitis. (J213.7.w2) See: Ulcerative Pododermatitis in Lagomorphs
  • Depilatory creams can be used but are messy and difficult to clean off properly. (B600.15.w15)
Wound lavage
  • Gentle lavage should be used to clean the wound, removing any debris. (J213.7.w2)
    • Avoid aggressive lavage; this can cause further damage to tissues and spread bacteria deeper into the tissues. (B601.3.w3, J213.7.w2)
  • Lavage solutions (sterile and ideally warmed to body temperature) include:
    • Isotonic (0/9%) saline (B601.3.w3, J213.7.w2)
    • Lactacted Ringer's solution (Hartmann's solution). (J213.7.w2)
    • Ringer's solution. (J213.7.w2)
  • Equipment
    • A container of lavage solution connected to an intravenous administration set, this then connected via a three-way stopcock to a 30 mL syringe with 18 gauge needle. (J213.7.w2)
Cleansing of wound and surrounding skin
  • Antiseptic detergents
  • Hydrogen peroxide
    • This "can be used as a one-time irrigation solution if an anaerobic infection is suspected, particularly Clostridium species, but hydrogen peroxide can be cytotoxic". (J213.7.w2)

For different types of wound:

Contaminated wounds

  • Saline or lactated Ringer's solution
    • Advantages of being sterile, isotonic and isosmotic but has no antimicrobial activity. (J213.7.w2)
  • Chlorhexidine
    • Advantages of being wide spectrum and having residual activity but it does precipitate and there is some gram negative resistance. (J213.7.w2)
    • Use as a diluted solution to cleanse the wound.
  • Povidone iodine (Iodophors)
    • Advantages of being wide spectrum and not causing irritation. However, organic material inactivates it. (J213.7.w2)
    • Use as a diluted solution to cleanse the wound.

Grossly contaminated wounds

  • Hydrogen peroxide
    • Advantages of removing dirt and being sporicidal but it has minimal antibacterial properties and causes cell injury. (J213.7.w2)
Wound Closure
  • Normal wound healing: in rabbits this generally takes 14 to 16 days to complete. (J213.7.w2)
  • Primary closure:
    • Ideally, wounds should be debrided and closed if circumstances allow. (J213.4.w4)
    • Performed if the wound is recent and there is minimal contamination. Debride any non viable tissue first. (J213.7.w2)
  • Other closures: Wounds can also be allowed to heal by delayed primary closure, secondary closure, or sometimes by second intention where an open wound heals by contraction and epithelialisation. (J213.7.w2)
  • Suture material:
    • In general, the non-absorbable monofilament sutures will create less intense tissue response in the rabbit than either the nonabsorbable multifilament sutures or the absorbable sutures. (J213.7.w2)
    • Polyglactin 910 is an absorbable suture which has been reported to have a prolonged extensive reaction in rabbits. (J213.7.w2)
    • Polypropylene: Although this suture material is nonabsorbable and monofilamented, it was reported to produce a relatively thick capsule response, possibly because of its rigidity and stiffness. (J213.7.w2)
    • In closure of infected / contaminated wounds: (e.g. when skin is sutured over AIPPMA beads) use fine monofilament materials with small knots and avoid burying suture material.
    • In noncontaminated skin closure, use an absorbable monofilament suture, e.g. polydioxanone or poliglecaprone 25, in a continuous subcuticular pattern and with a buried knot. Rabbits generally do not bother their wounds if the sutures are comfortable. However, some may remove skin sutures. Surgical staples may also be used. 
  • Elizabethan collars are often stressful to rabbits, and prevent normal coprophagy; they should be avoided where possible. (J213.7.w2, J83.29.w2, V.w128)
  • Preventing bandage or incision chewing: Noxious agents are generally not effective. (J213.7.w2)
Topical medication

The appropriate topical medication depends on the wound type.

  • Intrasite Gel (Smith & Nephew) can be used in a wide variety of wounds including lacerations, deep punctures, pressure sores etc. (B601.3.w3)

Small contaminated wounds

  • Bacitracin-Neomycin-polymyxin 
    • Advantages of being wide spectrum, having minimal toxicity and stimulating reepithelialisation. However, it is not effective on infected wounds; this product is often oil based and it has the potential to cause a local allergic reaction. Ingestion may be problematic in rabbits. (J213.7.w2)

Burns and wounds with necrotic tissue

  • Silver sulfadiazine
    • Advantages of being wide spectrum with antifungal properties and it is also painless and stimulates reepithelialisation. However, it may delay eschar separation (it impedes contraction) and it possibly causes bone marrow suppression if treating large wounds. (J213.7.w2)

Contaminated and infected wounds; chronic wounds

  • Hydrophilic agents 
    • D-Glucose polysaccharide (Intracell)
    • Advantages of promoting chemotaxis and enhancing epithelialisation. Glucose is provided for cell metabolism and the hydrophilic properties reportedly pulls fluid up  through the wound tissue and so bathes the wound from the inside. However, it has no direct antimicrobial activity. (J213.7.w2)

Contaminated and infected wounds; myiasis

  • Enzymes
    • Trypsin-balsam of Peru (Granulex)
    • Advantages of enzymatic debridement, angiogenesis and it also improves reepithelialisation. Useful in the initial management of wounds with necrosis or myiasis. "The preparation "bubbles" fly larvae out of wounds". However, it does cause local inflammation and a pyogenic reaction and so is not advised for long-term wound management. It also may produce a stinging sensation and it has no antimicrobial effects. (J213.7.w2)

Burns, ulcers and abrasions

  • Hydrogel wound dressing with acemannan
    • Advantages of stimulating angiogenesis and epithelialisation and is non toxic. It has been reported to enhance healing of burn wounds in guinea pigs. A freeze dried form of this product will apparently reduce tissue oedema by absorbing fluid from the wound as it converts to a gel. Daily application of this freeze-dried form is reported to stimulate the formation of granulation tissue over exposed bone and therefore enhancing wound contraction. However, it has no direct antimicrobial effects. (J213.7.w2)

Wounds in repair stage

  • Tripeptide-copper complex gel
    • Iamin-Vet Skin Care Gel (J213.7.w2)
    • Advantages of stimulating collagen synthesis and angiogenesis. It is also chemoattractant. Injections of this product into wounds have reportedly increased the wound healing when compared with the controls that just used saline. However, it has no direct antimicrobial effects. (J213.7.w2)

Urine or faecal scald wounds

  • Hexamethyldisiloxane acrylate copolymer
    • No Sting Barrier Film (J213.7.w2)
    • Advantages of producing a uniform, fast drying, transparent, noncytotoxic film. Useful as a skin protectant and allows the skin inflammation beneath the film to quickly resolve. Apply to dry, clean skin every third day. However, it has no direct antimicrobial effect. (J213.7.w2)

Contraindications

  • Topical steroid preparations may cause:
    • Adrenocortical suppression (J213.7.w2)
    • A significant delay in fibroblastic proliferation, angiogenesis, and synthesis of collagen and proteoglycans. Therefore impairing epithelialisation, wound strength, and the closure of open wounds. (J213.7.w2)
  • Nitrofurazone
    • Wide spectrum and hydrophilic, but may slow epithelialisation and is a known carcinogen. Therefore, it is not recommended in treating rabbit wounds. (J213.7.w2)
Dressings

Bandages usually have three layers to them: a primary layer that is in contact with the wound, a secondary layer that is absorptive and has stabilising properties, and a tertiary layer that holds the first two layers in place.

Note: body bandages may be poorly tolerated by rabbits, often slip, may interfere with normal breathing, and may cause overheating. (B601.3.w3)

Primary layer
  • There are various dressings that can be used as the first layer to speed wound healing:
  • Adherent dressings e.g. gauze pads, or wet-to-dry bandaging techniques, are useful in the initial wound management when debridement is necessary. (J213.7.w2)
  • Nonadherent dressings are needed during the reparative stage of wound healing. These types of dressings include occlusive and semiocclusive materials. (J213.7.w2)

    • Semiocclusive materials:
      • cotton nonadherent bandages
      • petrolatum-impregnated bandages
      • polyurethane foam sponge materials
      • tegaderm
    • Occlusive materials 
      • hydrocolloid dressing
      • hydrogel
      • calcium aginate
  • Other dressing materials
    Exogenous collagen that is used in wounds acts as a template for the migrating fibroblasts and allows for new deposition of collagen. 

(J213.7.w2)

Contaminated or infected wounds

  • Cotton nonadherent bandages
    • Advantages of being absorbent and helping to keep the wound dry. It allows excess fluid to seep into the secondary bandage layer. Also useful in wound debridement. However, due to adherence, they can disrupt the wound surface if left on for a prolonged period and removal may be uncomfortable. (J213.7.w2)
  • Polyurethane foam sponge material
    • Absorbs fluid and maintains a moist wound. Ideal for exudative wounds. Wetting agents or liquid medication can be applied to the foam for treatment of wounds. The foam provides a layer of padding in between the wound and the secondary bandage layer so is useful for wounds that need some additional padding, e.g. Ulcerative Pododermatitis lesions. The disadvantage to this product is that there may be edge adherence of this type of bandage. (J213.7.w2)

Wounds that are in repair stage with a healthy bed of granulation bed

  • Petrolatum-impregnated bandages
    • Comfortable and keeps wounds moist but it does slow epithelialisation. Less adhering than the cotton nonadherent bandages. (J213.7.w2)
  • Polyurethane foam sponge material
    • Absorbs fluid and maintains a moist wound. However, there may be edge adherence of this type of bandage. (J213.7.w2)
  • Tegaderm
    • A moisture or vapour permeable semiocclusive dressing. (J213.7.w2)
    • Maintains wound moisture and leads to a more rapid healing of the wound. It is adhesive and so may be used on its own over wounds without additional bandaging materials depending on the site. However, it does have a difficult application technique. (J213.7.w2)
  • Hydrocolloid (Duoderm; Dermaheal)
    • This dressing will absorb fluid to create a gel that enhances epithelialisation and needs less frequent changes than some types of bandage. Its disadvantages are that its not transparent; there may be reduced wound contraction; and difficult removal from the skin surrounding the wound. (J213.7.w2)
  • Hydrogel dressing (BioDres)
    • "A hydrophilic polyethylene oxide polymer composite". (J213.7.w2)
    • Also useful in noninfected eschar. (J213.7.w2)
    • This product absorbs fluid and enhances epithelialisation; it's transparent; and there is not a problem with adherence to the skin surrounding the wound. However, it may cause exuberant granulation tissue. (J213.7.w2)
  • Porcine small intestinal submucosa (Vet Biosist)
    • This product will act as a matrix for wound healing and it may also assist granulation over the bone. However, it can prolong contraction of the wound. (J213.7.w2)

Moderately to heavily exudative wounds

  • Calcium alginate (Curasorb)
    • This felt-like pad absorbs considerable wound fluid and it also enhances epithelialisation. However, a calcium alginate eschar may be produced if the wound is not producing enough fluid to convert the pad to a gel. (J213.7.w2)

Wounds in the late inflammatory or the early repair stage

  • Exogenous collagen matrix (Bovine collagen)
    • This product will act as a matrix for fibroblast migration by causing an inflammatory reaction that will enhance collagen deposition. (J213.7.w2)
Skin flaps and grafts

Skin flaps

  • A skin flap is "a partially detached segment of skin and subcutaneous tissue whose base maintains circulation to the skin during elevation and movement to a recipient bed". (J213.7.w2)
  • It is necessary to have knowledge of the vascular anatomy in order to preserve the circulation when creating a flap. (J213.7.w2)
  • In companion mammals, it is the subdermal plexus that is responsible for blood supply with simple flaps. (J213.7.w2)
  • The single pedicle advancement flap is used to repair cutaneous defects where the skin is advanced to cover a defect without movement laterally; this method relies on stretching of the elastic skin adjacent to the wound. (J213.7.w2)

Skin grafts

  • A skin graft is "a portion of dermis and epidermis that is completely detached from its original location and moved to a recipient site where its survival depends on the absorption of tissue fluid and the development of a new blood supply". (J213.7.w2)
  • Use of a skin graft is indicated when a cutaneous defect cannot be closed by moving local skin surrounding the wound, particularly on large wounds on the trunk or wounds on the distal limbs.
  • There are three physical types of skin grafts:
    • Full thickness: epidermis and dermis. These are useful for distal limb wounds, on flexor surfaces to prevent contracture, and on large skin defects on the trunk. (J213.7.w2)
    • Split thickness: epidermis with various dermal thicknesses. These heal quicker than the full thickness grafts because of "the increased number of capillaries on the derma surface of the graft available for inosculation with vessels of the recipient bed". (J213.7.w2)
    • Perichondrial cutaneous graft (PCCG): harvested from the ear with the following layers: perichondrial layer, scant subcutaneous tissue, dermis and epidermis. One study reported superior coverage (better contraction properties, thickness, and hair retention) with this type of graft in comparison with full thickness skin grafts. (J213.7.w2)
  • Meshing the skin will improve the contact between the wound surface and the graft. Also, the mesh holes allow serum, exudate, and blood to drain from under the graft. This prevents infection and decreases the distance for plasmatic absorption. (J213.7.w2)
    • However, one study of rabbit non-meshed and meshed split thickness skin grafts showed a significantly greater rate of growth and wound contraction in the non-meshed grafts. (J213.7.w2)
  • Contraindicated in areas that have:
    • infection
    • poor vascularity 
    • fat 
    • excessive movement 

    (J213.7.w2)

Ferret Consideration
Types of wounds
A variety of wounds can be found in ferrets depending on the cause of the trauma. (J213.7.w5)
Wound healing
  • There are four stages to wound healing: (J213.7.w5)
    • Inflammation (caused by histamine, thromboxane and growth factors) and haemorrhage. (J213.7.w5)
      • Inflammation may last up to five days. (J213.7.w5)
    • Debridement - the body clears away the rubbish with exudate (containing white blood cells and platelets). (J213.7.w5)
    • Regeneration of tissue (three to five days after injury). (J213.7.w5)
      • Fibroblasts deposit collogen. (J213.7.w5)
    • Maturation with myofiboblasts (seventeen to twenty days post surgery). (J213.7.w5)
      • This stage can last for several years. (J213.7.w5)
Wound management
  • The ferret should be stabilized prior to wound management. (J213.7.w5)
  • Wound management in ferrets is similar to that in other mammals. (J213.7.w5)
  • Bleeding should be stopped first. (B652.6.w6)
    • Incised wounds may bleed heavily. Apply direct pressure to stop the bleeding. Suture and apply a bandage. (B651.9.w9)
  • Pain relief should be given if necessary: (J213.7.w5)
  • Systemic antibiotics should be used, as with cats and dogs. Aminoglycosides should be avoided in ferrets. (J213.7.w5)
    • A culture and sensitivity may be run to find the appropriate antibiotic. (J213.7.w5)
  • A balanced diet should be given during the healing process. (J213.7.w5)
  • An older ferret or a ferret with a disease such as hyperadrenocorticism (Adrenocortical Neoplasia in Ferrets) may take longer to heal. (J213.7.w5)
  • A good blood supply to the wound increases the rate at which the wound will heal. (J213.7.w5)
  • Corticosteroids may slow wound healing down; infection may also be more likely if corticosteroids are used. (J213.7.w5)
  • If the wound is severely contaminated and has been left open for more than six to eight hours, the wound should be left open. (J213.7.w5)

Lavage

  • Warmer temperature lavage encourages wound healing, while cold temperature lavage fluid causes restriction of blood vessels. (J213.7.w5)
  • To ensure the wound is clear of debris, is important for wound healing. Use lactated ringer solution and high pressure, this will help ensure removal of bacteria. (J213.7.w5)
    • Chlorhexidene diacetate, also will kill bacteria. (J213.7.w5)
    • Povidine-iodine, will kill bacteria, yeast, fungi and viruses. (J213.7.w5)
    • Tris-EDTA assists the lavage solutions to kill bacteria. (J213.7.w5)

Topical antibiotic

  • Care should be taken when using antiseptic drugs, as this may cause delay in wound healing. (J213.7.w5)
  • To continue debridment, antibiotics or wet-dry dressing should be used on open wounds. (J213.7.w5)
    • Topical antibiotics:
      • Neobaciymx (effectiveness against pseudomonas sp. is poor). (J213.7.w5)
      • Silver sulfadiazene1% cream (good for treating burns). (J213.7.w5)
      • Gentamicin sulfate 0.3% ointment (good for gram negative bacteria, before and after grafting tissue). (J213.7.w5)
      • Nitrofurazone (this helps draw fluid away from the wound, as well as having antimicrobial effects). (J213.7.w5)
      • Aloe vera can be used as an antibiotic and to help vascular patency. (J213.7.w5)

Surgery

  • The wound should be closed as soon as possible after the trauma, once the wound is clear of bacteria. (J213.7.w5)
  • Debriding the skin is important before suturing the edges of the wound together. (J213.7.w5)
  • If bullet wounds are present, the shot will need to be surgically removed. (B651.9.w9)
  • A drain can be used to help remove fluid from the wound and reduce dead space (Penrose drains are commonly used for this purpose). (J213.7.w5)

Bandage

  • If there is tension on the wound, then a bandage should be used initially. (J213.7.w5)
  • Bandages may be used on wounds, dry or wet, to keep the wound clean, reduce haemorrhage and oedema. (J213.7.w5)
    • Wet-to-dry bandaging will encourage debridement. (J213.7.w5)
  • Bandages keep wounds warm, which increases circulation and improves the healing process. (J213.7.w5)
  • Always place the bandage on distal limbs, to reduce oedema. (J213.7.w5)
  • A buster collar may be required if a bandage is used, to prevent the ferret from interfering with it. (J213.7.w5)
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
  • Wounds due to intraspecific bites and play objects are common in bonobos. (J23.20.w2)

General primate/great ape information

  • Wounds in primates should be examined and cleaned thoroughly, using lavage, e.g. using saline, running water or hydrogen peroxide. (B10.44.w44i, D409.6.w6)
  • Debridement should be carried out. (D409.6.w6) Necrotic tissue should be debrided. (B10.44.w44i)
  • If possible, suture the wound to allow healing by primary intention.
    • If sutures are not under excessive tension they are more likely to be left alone by primates. (B10.44.w44i)
    • Assess the wound, considering depth, effect on a particular area and infection risks before carrying out primary closure with care not to trap debris or create an anaerobic environment. (D409.6.w6)
  • If suturing is not possible, leave healthy tissue to granulate. (B10.44.w44i)
  • A long-acting antibiotic should be injected (e.g.  Penicillin G benzathine) to reduce the risk of infection. (B10.44.w44i)
  • Positive reinforcement training (Mammal Handling & Movement - Husbandry Training) may allow post-operative lavage or topical wound treatment. (D409.6.w6)
Associated techniques linked from Wildpro

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Fluid Therapy

Fluid therapy is an important part of patient care, and is a vital part of initial patient stabilization, whatever the presenting problem of the patient in question. Dehydration and electrolyte losses may be severe and even life-threatening in an ill or injured animal, particularly in small species and in neonates.

Aims of Fluid Therapy
  • Fluid therapy aims to correct life-threatening hypovolaemia and hypoperfusion, maintain intravascular volume and osmotic pressure, treat dehydration, and correct electrolyte and acid-base imbalances. (J15.6.w1, J15.30.w4)
    • It is also necessary to provide normal daily fluid requirements and meet continuing losses. (J15.6.w1)
  • The goal of fluid therapy in shock is "to optimise vascular volume, restore circulatory function and tissue perfusion, and ultimately deliver oxygen to tissues". At the same time, the smallest amount of fluids needed should be given and volume overload needs to be avoided. (J29.13.w1)
  • It is important first to ensure adequate tissue perfusion (transport of fluid and oxygen through blood vessels to the capillaries, which requires a functioning cardiovascular system and adequate intravascular volume, then to ensure adequate interstitial hydration - the presence of fluid in the interstitial space, supporting cells and providing a transport medium for molecules. (J29.13.w1)
    • Perfusion can be assessed by resting heart rate, blood pressure, mucous membrane colour and capillary refill time. (J29.13.w1)
    • Hydration can be assessed by mucous membrane moisture, skin turgor and ocular globe position. (J29.13.w1)
    • If dehydration exceeds 10%, an intravascular fluid deficit may also occur. (J29.13.w1, V.w124)
  • For the treatment of dehydration, the aim is to provide maintenance levels of fluids plus replace the estimated deficit over a period of 12 to 24 hours. (J15.30.w5)
  • Maintenance levels are 1.0 - 2.0 mL/kg/hour using an isotonic crystalloid such as lactated Ringer's solution. (J15.30.w5, V.w124)
  • During surgery, in cats and dogs, 10 mL/kg/hour of an isotonic crystalloid is appropriate. (J15.30.w5, V.w124)
Signs of hypovolaemia, dehydration and volume overload
  • Hypovolaemia - a reduction in intravascular volume - is indicated by: tachycardia, abnormal pulse quality (increased amplitude with mild, compensatory hypovolaemia, moderate decrease with moderate hypovolaemia, and severe decrease with severe hypovolaemia) and decreased pulse duration), altered mucous membrane colour (normal or pinker with mild hypovolaemia, pale pink with moderate hypovolaemia and white or muddy with severe hypovolaemia), and increased capillary refill time with moderate or severe hypovolaemia. Depending on the severity of the hypovolaemia, there may also be tachypnoea, cool extremities and altered mental status. (J15.30.w4)
  • Dehydration - strictly, a loss of water, but generally used to indicate loss of iso- or hypotonic fluids from the body, can be estimated from skin turgor and mucous membranes. A deficiency of less than 5% is not clinically detectable; at 5 - 6% dehydration a slight loss of skin elasticity may be detectable. At 6 - 10% dehydration, skin elasticity is reduced and mucous membranes may be dry, as well as the eyes possibly being in a sunken position. At 10-12% dehydration loss of skin elasticity is very obvious (tented skin stays tented) the eyes are sunken in their orbits and mucous membranes are dry. At 10-12% dehydration, there are additional signs of shock such as tachycardia, cool extremities, weak and rapid pulse and prolonged capillary refill time. The level of dehydration may also be assessed based on PCV, total solids and urine specific gravity. (J15.30.w4)
  • Signs of volume overload may be seen with inappropriate fluid therapy and include increased distension of of jugular veins, increased central venous pressure, increased respiratory rate/effort, and crackles on thoracic ausculation. (J29.13.w1)
Types of Fluids
Crystalloids
  • Crystalloid fluids contain water, electrolytes and non-electrolyte solutes. (J15.30.w5)
  • Crystalloids may be used in the treatment of hypovolaemic or septic shock, dehydration, hyperkalaemia, hypercalcaemia and for replacement of ongoing fluid losses. (J15.30.w5)
  • Crystalloids are hydrators of the interstitial space rather than expanders of the intravascular volume. In the short term, crystalloids do expand the intravascular space. However, due to redistribution, by one hour after administration only 20% of the initial volume remains in the intravascular space. (J29.13.w1)
A variety of crystalloid fluids are available, including:

Isotonic fluids

  • 0.9% saline (physiological saline) may be used in the treatment of hypovolaemic or septic shock, dehydration, hyperkalaemia, hypercalcaemia and for replacement of ongoing fluid losses. Care must be taken that it does not exacerbate acidosis, or result in hypernatraemia or hyponatraemia. Plasma potassium and plasma sodium should be monitored, and potassium supplemented if necessary. (J15.30.w5)
  • Lactated Ringer's solution (Hartmann's solution, compound sodium lactate) may be used in the treatment of hypovolaemic or septic shock or dehydration, and for replacement of ongoing losses. It should not be mixed with blood products or with sodium bicarbonate. Plasma potassium should be monitored and supplemented if necessary. (J15.30.w5)
  • Plasma-Lyte 148 (Baxter Healthcare) and Normosol-R are commonly used in the USA but less common in the UK. They are broadly similar to Hartmann's solution, but with acetate and gluconate rather than lactate as buffer and bicarbonate precursor, slightly higher sodium (140 mmol/L rather than 130 mmol/L, slightly higher osmolarity (295 mOsm/L rather than 275 mOsm/L), and containing magnesium at 3 mmol/L (none in Hartmann's). (J15.30.w5)
  • Plasma-Lyte M and Normosol-M contain glucose, sodium, potassium, chloride, calcium, magnesium and lactate; they are suitable for use as maintenenace fluids. These fluids can be infused through the same line with blood products. (J15.30.w5, V.w124)
  • Dextrose 4% with saline (0.18%) has limited use. With the addition of potassium it could be used in patients unable to take water orally, but these would need e.g. a feeding tube for parenteral nutrition, and fluids could be given also by that route. (J15.30.w5)
  • 0.45% saline may be used in the treatment of dehydration and for replacement of ongoing losses, but glucose and potassium should be added. Care must be taken that it does not result in hypernatraemia or hyponatraemia. Plasma potassium and plasma sodium should be monitored, and potassium supplemented if necessary. (J15.30.w5) It can be used combined with 0.9% saline in the treatment of hyponatraemic individuals, to provide solutions with varying levels of sodium, allowing slow adjustment of the patient's sodium concentration. (J15.30.w5)
Hypotonic
  • Dextrose 5% (5% dextrose in water, DW5) is isotonic when given, but the dextrose is rapidly metablised, leaving water which is distributed through the three fluid compartments; therefore this is biologically hypotonic. It can be useful for pure water loss, such as may occur in pure dehydration e.g. associated with hyperthermia and panting. (J15.30.w5) It is not useful for restoration of circulating fluid volume since most of the fluid redistributes into cells, and it is does not provide an adequate source of energy. (J15.30.w5)
    • It is primarily used to carry drugs as sodium nitroprusside or dopamine for constant rate infusion. (V.w124)
    • 5% dextrose is rapidly absorbed if given orally. (B119.w2). 
    • Note: Dextrose 10% or 50% solution may be used to treat hypoglycaemia. Initial dose of 1mL/kg 50% dextrose may be used intravenously. N.B. Disadvantage of dextrose is the promotion of cellular acidosis (B119.w2).

Hypertonic

  • Hypertonic saline: Usually a 7.5% solution (2600 mOsm/L). A hyperosmolar (hypertonic) crystalloid used in the treatment of hypovolaemia along with hetastarch. Hypertonic saline acts by drawing fluid out of the interstitial spaces and intracellular spaces into the intravascular space. (J15.30.w5, J29.13.w1, P113.2008.w1, V.w124)
    • 4 mL/kg is given at up to 1 mL/kg/minute (J15.30.w5); 5 mL/kg is given over five to ten minutes (J29.13.w1) to provide a rapid increase in intravascular volume with only 0.25% of the volume which would be required to produce an equivalent intravascular fluid volume increase using colloids. However the effect is transient (less than 30 minutes (J29.13.w1); 30 - 120 minutes (J15.30.w5).
    • Hypertonic saline should be followed by replacement crystalloids to replace the fluid deficit (J15.30.w5); must be used with a colloid for continuing effect (hetastarch is given at 3 - 5 mL/kg). (J29.13.w1, V.w124)
    • The use of hypertonic saline should be avoided in individuals which are dehydrated (since the exravascular fluid volume is already depleted). (J29.13.w1)
    • Potential side effects include hypernatraemia, hyperchloraemia, hypokalaemia and dehydration. (J15.30.w5, J29.13.w1)
      • Relative contraindications include hypernatraemia, hypokalaemia and dehydration. (J15.30.w5)
    • Too rapid an infusion may cause hypotension, bradycardia, ventricular dysrhythmias, bronchoconstriction and rapid shallow respiration. (J15.30.w5)

Notes:

  • Fluids suitable for fluid replacement contain electrolytes at concentrations similar to extracellular fluid. Suitable fluids for fluid replacement include 0.9% saline, lactated Ringer's solution, Normosol-R, Plasmalyte-A. (J29.13.w1)
  • Fluids suitable for maintenance contain 40 - 60 mEq/L sodium and 15 - 30 mEq/L potassium. (J29.13.w1)
  • It has been suggested that fluids containing lactate should be avoided in patients with severe liver compromise, since this must be broken down in the liver. (J29.13.w1) However, it has also been noted that in practice this does not appear to be a problem in patients with hepatic impairment. (J15.30.w5)
Colloids
  • These fluids contain substances of large molecular weight, generally unable to pass through capillary membranes; they can be considered as intravascular volume expanders. (J29.13.w1)
  • Colloids are important in the treatment of shock, since patients in shock generally need sustained expansion of the intravascular volume. (J29.13.w1)
  • As well as synthetic colloids (e..g. hetastarch, Oxyglobin), biological colloids (whole blood, plasma, albumin) can be used.
  • Hetastarch (hydroxyethyl starch, HES solutions) are made from maize or sorghum. (J29.13.w1) They are modified polymers of amylopectin; intravascular hydrolysis is reduced because they are hydroxyethylated (J15.30.w6). These solutions contain large molecular weight particles with a negative charge, attracting sodium and water to expand the intravascular volume by about 1.4 times the volume infused. The average molecular weight is 450,000 Daltons; half life is 25 hours. If more than 40 mL/kg per day is given, increased incisional bleeding and coagulopathies have been reported. (J29.13.w1, V.w124)
  • Dextrans are glucose polymer solutions. They provide a marked, temporary expansion of plasma volume, by 1.5 to 2.0 times the volume given, but half the infused volume is lost from the vascular space over the following three hours. Solutions containing more large molecules (e.g. Dextrans 70) are cleared from the intravascular space more slowly (loss of 35% of polymers over 12 hours, by enzymatic degradation) than products with less large polymers (e.g. Dextrans 40). (J15.30.w6)
    • Renal failure has been reported with use of Dextran 40, particularly associated with hypovolaemia and pre-existing renal dysfunction. (J15.30.w6)
    • Anaphylactoid reactions to dextrans have been reported, but rarely severe reactions. (J15.30.w6)
  • Gelatins are polydisperse colloids from alkaline-modified bovine collagen. Haemaccel (Intervet) is a urea-linked gelatin, while Gelofusine (Braun) is a succinylatted gelatin. Gelatins have about 80% of their molecules smaller than 20 kDa, which are rapidly removed via the kidneys. These solutions have an intravascular persistence of about two to three hours. (J15.30.w6)
    • An anaphylactoid reaction may occur to gelatin-based products. (J15.30.w6)
  • Note: dilutional coagulopathy may occur when synthetic colloids are used (J15.30.w6, J29.13.w1); this is less of a problem with gelatin-based products. (J15.30.w6)
Whole blood and blood products
  • Blood is ideal if whole blood has been lost, to ensure adequate oxygen is delivered to cells. (J29.13.w1)
  • If clotting factors have been lost, whole blood or fresh frozen plasma are needed to replace clotting factors. Fresh frozen plasma is also useful if albumin has been lost; frozen albumin can also be used.
  • Note: a major limitation is the availability of adequate blood products to match the patient species. (J29.13.w1)
  • Can be used alongside crystalloids to avoid interstitial volume depletion; give 40-60% of the crystalloid dose which would be given if no colloids were used. (J29.13.w1)
  • Whole blood: give at 20 - 25 mL/kg. For treatment of acute haemorrhage of > 20% of blood volume. The aim is to stabilise clinical signs of chock, maintain a haematocrit of > 25% and keep clotting times within the normal range. (J29.13.w1)
    • Ideally, give over 2 - 4 hours, monitoring for transfusion reaction and avoiding volume overload. In practice give in boluses in life-threatening situations. (J29.13.w1)
  • Human albumin solution (HAS) is a monodisperse colloid containing albumin (molecular weight 69 KDa). It is of limited use in veterinary medicine, being expensive and having no clear advantages over other products. (J15.30.w6) 
    • It is being used in veterinary critical care patients that are hypoalbuminemic, given very slowly as a constant rate infusion. Anaphylactic and mild reactions have been reported. (V.w124)
  • Fresh frozen plasma is produced by centrifugation of whole blood and freezing within six hours of collection. It can be stored frozen for up to a year at - 30C. Once, thawed, it can be used within24 hours (V.w124) if kept chilled, or refrozen (then labelled as frozen plasma, rather than fresh frozen plasma). It contains albumin, alpha-2-macroglobulin, acute phase proteins, antithrombin and immunoglobulins, as well as fibrinogen, Von Willebrand's Factor and coltting factors II, V, VII, VII, IX, X and XI. (J15.30.w6)
    • This is particularly useful for severe cogulopathies (e.g. rodenticide toxicity, sepsis), also in systemic inflammatory response syndrome and in surgical patients (replacement of coagulation factors, support for wound healing, provision of drug binding capacity alongside pH buffereing and volume replacement). (J15.30.w6)
Haemoglobin-Based Oxygen Carriers (HBOC)
  • These provide oxygen carrying capacity to the tissues as well as acting as colloids.
  • Oxyglobin is a haemoglobin-based oxygen carrying solution. It contains purified polymerized haemoglobin, of bovine origin, in modified lactated Ringer's solution. it is iso-osmotic. (J29.13.w1, P3.2000b.w3)
  • It has a relatively low oxygen affinity and therefore readily offloads oxygen to tissues. (P3.2000b.w3, J29.13.w1)
  • The oxygen affinity is dependent on the concentration of chloride ions, not on 2,3,-diphosphoglycerae concentration. (P3.2000b.w3, J29.13.w1)
  • In dogs, a one-time dose is 30 mL/kg bodyweight, with a maximum administration rate of 10 mL/kg/hour. (P3.2000b.w3)
    • When used with crystalloids, a 5 mL/kg bolus is given, titrated to correct hypovolemia. (V.w124)
  • In cats, much lower doses must be given.
    • Use 2 mL/kg increments with crystalloids titrated to correct hypovolemia, but given slowly over 5-10 minutes. (V.w124)
  • Note: Oxyglobin acts as a colloid; to avoid fluid overload it is important to control the rate of administration if it is used in a normovolaemic anaemic animal. (P3.2000b.w3)
  • Side effects include: 
    • Skin, mucous membranes and sclera are discoloured yellow-orange, and urine becomes red-brown. This colouration, which is dose dependent, usually resolves over 3 - 5 days. (P3.2000b.w3)
    • All colourimetric tests, such as serum chemistry measurements and urine dipstick results, are inaccurate. (J29.13.w1, P3.2000b.w3)
    • Packed cell volume and haemogolbin concentration do not correlate; haemoglobin concentration must be measured directly. (J29.13.w1)
    • Mild gastro-intestinal tract effects have been recorded rarely. (J29.13.w1)
  • Note: this has not been approved for use in most species - approved for use in dogs only (J29.13.w1, V.w124)
  • Cats: give 2 mL/kg over a period of 10 - 15 minutes until normal heart rate and a systolic blood pressure > 90 mm Hg are reached, then give 0.2 - 0.4 mL/kg/hr as a continuous rate infusion. (J29.13.w1, V.w124)
  • This can be given through a 22- or 24-gauge catheter using a standard infusion pump; no pre-treatment or filtration is needed as there is no cellular debris in the fluid. (J29.13.w1, P3.2000b.w3, V.w124)
  • No cross-matching is needed. (P3.2000b.w3,)
  • It can be stored unopened for three years, but once opened, a 125 mL package needs to be used or discarded within 24 hours due to formation of methaemoglobin. (J29.13.w1, P3.2000b.w3)
  • Suitable for the treatment of acute blood loss/haemorrhagic shock e.g. following acute trauma or intraoperative blood loss. (J29.13.w1)
  • Because this is a colloid, use with caution in normovolaemic individuals (e.g. in the treatment of immune-mediated haemolytic anaemia) and never use in hypervolaemic individuals (e.g. congestive heart failure, or with oliguric or anuric renal failure); give slowly and monitor for signs of fluid overload. (J29.13.w1, V.w124)
  • Half-life is 30-40 hours; the primary clinical effect lasts about 24 hours and 90% is eliminated within five to seven days. (J29.13.w1)
Initial Calculations of Requirements
  • Requirements will vary depending on the physiological status of the patient: correction of perfusion deficits if these are present, then correction of dehydration, and finally maintenance. (J29.13.w1)
  • Note: initial choice of fluid type and volume should be based on the known or assumed types of fluid loss and the clinical status of the patient. Any protocol or "rule of thumb" should be considered as a guideline only and fluid therapy should be modified depending on the needs of the individual patient at a given time. (J29.13.w1)
  • Fluid therapy should be reassessed periodically and changed based on a variety of parameters including: blood pressure, hydration status, PCV, total proteins, urine output, acid/base balance and mental status of the patient. (J29.13.w1)
  • If calculated therapy is given and does not result in the required end-point parameters (normal hear rate and blood pressure, mucous membrane colour and capillary refill time), further evaluation is required. (J29.13.w1)
  • For wildlife casualties, it should be assumed that animals presenting with an injury will be shocked and dehydrated. (J213.9.w4)
  • Fluids should be given to meet maintenance fluid requirements (average 60 mL/kg/day for most mammals) plus the individual's fluid deficit (percentage dehydration x body weight in kilograms x 1000 x 0.8) plus ongoing losses (2 x estimated volume lost). This amount should be given over 24 - 48 hours, divided into e.g. four treatments if giving subcutaneously. (J213.9.w4)
    • Average maintenance 48 mL/kg/day; estimate losses due to e.g. vomiting and diarrhoea. (V.w124)
Routes:

The appropriate route for fluid therapy depends on factors such as the severity and duration of the disease process causing the need for fluid therapy, the goal of the fluid therapy, the type of fluid to be given, the costs and availability of fluids and equipment and the technical skills of personnel. (B150.w3)

  • Oral: 
    • This route is useful in individuals that are not severely dehydrated, for example to replace fluids and calories being lost in diarrhoea, and when parenteral administration of fluids cannot be carried out. It is generally safe and inexpensive. (B150.w3)
    • Oral rehydration fluids should contain sodium at 60 - 90 mmol/L and glucose at 60 - 110 mmol/L. Preferably, other electrolytes should be present to replace lost potassium, chlorine and bicarbonate as well as sodium. (B150.w3)
    • The fluid should be both inexpensive and palatable. (B150.w3)
    • Also offer plain water. (B150.w3)
  • Subcutaneous: 
    • For individuals which are not severely dehydrated; to prevent dehydration in anorectic patients, and in transition from intravenous to oral fluids. (B150.w3)
    • Not suitable in individuals with marked peripheral vasoconstriction since absorption of fluid will be delayed. (B150.w3)
    • Not suitable for individuals which are hypothermic, severely dehydrated or with acute severe fluid loss. (B150.w3)
    • Fluids need to be isotonic or mildly hypotonic and should be warmed before administration. (B150.w3)
    • Not a suitable route for solutions such as 5% dextrose in water, which lack electrolytes, as electrolytes will move from the extracellular space into the subcutaneous space before the fluid is absorbed. (B150.w3)
    • The amount of fluid which can be given depends on skin elasticity. If large quantities are required, several sites should be used to avoid overdistention of skin, which may cause discomfort. (B150.w3)
    • There is a risk of cellulitis and infection; sterile technique should be used. (B150.w3)
  • Intraperitoneal:
    • For individuals which are not severely dehydrated; to prevent dehydration in anorectic patients, and in transition from intravenous to oral fluids. (B150.w3)
    • Absorption will be delayed in hypotensive or hypothermic individuals. (B150.w3)
    • Fluids need to be isotonic and non-irritating, and should be warmed before administration. (B150.w3)
    • Not effective in treatment of haemorrhagic shock, probably due to constriction of the mesenteric capillaries. (B150.w3)
    • Care is required to avoid injuring abdominal organs when placing a needle or catheter into the peritoneal space. (B150.w3)
    • Note: There is a risk of trauma to viscera, infection, inflammation, leakage of fluid into adjacent subcutaneous tissues, and failure to restore vascular volume. (B150.w3)
  • Intravenous:
    • For treatment of severe dehydration, shock and acute fluid loss. Large volumes of fluids can be given rapidly by this route (unless there is underlying cardiac dysfunction). (B150.w3)
    • Isotonic hypotonic and hypertonic crystalloids, colloids, plasma, blood etc. can all be given by this route. (B150.w3)
    • There is a risk of infection if good sterile technique is not used for catheter placement. (B150.w3)
  • Intraosseous: 
    • For administration of fluids when intravenous access is difficult or cannot be obtained. (B150.w3, J34.23.w1)
    • Note: this route is becoming more popular. (V.w124)
    • All fluids can be given by this route. (B150.w3, V.w124)
    • There is a risk of osteomyelitis if good aseptic technique is not used. (B150.w3)
Temperature of Fluids
  • When giving fluids to small animals in particular, it is important to use warm fluids, since use of cold fluids may contribute to development of hypothermia.
  • This is particularly important in the perioperative period. 
  • Fluids should be pre-warmed e.g. in a water bath or (carefully) by using a microwave oven (J3.159.w4) or by use of an intravenous fluid warmer. (V.w124)
  • A heat retention cover (thermal jacket) is useful to maintain the heat of fluids within an intravenous fluid bag.
  • Note: considerable loss of temperature can occur while fluids are passing through drip tubing.
  • Loss of heat through the giving set can be reduced by pre-warming the giving set then wrapping 30 cm of the giving set around a "hot hand": a large size latex glove, filled with 200 mL water at 38 C, knotted at the wrist, and with the fingers tied to minimise air in the glove.

(J3.159.w4, V.w124)

The following has been provided by Marla Lichtenberger DVM, DACVECC, Milwaukee Emergency Center for Animals and Speciality Services (V.w124)

Perfusion Deficit Corrections

Decompensatory Phase of Shock (Bradycardia, hypotension, hypothermia)

-Slow bolus over 10 minutes of hypertonic saline 7.2/7.5% (3 ml/kg) + Hetastarch (3 ml/kg) IV (intraveous) or IO (intraosseous)

-External and core body temperature warming over 1-2 hrs
-Crystalloids at maintenance (3-4 ml/kg/hr)

-When patient is warmed to 98F (36.7C), use slow IV/IO fluids (see below) to correct indirect systolic blood pressure to >90 mmHg (after each bolus recheck blood pressure-repeat bolus 3-4 times until blood pressure is normal):
1.Crystalloids (LRS, normasol, Plasmalyte) at 15 ml/kg
2.Hetastarch at 3-5 ml/kg

-Unresponsive shock to above protocol:

1.Consider Oxyglobin at 2 ml/kg slow bolus, and if systolic blood pressure is >90 mmHg: 
a.Start crystalloid constant rate infusion (CRI) to correct dehydration or if not dehydrated then at maintenance (3-4 ml/kg/hr)
b.Start Oxyglobin CRI at 0.2 ml/kg/hr
2.If the patient continues to be unresponsive:
a.Check blood glucose, BUN, acid base and electrolytes-correct if abnormal
b.Check packed cell volume (PCV) and total protein consider whole blood transfusion if PCV is < 20 (see blood transfusion in text)
c.Check echocardiogram for abnormal heart function and correct contractility if abnormal
d.Recheck temperature of patient and warm again if hypothermic
3.If the patient continues to be unresponsive and systolic blood pressure is less than 90 mmHg (but patient is normothermic):
a.Consider vasopressors in the doses recommended for small animals (i.e., norepinephrine, doxapram)


Dehydration Deficit Corrections

Estimation of percentage dehydration:
>10%=dry mucous membranes, suction eyes, altered mentation, very significant skin tenting.
7-9%=dry mucous membranes, skin tenting.
5-7%=dry mucous membranes and mild skin tenting.
4-5%=dry mucous membrane .

Fluid requirements of dehydration deficits calculation:
%dehydration x Kg x 1000 ml/L=fluid deficit (L)
This amount is added to maintenance requirements (3-4 ml/kg/hr) + any losses (e.g. diarrhea).
Replacement over how many hours based on how fast the losses occurred:
Losses occurred in <24 hours= acute so replace dehydration deficit over 6-8 hours.
Losses occurred over 24-72 hours=chronic, so replace over 24 hours.


Diuresis Protocol for Acute Renal Failure

A. Correct perfusion deficits if perfusion is inadequate (see above) with crystalloids and colloids.
B. Correct dehydration deficits if present (see above) with crystalloids.
C. Diuresis phase is continued until the azotemia (creatinine and BUN), electrolytes and acid-base are normal.

-Estimate urine output (UO) by urinary catheter volume or diaper weight changes under the small mammal or rodent:
1.UO in mL/hr + Maintenance fluids requirements + losses in ml/hr + add an extra 3-5% ml/kg= total amount of crystalloids needed over an hour.
2.Continue .
D. Maintenance fluids or nasogastric tube feedings until eating and drinking on their own.

(V.w124)

Waterfowl Consideration

FOR BIRDS

"It has been suggested that all birds suffering from trauma and disease can be assumed to be at least ten percent dehydrated." (B13.39.w16)

Fluid therapy is a vital part of initial patient stabilization, whatever the presenting problem of the patient in question. Dehydration and electrolyte losses may be severe and even life-threatening in an ill or injured bird.

AIMS
  • Correct any existing fluid deficit.(B119.w2)
  • Correct and existing electrolyte disorders.(B119.w2)
  • Provide daily requirements.(B119.w2)
CONSIDER
  • Hydration status.(B119.w2)
  • Electrolyte balance.(B119.w2)
  • Acid-base status.(B119.w2)
  • Haematology and biochemistry.(B119.w2)
  • Caloric balance.(B119.w2)
SIGNS OF DEHYDRATION
  • Ulnar artery and vein easily compressible, with small diameter and slow (more than 1-2 seconds) filling time indicates dehydration of more than 7% (B119.w2).
  • Dry mucous membranes (B119.w2).
  • Sunken eyes. (B119.w2)
  • Reduced skin elasticity (B12.39B.w9)
  • General: weakness, reluctance to move, signs of central nervous system depression (B119.w2).
  • Serum osmolarity (B119.w2)
  • Plasma urea (may be greatly increased)(B119.w2)
  • Packed cell volume. May not reflect acute changes in hydration status (B119.w2)
ROUTE
  • Oral, subcutaneous, intravenous or intraosseous. (B119.w2)
  • Oral fluids are appropriate in an individual which is conscious and able to perch/stand and keep its head up. Stressed individuals may tolerate the administration of only small volumes e.g. 5-10ml/kg initially. The frequency of administration may be decreased, and the volume increased gradually. (B119.w2) (See: Gavage / Tubing of Birds)
  • Subcutaneous fluids are appropriate for individuals which are not critically compromised. (See: Subcutaneous Injection of Birds).
  • Intravenous fluids are appropriate for birds with hypotensive or hypovolaemic shock, and for critically ill individuals (See: Intravenous Injection of Birds). (B119.w2)
  • Intraosseous fluids are appropriate e.g. for individuals with very small or collapsed veins (P3.1999b.w2, B119.w2).
  • Intraosseous and intravenous routes provide rapid expansion of circulatory volume and rapid kidney perfusion: these routes should be used in shock or haemorrhage (P3.1999b.w2).
CALCULATION OF REQUIREMENTS
  • Fluid deficit in millilitres may be calculated as equal to normal body weight in grams, times 0.1 (i.e. one tenth of body weight) for estimated 10% dehydration.
  • Maintenance fluids required may be estimated at 50ml/kg/day (40-60ml/kg/day).
  • Fluid therapy should be given to correct 1/4 to1/2 of the deficit within the first 4 to 6 hours, with the remaining replacement volume given over the following 20 to 28 hours (B119.w2); correction of 50% of the estimated deficit in the first 24 hours and the remainder of the deficit over the following 48 hours (together with the maintenance requirement) (B11.3.w10).
  • "Rule of thumb": 3% of body weight three times daily (allows for 10% dehydration plus maintenance requirements (P7.1.w6). Normal saline warmed to 39 C may be given by subcutaneous injection at 20.5ml/kg body weight four to five times daily (B14).
  • N.B. Maximum acute fluid load tolerated by a healthy individual is 90ml/kg/hour, but most avian patients cannot tolerate such a high rate.
  • Lactated Ringer's solution is the fluid of choice, protecting renal function better than do sugar solutions. 5% dextrose is not considered a satisfactory replacement solution since free water is left as the dextrose is metabolized.
  • Fluids should be warmed to 96 F (37 C) before being administered to anaesthetized birds, to avoid hypothermia. (B13.39.w16); (38-39 C (100.4-102.2 F)) (P3.1999b.w2).
Crane Consideration
  • Depending on the crane's level of dehydration, electrolyte imbalance, emaciation, stress or shock, oral, subcutaneous or intravenous routes of fluid therapy may be appropriate. (B12.56.w14)
  • For wild cranes coming in for rehabilitation, dehydration is common. The hydration status should be checked and fluids given immediately if the bird is dehydrated. (J311.21.w1)
Types of fluids
  • Depending on the crane's physiological state (dehydration, electrolyte imbalance etc.), 2.5% dextrose, Ringer's solution, lactated Ringer's solution (Hartmann's), 5% dextrose, normal saline etc. may be appropriate. (B12.56.w14)
  • Lactated Ringer's solution can be used for this (similar to avian plasma) or normal saline. (B115.8.w4)
Shock treatment
  • In the treatment of shock, use of bolus intravenous fluids can be useful. (B115.8.w4)
Rehydration
  • The basic fluid requirement is 44 mL/kg bodyweight per day, plus replacement of losses due to diarrhoea and additional requirements if the crane is dehydrated. (B12.56.w14, B115.8.w4)
    • Assume 10% dehydration of dehydration is evident. (B115.8.w4)
  • About 50% of the deficit should be administered in the first 12 hours of treatment and the remainder, together with the crane's maintenance requirements (calculated) over the next two days. (B115.8.w4)
  • As a rule of thumb, 3 - 5% of body weight of a fluid such as lactated Ringer's solution should be given initially as an intravenous bolus, subcutaneously or intraosseously. (J311.21.w1)
  • Keep fluids warm using an incubator or water bath. (B115.8.w4)
Correction of acidosis
  • Acidosis may be present in cranes with dehydration and capture myopathy. B115.8.w4
  • The presence and level of acidosis ideally should be confirmed by laboratory testing.(B115.8.w4)
  • To treat known or suspected accidosis, give sodium bicarbonate, 1 mEq/kg bodyweight subcutaneously with fluids, every 30 minutes to a maxmum of 4 mEq/kg. (B115.8.w4)
Correction of hypoglycaemia
  • Rarely needed in adult cranes, but in young chicks this must be considered. (B115.8.w4)
  • Correction of hypoglycaemia may be critical for the survival of young chicks. (B115.8.w4)
  • Glucose or dextrose should be given orally or parenterally. (B115.8.w4)
Routes
  • Intravenous. The initial dose of fluid calculated can be given as a slow intravenous bolus injection, generally into the jugular or median tarsal vein (alternatives are the brachial or ulnar veins). (B115.8.w4) See: Venipuncture in Cranes (Techniques)
  • Subcutaneous. See Subcutaneous Injection of Birds (Techniques)
    • Not suitable for critical care, but can be used otherwise. (B115.8.w4)
    • The main sites used are the flank just in front of the legs, the intrascapular area and the base of the neck. (B115.8.w4) 
    • This route can be used repeatedly with minimal trauma to the crane. (B115.8.w4)
    • Absorption is relatively slow: up to 30 minutes, and sometimes much longer in a severely debilitated crane. (B115.8.w4)
  • Intraosseous. (B115.8.w4)
Bear Consideration Fluid therapy can follow the usual recommendations for domestic animals such as cats and dogs.
  • Bears may require fluids when obviously dehydrated (B16.9.w9, B64.26.w5, P62.18.w1) or while undergoing prolonged surgical procedures. (P3.2006a.w1, P62.18.w1)
    • Intravenous catheters may be placed in the cephalic (P62.18.w1) or medial saphenous (V.w6) veins to give fluids intravenously. 
    • Lactated Ringer's solution and 5% dextrose may be given as fluid support during surgery. (P62.18.w1)
  • Fluids may be given as supportive treatment for bears which have diarrhoea and vomiting and/or are refusing food and water. (J3.145.w4, J11.83.w1, J142.19.w1) 
    • Cubs with bacterial gastroenteritis [Bacterial Gastroenteritis in Bears] may need intravenous or subcutaneous fluid therapy with lactated Ringer's solution (Hartmann's Solution). (B64.26.w5)
    • In cases of severe dehydration with gastritis, supportive fluid therapy intravenously: (B16.9.w9, B64.26.w5) isotonic saline or saline dextrose, 50-100 mL/kg body weight (B64.26.w5). 7.5-10 mL per kg body weight. (B16.9.w9)

Lagomorph Consideration

Fluids may be useful:
  • In obviously dehydrated rabbits.
  • In rabbits with diarrhoea.
  • In rabbits with gastric stasis.
  • Prior to anaesthesia, if a fluid deficit is suspected. (J15.20.w2)
  • During surgery, and after if significant blood loss has occurred.
  • Immediately after surgery to avoid development of dehydration before the rabbit returns to drinking normally.

(B600.10.w10, B601.8.w8, B601.3.w3, B609.2.w2, J15.20.w2, J60.8.2, J213.8.w2)

Types of Fluid
  • Crystalloids are commonly used; these are inexpensive and physiological.
  • Colloids are useful in hypoproteinaemic rabbits (normal total protein 5.4 - 7.5 g/dL) or when crystalloids are ineffective to restore blood pressure. (J213.1.w1, J215.21.w3)
  • Oxyglobin can be given to rabbits in small volume boluses, but not in large volume boluses as used in the dog. (J29.13.w1)
Route
  • Fluids may be given orally or by subcutaneous, intraperitoneal, intravenous or intraosseous administration. (J15.30.w2)
    • Oral: divide the total into four to six doses. Give slowly to avoid inhalation. (J15.30.w2)
    • Subcutaneous: large volumes can be given, but absorption from this site is slow. (J15.30.w2)
    • Intraperitoneal: ensure aseptic technique. Absorption is rapid. (J15.30.w2)
    • Intravenous. Crystalloids, colloids and blood can be given by this route. The lateral saphenous, marginal ear vein or cephalic vein can be used. If giving fluids in boluses, divide up the daily total into several doses. Alternatively, give by continuous infusion. (J15.30.w2)
    • Intraosseous: Crystalloids, colloids and blood can be given by this route. It is useful in collapsed/hypotensive  individuals and those with very small or fragile veins. (J15.30.w2, J215.21.w3)
  • An infusion pump can be used to ensure an accurate flow rate for continuous infusion. For small rabbits, a syringe pump may be needed. (J213.1.w1)
Initial Calculation of Requirements
  • Fluid maintenance rate for rabbits is 120 mL/kg/day. (J15.30.w2); 75 - 100 mL/kg/day (B601.3.w3, P113.2005.w3)
  • For a debilitated rabbit, assume about 10% dehydration. (P113.2005.w3)
    • In general, replace 50% of the deficit (plus ongoing required volumes for maintenance and ongoing losses over the first 12 hours, then replace the remainder of the deficit (with ongoing maintenance etc. over 48-72 hours. (P113.2005.w3)
    • Required volume for rehydration (litres) = hydration deficit x body weight (kg) x 1000. (P113.2008.w1)
      • As much as 80% of the calculated deficit can be administered in the first 24 hours. (P113.2008.w1)
      • Once calculated losses are replaced, continue giving maintenance fluids until the rabbit is taking in fluids normally and maintaining hydration. (P113.2008.w1)
  • Note: when giving fluid therapy to treat a deficit (e.g. hypovolaemia or dehydration) it is important to continue treatment (and if necessary amend treatment) until the required effect (correction of the hypovolaemia or dehydration) has been produced, not simply give according to a formula then stop. (J29.13.w1)
  • Blood loss
    • Rabbits have a blood volume of about 60 mL/kg (J29.13.w1) (55 - 65 mL/kg (B601.3.w3) 50 - 60 mL/kg (P113.2008.w1) (compared with 90 mL/kg in dogs). (J29.13.w1, P113.2008.w1)
    • Loss of more than 20 - 25% of blood volume causes shock. (B601.3.w3)
    • If PCV falls below 12% (J213.1.w1) 10-15% (B601.3.w3), give a blood transfusion. (B601.3.w3, J213.1.w1)
    • No blood typing has been established for rabbits. (B601.16.w16, J213.1.w1)
    • In practice, acute transfusion reactions in response to an initial transfusion from a single donor are rare. (B601.3.w3, B601.16.w16)
    • Blood collection: 
      • Ideally the blood donor should be:

        (B601.3.w3)

      • Up to 1% bodyweight in blood can be taken from the donor. (B601.3.w3)
      • Blood should preferably be taken from the jugular. (B601.3.w3)
      • Collect with citrate phosphate dextrose (CPD) adenine anticoagulant at 0.14 mL per 1 mL blood. (B601.3.w3)
      • Ideally transfuse within 4 - 6 hours. Can be stored at 4 - 6 C for up to 28 - 35 days. (B601.3.w3)
      • Whole blood can be collected and stored with 1 part acid citrate dextrose (ACD) to 3.5 parts whole blood. (B601.16.w16)
    • Blood transfusion
      • Whole blood can be given at 10 - 20 mL/kg (amount based on cat and dog medicine). (B601.3.w3)
      • Initially give 0.25 mL/kg over 15 minutes and check for transfusion reactions. (B601.3.w3)
      • Maximum rate of 22 mL/kg/hr. (B601.3.w3)
      • Any vein which can be used for intravenous injections (e.g. marginal ear vein) can be used. (B601.3.w3)
      • Preferably use an in-line filter; direct injection without a filter can be used in an emergency. (B601.3.w3)
      • Monitor respiration and heart rate. (B601.3.w3)
      • Monitor for rigor, jaundice, abnormal bleeding (due to intravascular coagulation) and signs of renal impairment, indicating transfusion reaction. (B601.3.w3)
  • During surgery: 
    • If loss of blood or fluids is expected, initially give 10 - 15 mL/kg/hr of warmed lactated ringers solution (Hartmann's). (B601.16.w16)
    • Monitor blood loss by weighing swabs. Whole blood can be given if severe blood loss occurs (see above), or Oxyglobin can be given if blood is not available. (B601.16.w16)
    • If whole blood is unavailable, Oxyglobin can be given to rabbits in small volume boluses, but not in large volume boluses as used in the dog. A suggested rate is 2 mL/kg over 10 - 15 minutes until heart rate returns to normal and systolic blood pressure rises above 90 mm Hg, then 0.2 - 0.4 mL/kg/hr as a constant rate infusion (CRI). (J29.13.w1)
      • The CRI of Oxyglobin can continue for up to 24 hours (remembering that Oxyglobin cannot be given more than 24 hours after opening due to conversion to methaemoglobin). (V.w124)
  • For hypovolaemic shock: 
    • In hypovolaemic shock, a rabbit generally has a heart rate under 200 bpm, hypotension (systolic blood pressure under 90 mm Hg) and hypothermia (body temperature less than 36.6 C/98 F). The mucous membranes may be white or grey, capillary refill absent, and the pulse weak or not palpable. (J29.13.w1, P113.2008.w1)
    • Give a combination of crystalloids and colloids: isotonic crystalloids at 10 - 15 mL/kg as a rapid infusion, plus colloids (Hetastarch suggested) at 5 mL/kg over a period of 5 - 10 minutes. (J29.13.w1, P113.2008.w1)
    • Once the systolic blood pressure is above 40 mm Hg, continue maintenance crystalloids plus patient warming over 30 - 60 minutes using e.g. warm water bottles or warm forced air, and warmed fluids. (J29.13.w1, P113.2008.w1)
    • Once the rabbit's core body temperature reaches 99 F, check the blood pressure and give more colloid (hetastarch) as required: 5 mL/kg at a time over 15 minutes, until systolic blood pressure increases to above 90 mm Hg.
    • Once this level is reached, monitor the rabbit. Often maintaining body temperature and giving warmed maintenance crystalloids is all that is required. If blood pressure drops again, another 5 mL/kg hetastarch may be given, followed by a constant rate infusion of 3 - 5 mL/kg/hr of hetastarch. (J29.13.w1)
    • Note: use of crystalloids alone, without colloids, to treat hypovolaemia, "can result in significant pulmonary and pleural fluid accumulation." (J29.13.w1, P113.2008.w1)
    • Alternative:
      • Give 100 mL/kg (one blood volume) crystalloids over a period of one hour. (J213.1.w1, P113.2005.w3)
        • Carefully monitor pulse and heart rate. (J213.1.w1)
Temperature of fluids
  • Give fluids warm to avoid development of hypothermia. (J15.30.w2, J213.1.w1, P113.2005.w4)
  • NOTE: In the treatment of shock, raising the rabbit's body temperature to the normal range (it is likely to be hypothermic) is an important part of treatment, since low body temperature appears to make the adrenergic receptors refractory to catecholamines, probably preventing compensatory mechanisms such as vasoconstriction in response to hypovolaemia. (J29.13.w1)
Ferret Consideration
  • Fluid requirements are 75-100 mL per kg per day, plus allowances for losses due to vomiting, diarrhoea etc., losses during surgery, and any pre-existing dehydration. (J15.24.w5)(J15.24.w5)
Route
  • Fluids may be given subcutaneously or intravenously. (B602.2.w2)
  • In a mildly dehydrated ferret, fluids can be given subcutaneously over the dorsum (under the scruff). (B631.18.w18, P120.2006.w6)
    • If giving subcutaneously, give fluids divided into two or three sessions (give every 8-12 hours). (B602.2.w2, P120.2006.w6)
    • Note: when fluids are given subcutaneously, the ferret may find this painful; good restraint is important so the ferret does not bite anyone. (B602.2.w2)
  • Fluids can be given as a bolus by intraperitoneal injection: with the ferret held vertically, insert the needle into the caudolateral abdomen, with the needle held at a 45 degree angle. Up to 30 mLkg may be given by this route at one time. This can be useful perioperatively, and provides rapid absrption, but if carried out repeatedly the ferret may start resenting the procedure and struggling (increasing the risk of iatrogenic injury). This route should not be used in a ferret with an intra-abdominal mass or intra-abdominal fluid. (B631.18.w18)
  • For severely dehydrated or very ill ferrets administer fluids intravenously or by the intraosseous route using a fluid pump. (B602.2.w2, B631.18.w18, P120.2006.w6)
    • Continuous rate infusion is preferred if available. (B602.2.w2)
    • Alternatively, the daily requirement can be divided into two or three and delivered using a syringe pump or a Buretrol (Baxter Healthcare, Glendale, California, USA). (B602.2.w2)
Crystalloids
  • For maintenance, crystalloids should be given at 60-70 mL/kg per day. (B602.2.w2, P120.2006.w6) 70 ml/kg/day (J29.6.w3) 75-100 mL/kg per day. (B232.18.w18)
    • Additional fluids should be given as required to correct any dehydration and/or allow for additional ongoing losses. . (B602.2.w2, P120.2006.w6)
    • In shock, 60 mL/kg can be given over the first hour, before reducing to maintenance rates. (B631.18.w18)
  • Give potassium, dextrose (2..5 - 5%), B-vitamins and other supplements as required - use the same clical criteria and calculations as in a cat or dog. (B602.2.w2, P120.2006.w6)
  • For middle-aged and older ferrets (in which Insulinoma in Ferrets are common) undergoing surgery, consider using 2.5-5% dextrose in saline, rather than lactated Ringer's solution. (J29.6.w3)
Colloids
  • For a ferret with hypoproteinaemia or shock, give a colloid, such as hetastarch (Hydroxyethyl starch) at 10-20 mL/kg/day  intravenously. (B602.2.w2, P120.2006.w6)
    • If also giving crystalloids, reduce the volume of crystalloid fluids by 33-50%. (B602.2.w2, P120.2006.w6)
    • In severe shock, give 5 mL/kg intravenously as a bolus over a period of 15 minutes. Repeat as needed, up to a maximum of 20 mL/kg/day. (B602.2.w2, P120.2006.w6)
Blood transfusion
  • Blood transfusion may be needed if the ferret:
    • Is anaemic due to blood loss, chronic disease, or oestrogen toxicosis, also in thrombocytopaenia. (B602.2.w2)
    • Has a PCV under 25% and shows clinical signs of anaemia (e.g. tachycardia, tachypnoea, weakness), or is to undergo surgery. (B602.2.w2, P120.2006.w6)
    • Is thrombocytopaenic with petechiae, ecchymoses or bleeding evident. (B602.2.w2, P120.2006.w6)
    • If major blood loss occurs during surgery and the haematocrit falls below 25%. (B631.22.w22)
  • Ferrets have no detectable blood groups, therefore there appears to be no risk from transfusing blood between ferrets, with no cross-matching required. (B232.18.w18, B602.2.w2, J4.197.w2, J29.6.w3) 
    • Ferrets can be transfused with blood from several donors if necessary. (J29.6.w3)
    • Consider pretreating with corticosteroid (J513.2.w3), e.g. prednisolone sodium succinate 22 mg/kg intravenously, or dexamethasone sodium phosphate 4 - 8 mg/kg intravenously or intramuscularly. (P120.2006.w6)
  • Collect blood from the jugular vein of the donor, using a butterfly catheter and a syringe containing an anticoagulant - either heparin or acid-citrate-dextrose. (J29.6.w3)
    • A 1 kg ferret has about 100 mL blood. Up to 10% of that (i.e. 10 mL) can safely be taken. (J29.6.w3)
    • Large male ferrets are preferred as blood donors. (B602.2.w2, P120.2006.w6)
    • Use 1 mL of acid-citrate-dextrose anticoagulant to 6 mL blood. (B232.18.w18, B602.2.w2, P120.2006.w6)
  • Give the blood to the recipient ferret immediately (J29.6.w3); use a filter. (B602.2.w2); give 6 - 12 mL. (B232.18.w18)
Blood alternatives
  • Haemoglobin-based oxygen-carrying solutions can be used, e.g. Oxyglobin (Biopure Corp., Cambridge, Massachusetts, USA). (B602.2.w2, P120.2006.w6)
  • This can be given at 11-15 mg/kg infused over a period of four hours. (B602.2.w2, J513.2.w3, P120.2006.w6)
  • Over a period of 24 hours, this dose can be given twice in total if needed. (B602.2.w2, P120.2006.w6)
  • No donor is required, and there is no delay while blood is collected from a donor. (J513.2.w3)
  • No filter is needed for administration. (B602.2.w2)
  • Oxyglobin can be given through a smaller catheter. (J513.2.w3)
  • Note: Oxyglobin has colloid properties. To avoid volume overload, it is important to control of the rate of administration in normovolaemic anaemia or when the blood volume loss is undetermined. (J513.2.w3)
    • Particular care is required in patients with severe cardiac function impairment, renal impairmens with oliguria or anuria, or individuals predisposed to pulmonary oedema development. (J513.2.w3)
  • Individuals given oxyglobin develop dose-dependent yellow-orange colouration to the skin, sclera and mucous membranes, and red-brown urine; this resolves in three to five days. (J513.2.w3)
  • Once Oxyglobin has been given, oxygen carrying capacity of the blood needs to be measured on the basis of total haemoglobin concentration, not just PCV. (J513.2.w3)
  • Carry out serum chemistry tests before giving Oxyglobin, since test results can be artifactually increased or decreased in th presence of Oxyglobin (results depend on the reagents and analyzer used). (J513.2.w3)
  • The colloid properties as well as the oxygen-carrying properties are beneficial in individuals with, for example, gastro-intestinal bleeding due to an ulcer. (J513.3.w3)
Treatment of Hypovolaemic Shock

In hypovolaemic shock, usually the ferret will have a heart rate under 200 bpm, systolic blood preeure less than 90 mmHg, hypothermia under 36.6 C (98 F), grey or white mucous membranes, no visible capillary refill, and will show profound mental depression. (J513.7.w3)

  • Initially give an isotonic crystalloid via an intravenous or intraosseous catheter. (J513.7.w3)
  • Next give Hetastarch (6%), 5 mL/kg over a period of 5 - 10 minutes. (J513.7.w3)
  • If the ferret's systolic pressure is at least 40 mm Hg, aggressively warm the ferret (using a forced air blanket, hot water bottles or an incubator) while giving warmed fluids at maintenance rates. (J513.7.w3)
    • Hypothermia appears to play a significant role in hypovolaemic shock. Rewarming is an important component of shock treatment. (J513.7.w3)
    • Always monitor body temperature closely during rewarming to avoid hyperthermia. (J513.7.w3)
  • Check cardiac function; correct any blood glucose, acid-base and electrolyte abnormalities. (J513.7.w3)
  • For non-responsive shock, consider giving Oxyglobin, 2 mL/kg over 10-15 minutes, repeated as necessary to produce systolic blood pressure of over 90 mm Hg. Usually two such injections are needed. (J513.7.w3)
  • Once blood pressure and heart rate have normalised, give crystalloids at 2 mL/kg/hour (maintenance rate) plus oxyglobin at 0.2 - 0.4 mL/kg/hour as a constant rate infusion for about 24 hours. (J513.7.w3)
  • Note: using crystalloids alone may result in pulmonary and pleural fluid accumulation with resultant hypoxaemia. (J513.7.w3)
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and health care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
  • Fluid therapy is commonly a critical need in debilitated primates, particularly those with diarrhoea, (B10.44.w44g, D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
Choice of fluid therapy route
  • In a primate with mild dehydration (reduced urinary output, increased thirst), provide oral rehydration solution or maintenance solution, giving 10 ml/kg/hour every four hours; an additional 5-10 mL/kg may be given after each episode of diarrhoea. (D426.2.11.w2k)
    • In infants, mild dehydration may be treated by increased oral intake, but the sicker the infant, the more difficult it is to get it to drink. Oral rehydration solution can also be given by stomach tube if the infant is not vomiting. (B678.w8)
  • In a primate with moderate dehydration (eyes sunken, somewhat reduced skin turgor (abdominal skin tents for less than two seconds when lifted and released), buccal mucous membranes dry), oral rehydration solution should be given at 15-20 mL/kg/hour and the primate re-assessed every four hours. (D426.2.11.w2k)
  • In a primate with severe dehydration (eyes sunken, reduced skin turgor with tenting of the skin for more than two seconds, buccal mucous membranes dry, capillary refill time prolonged, with or without signs of shock: rapid breathing, lethargy, rapid thready/weak pulse, cool extremities or coma, the individual's blood pressure should be measured and intravenous fluid therapy started: (D426.2.11.w2k)
    • Administer fluids via a large bore indwelling intravenous catheter. (D426.2.11.w2k)
    • Use saline, plasma or colloids. Infusion of 10-20 mL/kg over 30 minutes if necessary, to restore circulating blood volume.  (D426.2.11.w2k)
Oral rehydration
  • If possible: if the primate is not vomiting excessively, and the kidneys are working effectively, then oral rehydration is preferable. This route is commonly used in primates with diarrhoea. (D426.2.11.w2k)
  • In a primate with diarrhoea but normal urinary output and no clinical signs of dehydration, fluids should be given ad libitum. Oral rehydration solution should be offered, while undiluted fruit juice and other high osmolarity fluids should be avoided. (D426.2.11.w2k)
    • In most cases, oral rehydration therapy is effective in the treatment of watery diarrhoea of whatever cause. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
    • Oral rehydration fluids contain glucose as well as sodium. Glucose is absorbed through the intestinal wall even when this is damaged, and a co-transport system means that sodium is also absorbed, which pulls water through as well. (D425.3.14.w3n)
    • Ideally, the oral rehydration solution contains glucose and sodium in a 1:1 ratio of molarity. (D425.3.14.w3n)
  • Limitations: oral rehydration is not suitable for individuals with: (D425.3.14.w3n,  P3.2005b.w2)
    • protracted vomiting despite frequent small feeds.
    • diarrhoea which is getting worse, with oral rehydration volumes insufficient to match losses.
    • stupor or coma
    • intestinal ileus.
Parenteral fluid therapy
  • Intravenous rather than oral fluids will produce faster improvement in the clinical condition of dehydrated infants. (B678.w8)
  • If intravenous catheterisation is difficult due to low blood pressure, a sterile cut-down onto the vein is recommended. (D425.3.14.w3n, P3.2005b.w2)
  • Where potassium loss is a feature of dehydration, as long as 12-24 hours may be needed between oral administration and clinical improvement. (B678.w8)
  • Intraosseous administration of fluids may be required if intravenous access cannot be obtained. Note: in humans, intraosseous administration of fluids is known to be extremely painful. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
    • There is also a higher risk of infection. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
    • This route may be useful in an anaesthetised individual. (P3.2005b.w2)
  • Subcutaneous administration of fluids is a poor choice. Apes have little subcuticular space in which to place fluids, and individuals with low circulating blood volume or acidosis will have constriction of blood vessels supplying the skin, therefore fluids given by this route will not be absorbed. Note that e.g. 5% dextrose is contraindicated because it will actually cause vasoconstriction and draw fluid out of the vessels into the subcuticular space. (D426.2.11.w2k)
    • This route can be used for maintenance fluids. (P3.2005b.w2)
    • Only isotonic fluids should be used.  (P3.2005b.w2)
    • 5% dextrose is not suitable for this route: the extracellular fluid volume needs to equilibrate with the electrolyte-free fluid, which may make electrolyte imbalances worse. (P3.2005b.w2)
  • The intraperitoneal route can be used if no other route is available. However, the peritoneal cavity is normally only a potential space, therefore there is a high risk of the needle piercing an organ when introduced into the abdomen. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
    • There is a risk of chemical peritonitis (depending on the pH of the fluid)
    • There is a risk of infection if aseptic technique is inadequate.
    • Uptake of fluid is not well controlled.
    • Fluid given by this route which has not been appropriately warmed risks causing shock or vomiting.
Choice of fluid for parenteral fluid therapy
  • Fluid choice for the parenteral treatment of dehydration depends on the losses involved. (D425.3.14.w3n, D426.2.11.w2k, P3.2005b.w2)
  • In most situations, lactated Ringer's solution (Hartmann's solution) is appropriate for parenteral fluid therapy. (D425.3.14.w3n, P3.2005b.w2)
    • Not suitable in an individual with pulmonary or cerebral oedema.
    • For an individual with congestive heart failure, use 0.45% saline plus 2.5% dextrose, or use half-strength LRS plus 2.5% dextrose.
    • For an individual with ruptured bladder use 0.9% saline.
  • Haemorrhage (all blood components lost). For replacement of mild blood loss, colloids can be used (and crystalloids). For severe blood loss, fresh whole blood is preferable if this is available.
  • Dehydration (due to insufficient fluid intake; water lost). Replace with sodium chloride 0.18% plus+ dextrose 4%, or with dextrose 5%. Add potassium chloride 10 - 20 mmol/L after two days of rehydration.
  • Vomiting (water, sodium, potassium and chlorine lost, acid lost). Sodium chloride 0.9%, or Ringer's solution. Add potassium chloride 10-20 mmol/L after two days.
  • Diarrhoea (water, sodium, potassium and chlorine lost, bicarbonate lost). Oral fluids using an oral rehydration solution. Lactated Ringer's solution (Hartmann's). Add potassium chloride 10 - 20 mmol/L after two days of rehydration.
  • Severe vomiting and diarrhoea (water, sodium, potassium and chlorine lost, bicarbonate lost). Colloid plus lactated Ringer's solution (Hartmann's).
  • Peritonitis (plasma and extracellular fluid). Colloid plus lactated Ringer's solution (Hartmann's).
  • Gastrointestinal obstruction (water, bicarbonate, sodium and chloride lost). Colloid plus lactated Ringer's solution (Hartmann's).
  • Urethral obstruction (potassium and acid retention). Sodium chloride 0.9% plus dextrose 5%.
Additional components
  • Treatment of acidosis: Sodium bicarbonate, 1 meq/kg intravenously over a period of 10-15 minutes. (D425.3.15.w3o)
  • Treatment of hypokalaemia: Potassium chloride 20 - 40 meq per litre of fluids intravenously. Maximum 1 meq per kg per minute. (D425.3.15.w3o)
Associated techniques linked from Wildpro Birds

Mammals

Bears

Rabbits

Ferrets

Bonobos

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Analgesia

Analgesia - pain relief - is an important part of treatment of the individual animal.

Pain and associated distress is a welfare problem. It is generally agreed that we have a duty to minimise the pain and suffering of animals that are under our care. (J4.221.w2) The "Five Freedoms" described by the Farm Animal Welfare Council in the UK include "Freedom from pain, injury or disease" and "Freedom from fear and distress."  (J35.161.w2, W550.Dec04.w1) The same Five Freedoms are accepted as principles providing a framework for the Secretary of State's Standards of Modern Zoo Practice (D15 - full text included in Wildpro) which zoos in the UK are expected to adhere to. It is therefore expected that pain will be prevented if possible and rapidly diagnosed and treated if it occurs. (J35.161.w2)

  • Veterinary surgeons in the UK (members of the Royal College of Veterinary Surgeons) have a responsibility to ensure the welfare of animals under their care, including a specific responsibility to their patients, set out in the RCVS Guide to Professional Conduct, not to cause any patient to suffer "by failure to maintain adequate pain control and relief of suffering." (D155.1C.w3)
Advantages of analgesia in the treatment of animals include:
  • Improved comfort. (J16.28.w1)
  • Faster recovery from surgery or trauma. (J16.28.w1, J16.36.w1, J288.59.w2, B207.2.w2, B322.1.w1, P53.24.w1)
  • Faster return to normal intake of food and water. (J16.28.w1)
  • Reduced susceptibility to disease. (J284.71.w1)
  • Avoidance of development of hyperalgesia and chronic pain. (J16.36.w1; B207.2.w2, P53.24.w1)
  • Reduced stress. (J4.221.w3, B207.2.w2)

NOTE: It is important to remember that pain, and suffering which may be caused by unrelieved pain, are associated with maladaptive physiological responses and behaviours: "there are no beneficial effects of unrelieved pain in animals under veterinary care." (J4.213.w2, W513.Jun04.w1)

Analgesia should be given whenever an animal has an injury which would, in domestic animals, be expected to be painful, or is subjected to a procedure which damages tissues and/or would generally considered to be painful, or displays abnormal behavioural responses. (J29.15.w1, V.w5)

  • Note: it can be difficult to assess pain in animals, particularly prey species. (J213.10.w2, J29.15.w1, J290.21.w1, J303.7.w1)
  • Observer experience is important for accurate assessment of pain. (P54.2.w16)
  • An understanding of the normal behaviour for the species is required in order to properly interpret behavioural signs of pain. (B322.4.w4)
    • Knowing the normal behaviour of the individual is also important. (J29.15.w1)
  • Wild animals are very likely to hide signs of pain even when presenting with severe injuries. (J34.24.w2)

Pre-emptive analgesia should be given before a painful procedure, to avoid sensitisation. (J29.15.w1)

Multimodal analgesia (combined therapy) is recommended. This involves the use of different types of analgesic agents acting at different points along the pain pathway. (J213.10.w2)

  • Use of multimodal analgesia allows use of a smaller dose of each drug, since their actions are additive or synergistic; this reduces undesirable side effects (which are usually dose-dependent). (J213.10.w2)
  • Further information on different types of analgesics is given in Groups of Analgesic Drugs
Crane Consideration
  • For minor procedures (e.g. wound suturing, implantation of subcutaneous transmitters), infiltration of local anaesthetic into the area can be used: lidocaine 0.5 mL, xylocaine 0.5 mL or bupivacaine up to 2 mg/kg. (B703.10.w10)
    • Small amounts should be used, e.g. 0.25 - 0.5 mL lidocaine in an adult care. (B115.8.w4)
  • Use of local anaesthetic agents at surgical sites may reduce post-operative pain. (B703.10.w10)

Lagomorph Consideration

  • Pain management is very important in rabbit medicine, particularly critical care, since pain can affect the responses of the rabbit to care, and can affect the rate of recovery. (J213.1.w1)
  • Unfortunately it is probable that rabbits seen in general veterinary practice commonly are still not given adequate analgesia, as indicated by the results of a 1999 survey. (J3.145.w3, J213.4.w5)
  • In wild lagomorphs, pain management is just as important as in domestic rabbits; the same analgesics and doses should be used as in domestic rabbits. (B284.10.w10)
Analgesics used in rabbits

Both NSAIDs and opioid analgesics are used in rabbits.

  • Buprenorphine (Opiate analgesic)
    • Mixed agonist/antagonist properties. (B600.5.w5)
    • Can be given at the start of anaesthesia as pre-emptive analgesia before potentially painful procedures. (B600.5.w5)
      • May reduce the amount of volatile anaesthetic agent (e.g. Isoflurane) required to maintain anaesthesia.
    • May be sedative at higher doses. (J15.30.w2)
    • Dose rate and route: 
      • 0.01 - 0.05 mg/kg subcutaneously or intravenously. (B600.5.w5, B601.16.w16)
      • 0.01 - 0.05 mg/kg subcutaneously. (J15.20.w2)
      • 0.01 - 0.05 mg/kg subcutaneously, intravenously or intramuscularly. (B604.3.w3, J15.30.w2, J29.15.w1)
    • Duration of action 
      • Provides analgesia in rabbits for seven hours. (B600.5.w5)
      • 6 - 12 hours. (B600.5.w5, B601.16.w16, J15.30.w2, J29.15.w1, J290.17.w2)
    • Note: onset of analgesia after 30 minutes, therefore if used for post-operative pain relief, this should be given before recovery from anaesthesia. (J290.17.w2)
  • Butorphanol (Opiate analgesic)
    • Mixed agonist/antagonist properties. (B600.5.w5)
    • Provides analgesia and mild sedation. (B600.5.w5)
    • Dose rate and route:
      • 0.1 - 0.5 mg/kg subcutaneously (B601.16.w16, J15.20.w2, B604.3.w3) or intravenously. (B600.5.w5, B601.16.w16, B604.3.w3) or intramuscularly. (B600.5.w5, B604.3.w3)
      • 0.1 - 1.0 mg/kg subcutaneously, intramuscularly or intravenously. (J29.15.w1)
    • Duration of action 2 - 4 hours. (B600.5.w5, B601.16.w16, J29.15.w1, J290.17.w2)
      • Longer elimination half-life following subcutaneous administration than following intravenous administration. (B600.5.w5, J13.53.w1)
  • Morphine (Opiate):
    • 2.5 mg/kg subcutaneously or intramuscularly every four hours. (J15.30.w2, J29.15.w1)
  • Pethidine (Opiate) (Meperidine) 
    • Opioid agonist, mainly acting on mu receptors. Relatively short acting/ Less potent than morphine. (B600.5.w5)
    • May be useful when other opioid are not available. (B600.5.w5)
    • Dose rate and route: 
      • 5 - 10 mg/kg intramuscularly or subcutaneously. (B600.5.w5, B601.16.w16)
      • 5 - 10 mg/kg subcutaneously or intraperitoneally. (B604.3.w3)
    • Duration of action 2 - 4 hours. (B601.16.w16); 2-3 hours. (B600.5.w5, B604.3.w3)
  • Nalbuphine (Opiate)
    • 1 - 2 mg/kg intravenously. Duration of action about 4 hours. (B601.16.w16)
    • 1 - 2 mg/kg intramuscularly or intravenously every 2 - 4 hours. (J29.15.w1)
  • Aspirin (NSAID)
    • 100 mg/kg orally every 48 hours. (B604.3.w3
    • 100 mg/kg orally as "first aid" pain relief. (B284.10.w10)
  • Carprofen (NSAID)
    • 1.5 mg/kg orally or 1-2 mg/kg subcutaneously or intravenously. Duration of action possibly 24 hours. (B601.16.w16)
    • 1.5 mg/kg orally twice daily. (B600.5.w5, J15.20.w2)
    • 4 mg/kg subcutaneously once daily. (B600.5.w5)
    • 1.0 - 2.2 mg/kg orally every 12 hours. (J15.30.w2)
    • 1.0 - 2.0 mg/kg subcutanelously or orally every 12 hours. (J29.15.w1)
  • Flunixin meglumine (NSAID)
    • 1.1 mg/kg subcutaneously. Duration of action possibly 12-24 hours. (B601.16.w16)
    • 1.1 mg/kg subcutaneously twice daily. (J15.20.w2)
    • 1.1 mg subcutaneously or intramuscularly every 12 - 24 hours. (J15.30.w2)
  • Ibuprofen (NSAID)
    • 10 - 20 mg/kg orally; lasts four hours. (B604.3.w3)
  • Ketoprofen (NSAID)
    • 3 mg/kg intramuscularly. (B601.16.w16)
    • Duration of action possibly 12 - 24 hours. (B600.5.w5, B601.16.w16)
    • 3 mg/kg intramuscularly. (J15.20.w2)
    • 1 - 3 mg/kg subcutaneously twice daily. (B600.5.w5)
    • 1.0 mg/kg intramuscularly or subcutaneously every 12-24 hours. (J15.30.w2)
    • 1 mg/kg intramuscularly every 12 hours. (J29.15.w1)
    • Use with care in hypotensive individuals. (B284.10.w10, B600.5.w5)
  • Meloxicam (NSAID)
    • 0.2 - 0.6 mg/kg orally or subcutaneously. Duration of action possibly 24 hours. (B601.16.w16)
    • 0.3 - 0.6 mg/kg every 24 hours. (J15.30.w2)
    • 0.1 - 0.3 mg/kg subcutaneously or orally every 24 hours. (J29.15.w1)
    • The oral suspension is generally palatable to rabbits. (B601.16.w16, J15.30.w2)
Peri-operative and post-operative analgesia
  • Peri-operative analgesia is particularly important in rabbits as rabbits pain following surgery is a major cause of post-surgical anorexia. (J213.4.w5)
  • Provision of pre-operative analgesia may (a) provide more effective pain relief and (b) reduce the anaesthetic dose needed. (B601.16.w16, J15.20.w2)
    • Pre-operative buprenorphine can reduce the required concentration of isoflurane by 0.25 - 0.5% and the required concentration of sevoflurane by 1.0 - 1.5%. (B601.16.w16)
    • It is not possible to reduce the anaesthetic dose so easily if using injectable anaesthetics. (B601.16.w16)
  • For major surgical procedures such as orthopaedics, mandibular or maxillary surgery, give buprenorphine and consider also giving a NSAID (carprofen, ketoprofen or meloxicam). (B601.16.w16, P113.2005.w3)
    • As an adjunct, the surgical site may be infiltrated with a local anaesthetic e.g. bupivacain. Note measure the amount given carefully to avoid overdose - maximum 2 mg/kg bupivacaine, or 10 mg/kg Lidocaine (Lignocaine). (B601.16.w16)
  • For e.g. castration or ovariohysterectomy: potent NSAID at the time of surgery and a second dose orally 12-24 hours later. (B601.16.w16)
  • Note: provision of analgesia does not predispose rabbits to remove skin sutures. (B601.16.w16)
  • In general, pain relief should be given for 24 - 48 hours after surgery; a longer period may be required after extensive dental-related surgery. (B601.16.w16, P113.2005.w3)
Ferret Consideration Management of pain in ferrets is often inadequate due to concerns regarding side-effects of analgesic drugs such as opiates and NSAIDs. (J513.7.w3)
  • Ferrets require analgesia for example following traumatic injury as well as in the post-surgical period. (B602.2.w2, P120.2006.w6)
  • As with other species, multi-modal analgesia, acting at various levels of the nociceptive pathway, is preferable, allowing smaller doses of each drug, thereby reducing the risks of undesirable side effects. (J513.7.w3)
  • Provision of analgesia before the pain is perceived by the ferret is more effective. (B232.18.w18)
  • Assess pain before and after administering an analgesic: check that it is producing the desired analgesic effect. (B232.18.w18)
  • Note: ferrets are stoic and may not show pain; if in doubt, give analgesia.(J15.24.w5)
Signs of pain include
  • Anorexia (J29.6.w3) is seen with visceral pain; the ferret may show bruxism (teeth grinding) when offered food. (J29.14.w1)
    • Sometimes the ferret will eat only if hand-fed. (B631.22.w22)
  • Depression, immobility, lethargy, reluctance to move, reluctance to wake, loss of normal play behaviour. (B631.22.w22, J29.6.w3, J29.14.w1, J513.7.w3)
  • Curling up in a tight ball. (J29.14.w1)
    • OR inability to curl up into a proper sleeping posture. (J29.6.w3)
    • OR hunched posture. (B631.22.w22)
  • Silence (J513.7.w3), crying. (J29.6.w3) or screaming (B631.22.w22) - high pitched vocalisation; (J29.14.w1) may grunt when handled. (J29.14.w1)
  • Squinting. (J29.6.w3)
  • Lack of response to the environment, to other ferrets, to human attention and to being petted. (B631.22.w22, J513.7.w3)
    • OR aggressive response to disturbance (teeth-baring, biting by a normally gentle individual ). (B631.22.w22, J29.14.w1)
    • Sometimes wanting more cuddling; burrowing into the owner's clothing. (B631.22.w22)
    • May cringe and/or vocalise when touched. (B631.22.w22)
  • Hiding. (J513.7.w3)
  • Sometimes hyperventilation. (J513.7.w3)
  • Shivering or trembling despite normal body temperature. (B631.22.w22, J29.14.w1)
  • Tail fur bristling. (J29.14.w1)
  • Dull expression and dull or glassy eyes. (B631.22.w22)
  • Half-closure of the eyelids. (J29.14.w1)
  • Focal muscle fasciculations. (J29.14.w1)
  • Changes in gait (B631.22.w22): stiff movements; (J29.6.w3) lameness. (J29.14.w1)
  • Reduced or absent grooming. (B631.22.w22)
  • Bruxism, hypersalivation, pawing at the mouth. (B631.22.w22)

(J29.6.w3, J29.14.w1, J513.7.w3)

Note: 

  • Signs of acute pain are likely to be more obvious than signs of chronic pain. (J29.14.w1)
  • Effective management of chronic pain requires commitment from the owner; many owners are capable of developing very good recognition of pain in their ferret and dosing appropriately. (B631.22.w22) 

Appropriate analgesics include:

Opiates
  • Opiates can be used for up to three days in the treatment of acute pain, for example following surgery or traumatic injury. (B631.22.w22)
  • Both buprenorphine and butorphanol at the higher end of the dose rates will produce sedation (and depression); this can be useful to reduce movement of the ferret. (J29.6.w3)
  • Although opiates have a reputation for producing respiratory depression in ferrets, a contrary view suggests that ferrets become comfortable when post-surgical opiates are administered, therefore sleep normally. (J513.7.w3)
  • Buprenorphine  
    • 0.01-0.03 mg/kg intravenously, intramuscularly or subcutaneously, every 8 - 12 hours. (P120.2006.w6)
    • 0.01-0.05 mg/kg intravenously or subcutaneously every 8-12 hours. (J29.6.w3)
    • 0.01 - 0.03 mg/kg intravenously, subcutaneously or transmucosally (applied to the space between the molars and the buccal mucosa) every 6 - 10 hours. (J29.14.w1)
    • 0.05 mg/kg subcutaneously or intramuscularly; effective for up to 12 hours. (B232.18.w18)
    • 0.01 mg/kg subcutaneously, intramuscularly or intravenously every 8 - 12 hours. (B631.22.w22)
    • This can be used to reduce ketamine and medetomidine dosages in anaesthetic protocols. (B339.9.w9)
    • Reversal of the sedation and depression can be carried out using naloxone, 0.04 mg/kg  intravenously, intramuscularly or subcutaneously. (J29.6.w3)
    • This can be used concurrently with a NSAID. (B631.22.w22)
    • Minimal sedative effect. (B631.22.w22)
  • Butorphanol  
    • Butorphanol appears to be an effective visceral analgesic but a poor somatic analgesic in ferrets. (J513.7.w3)
    • Note: ferrets may be very lethargic after being given butorphanol, and remain immobile for long periods. (B602.2.w2)
      • Monitor careful if e.g. under a heat lamp, to avoid the risk of overheating. (B602.2.w2)
    • A ferret resting quietly, pain-free, after being given an opiate may be thought (incorrectly) to be showing respiratory depression. (J513.7.w3)
    • More sedative than buprenorphine. (B631.22.w22)
    • 0.1-0.5 mg/kg intramuscularly or subcutaneously, every 12 hours. (P120.2006.w6); intravenously, subcutaneously or intramuscularly every 4-6 hours. (J29.6.w3)
    • 0.1 - 0.4 mg/kg intravenously, intramuscularly or subcutaneously, every 2-4 hours. (J29.14.w1)
    • 0.2 - 0.4 mg/kg subcutaneously or intramuscularly every 6 - 8 hours. (B631.22.w22)
    • 0.25 mg/kg subcutaneously. (B232.18.w18)
    • Pre-operatively, 0.2 - 0.8 mg/kg subcutaneously, intramuscularly or intravenously. (J513.7.w3)
    • Post-operatively, initial 0.1 - 0.2 mg/kg loading dose followed by 0.1 - 0.2 mg/kg/hour by CRI. (J513.7.w3)
      • 0.1-0.2 mg/kg intravenously by CRI. (J29.14.w1)
  • Fentanyl 
    • Fentanyl is very effective against both visceral and somatic pain in ferrets. Due to rapid metabolism, constant rate infusion (CRI) is needed. (J513.7.w3)
    • 20 - 30 g/kg/hour intravenously by CRI during anaesthesia, to reduce inhalant anaesthesia concentrations. (B631.22.w22, J29.14.w1)
      • Monitor the ferret's blood pressure. (B631.22.w22)
    • Fentanyl can be used intraoperatively together with ketamine, in micro-doses via a constant rate infusor (CRI). Use of this combination reduces the concentration of anaesthetic inhalant (gaseous) anaesthetic agent required and reduces  hypotension associated with gaseous anaesthesia. Following a 0.1 mg/kg loading dose of ketamine, ketamine is given intra-operatively via CRI at 0.3 - 0.4 mg/kg/hour with fentanyl at 10-20 g/kg/hour, then post-operatively ketamine at 0.3 - 0.4 mg/kg/hour, with fentanyl at 5 - 10 g/kg/hour. (J513.7.w3)
  • Morphine 
    • 0.2 - 2.0 mg/kg intramuscularly as a single pre-operative dose. (J29.14.w1)
    • 0.5 - 5.0 mg/kg subcutaneously or intramuscularly every 2-6 hours. Usually just given as a single pre-operative injection. (B631.22.w22)
    • Morphine given at 0.22 mg/ kg provides up to 24 hours of post-operative analgesia. (J513.7.w3)
    • 0.1 mg/kg once, as an epidural for surgical analgesia; the effects last 12-24 hours. (B631.22.w22)
  • Hydromorphone is very effective against both visceral and somatic pain in ferrets. (J513.7.w3)
    • This can be used in management of pain due to trauma or another painful medical condition, as a pre-emptive analgesic prior to surgery, and in the management of moderate to severe post-surgical pain. (J29.14.w1)
    • 0.1 - 0.2 mg/kg intravenously, intramuscularly or subcutaneously, every 6 - 8 hours. (B631.22.w22, J29.14.w1)
    • Due to rapid metabolism, constant rate infusion (CRI) is needed. Hydromorphone can be used pre-operatively at 0.05 - 0.1 mg/kg intravenously. It can be used post-operatively at 0.05 mg/kg intravenously as a loading dose followed by CRI at 0.05 - 0.1 mg/kg/hour. (J513.7.w3)
  • Meperidine
    • 5 - 10 mg/kg subcutaneously, intramuscularly or intravenously every 2 - 4 hours. (B631.22.w22)
    • This may make some individuals drowsy. (B631.22.w22)
  • Nalbuphine
    • 0.5 - 1.5 mg/kg intramuscularly or intravenously ever 2-3 hours. (B631.22.w22)
  • Oxymorphone
    • 0.05 - 0.2 mg/kg subcutaneoulsy, intramuscularly or intravenously every 8 - 12 hours. (B631.22.w22)
  • Pentazocine
    • 5 - 10 mg/kg intramuscularly every four hours. (B631.22.w22)
  • Tramadol
    • 5 mg/kg orally every 12-24 hours. (B631.22.w22)
    • Useful for mild to severe pain; can be used together with an NSAID and can be used long term in management of neoplasia-associated pain. (B631.22.w22)
NSAIDs
  • Ferrets, like cats, are deficient in enzymes of the glucuronidation pathway, therefore NSAIDs must be used carefully to avoid overdosing. (J513.7.w3)
  • These drugs are useful in alleviation of traumatic pain and postoperative pain; COX-2 inhibitors are preferred. (J513.7.w3)
  • NSAIDs are useful in treatment of acute and chronic pain. (J29.14.w1)
  • NSAIDs can be used for up to seven days in the treatment of post-surgical pain or traumatic pain. (B631.22.w22)
  • Carprofen  
    • 1 mg/kg orally every 12-24 hours. (B631.22.w22, P120.2006.w6)
      • Use with an H2-blocker. (B631.22.w22)
    • 1 - 5 mg/kg orally. (B339.9.w9)
    • 4 mg/kg orally every 24 hours. (J513.7.w3)
    • This has been found safe for use in ferrets. (J513.7.w3)
  • Meloxicam  
    • This is the NSAID used most commonly in ferrets. (J513.7.w3)
    • The palatable liquid is generally acceptable to ferrets. (J29.14.w1)
    • Meloxicam is safe for short term use. (J29.14.w1)
    • Caution is advised regarding longer term use; liver and kidney function should be monitored via blood biochemistry. (J29.14.w1)
    • 0.1 mg/kg orally every 24 hours. (P120.2006.w6)
    • 0.1 - 0.2 mg/kg subcutaneously or orally every 24 hours. (J29.14.w1)
    • 0.2 mg/kg orally. (B339.9.w9)
    • 0.2 mg/kg subcutaneously, intravenously or orally as an initial dose, then 0.1 mg/kg every 24 hours. (J513.7.w3)
    • 0.2 mg/kg orally, subcutaneously or intramuscularly every 24 hours. Use with an H-2 blocker. This can be used long-term, but liver parameters should be monitored. (B631.22.w22)
  • Ketoprofen.
    • Can be used postoperatively; due its COX-1 effects, should not be used pre-operatively. (J513.7.w3)
    • 1 - 2 mg/kg every 24 hours post-operatively. (J513.7.w3)
    • 0.5 - 1.0 mg/kg orally or subcutaneously every 24 hours. This should be used with an H2 blocker, and cautiously, for less than five days. (B631.22.w22)
    • 2 mg/kg subcutaneously every 24 hours. (B232.18.w18)
  • Aspirin
    • 10-20 mg/kg orally every 24-48 hours. (J29.6.w3)
    • 10-20 mg/kg orally every 24 hours. Give an H2-blocker concurrently. (B631.22.w22)
  • Flunixin meglumine (NSAID)
    • 1.1 mg/kg subcutaneously or orally every 12-24 hours. (J29.6.w3)
    • 0.5 - 2.0 mg/kg subcutaneously. (B232.18.w18)
    • 0.3 mg/kg orally or subcutaneously every 24 hours. Use with an H2 blocker, and for less than three days. (B631.22.w22)
    • The newer COX-2 inhibitors (e.g. carprofen, meloxicam) may be safer. (J513.7.w3)
    • Potential renal damage; not recommended. (B631.22.w22)
  • WARNINGS:  
    • Contraindicated in pregnant ferrets. (J29.14.w1)
    • Contraindicated in ferrets with renal disease, hypovolaemia or bleeding disorders. (J513.7.w3)
    • Contraindicated in ferrets with hepatic dysfunction. (J29.14.w1)
    • Contraindicated in ferrets with known gastrointestinal ulceration. (J29.6.w3, J29.14.w1)
      • Avoid using concurrently with corticosteroids.  (J29.6.w3)
    • Contraindicated with limited organ perfusion, for example with shock. (J29.14.w1)
    • Avoid using if severe haemorrhage is likely during surgery. (J513.7.w3)
    • To reduce the risk of gastric ulceration, 
    • As in cats, do not give paracetamol (acetaminophen) to ferrets because hepatic glucuronidation of this drug is very slow in these species (low UDP-glucuronosyltransferase activity), leading to toxicity. (P120.2006.w6)
    • Ibuprofen Toxicity in Ferrets has been reported on a number of occasions and a single human tablet can provide a fatal dose. (J4.216.w1, J513.3.w4)
  • Use of H2 blockers is recommended when using NSAIDs:
    • Cimetidine, 10 mg/kg orally, subcutaneously, intramuscularly or by slow intravenous injection, every eight hours. The oral form is unpalatable to ferrets. (B631.22.w22)
    • Famotidine, 2.5 per ferret, orally, subcutaneously or intravenously every 24 hours. Available as a non-prescription tables, palatable. (B631.22.w22)
    • Ranitidine 3.5 mg/kg orally every 12 hours. the oral form is unpalatable to ferrets. (B631.22.w22)
Peri-operative and post-operative analgesia

Pre-emptive analgesia is recommended. Pain management is generally more successful when pre-emptive analgesia is used rather than analgesia in response to pain. Also, pre-surgical analgesia usually reduces general anaesthetic requirements. (J29.14.w1)

  • Ketamine can be used as part of the anaesthetic induction protocol to reduce "wind-up". (J513.7.w3)
    • 0.5 mg/kg intravenously pre-surgery. (J29.14.w1)
    • 4 - 10 mg/kg intravenously for induction. (J513.7.w3)
    • 10 g/kg/minute intravenously by CRI during surgery. (J513.7.w3)
    • 2 g/kg/minute intravenously by CRI postoeratively, in combination with an opioid, for 24 hours after surgery. (J513.7.w3)
    • Micro-dosing with Ketamine plus Fentanyl has been used, via a constant rate infuser (CRI), during and after surgery. Use of this combination reduces the concentration of anaesthetic inhalant (gaseous) anaesthetic agent required and reduces  hypotension associated with gaseous anaesthesia. Following a 0.1 mg/kg loading dose of ketamine, ketamine is given intra-operatively via CRI at 0.3 - 0.4 mg/kg/hour with fentanyl at 10-20 g/kg/hour, then post-operatively ketamine at 0.3 - 0.4 mg/kg/hour (with fentanyl at 5 - 10 g/kg/hour). (J513.7.w3)
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.

In primates

  • Post-operative self-mutilation may indicate pain. (B671.13D.w13d)
  • Appropriate analgesic drugs should be administered at appropriate doses and dose intervals to ensure analgesia. (B671.13D.w13d)
  • A combination of a NSAID and an opiate is often appropriate for the relief of post-surgical pain. (B671.13D.w13d)
  • Information on drug dosages is provided in the pages linked from Drugs used in the treatment of Bonobos - Painkillers (Analgesics), Anti-inflammatories and Symptomatics
Associated techniques linked from Wildpro

Injection Techniques

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Anaesthesia and Chemical Restraint

NOTE: Before using any anaesthetic agent or combination of agents, the manufacturer's data sheet on the agent or agents concerned should be consulted, taking particular note of any contra-indications and operator warnings.
  • N.B. Whenever an anaesthetic is undertaken, the anaesthetist must be familiar with emergency protocols. Consideration must be given as to the availability of equipment required to monitor the anaesthetic plane of the animal being anaesthetized and any equipment/drugs required for revival. It is advisable to calculate the doses of any revival agents which may be required in an emergency BEFORE COMMENCING the anaesthetic. An accurate body weight should be determined to allow accurate dosage. (J34.23.w1, P3.1999b.w2, V.w6).
  • Always consider whether the risks of the anaesthetic are outweighed by the benefits gained by the immobilization. (P106.2007.w5)
  • The ideal anaesthetic produces a smooth, reliable induction, provides relief of the patient from fear and anxiety, produces appropriate levels of restraint, analgesia and relaxation for the procedure to be performed, for the length of time required for the procedure, and a rapid, uneventful and full recovery.
  • The required degree of restraint and relaxation may vary from minimal, e.g. for radiography, to considerable for orthopaedic or abdominal surgery. Both intra-operative and post-operative analgesia are usually required for surgery, and anaesthetic agents which provide good analgesia often allow better muscle relaxation and restraint; they may also allow maintenance on a lower (and safer) anaesthetic plane.
  • In general, premedications which produce a longer recovery should be avoided unless specifically indicated. (J34.23.w1)

Pre-anaesthetic assessment and care

  • Assess the patient's general physiological status before anaesthetic induction, if possible. 
  • When dealing with wild animals for which there is no history available, it is particularly important to ensure that a proper assessment has been carried out and the patients stabilised before anaesthesia, if possible. (B545.8.w8, V.w5)
    • Note: in many circumstances when dealing with wild animals (e.g. wildlife casualties, animals in zoos) it is necessary to anaesthetise the animal without performing a pre-anaesthetic physical examination or any other tests to determine health status.
  • Basic cardiac and respiratory function may be assessed by observation of the awake animal. If the animal is in a quiet, thermoneutral environment and is undisturbed, but is showing rapid, shallow respiration, then further evaluation is needed.
  • Dehydration can be assessed by rolling the skin between the fingers - this is more difficult as dehydration increases. (J34.23.w1)
  • If possible, the animal should be physiologically stabilised before anaesthesia. When this is not possible, stabilisation should commence as soon as the animal is anaesthetised.
  • If the patient has cardiopulmonary compromise, they should be oxygenated (e.g. in an oxygen cage) before induction.

Anaesthetic monitoring and support

  • During anaesthesia, the animal should be placed in a position which promotes respiration and minimises the risk of aspiration of saliva or regurgitated material.
  • The limbs should be positioned to minimise the development of circulatory impairment.
  • The eyes should be covered to protect them and t minimise visual stimuli. 
  • Cotton wool may be placed in the ears to minimise stimulation due to noise; it is important to ensure this is removed prior to anaesthetic recovery.
  • Monitoring of cardiovascular function, respiratory function and temperature is extremely important.

(B11.9.w20, B13.49.w16, B14, B121, B486.11.w11, J1.5.w5, J34.23.w1, P3.1999b.w2, P106.2007.w5, V.w5, V.w6)

Further information is available in this section (see below) on:

Waterfowl Consideration

(Information on ANAESTHETIC EMERGENCIES is at the end of these Waterfowl Considerations.)
HANDLING AND RESTRAINT
  • See also Manual Restraint information in the Bird Handling & Movement - Holding & Carrying.
  • Waterfowl show a very variable response to stimuli; feather follicles are sensitive and there may be violent reaction to feather plucking, but little reaction to suturing or cutting the skin, or even handling viscera. The bill, head and feet are also sensitive. (B10.26.w3, B11.9.w20, B13.46.w1, B14).
PRE-ANAESTHETIC PREPARATION
  • Pre-anaesthetic handling should be minimal, as gentle as possible and as stress-free as possible. (B11.9.w20)
  • A bird should be in as good a state of health as possible before being anaesthetized (B11.9.w20). A blood sample should be taken if there is any doubt as to the health status of the bird, if time allows. Minimum clinical profile of AST, bile acids, LDH, urea, uric acid, full haematology and clotting time is suggested (B14).
  • Hydration status should be considered: fluid therapy should be given before anaesthetic if the PCV is above 55%. If isoflurane anaesthesia is used, fluid therapy may be started immediately after induction, rather than prior to induction. (B11.9.w20, B14)
  • Possible hypoglycaemia: intravenous 5% glucose should be given before, during and following surgery if the blood glucose level is below 16mmol per litre. (B11.9.w20)
  • Liver and kidney functions should be considered: halothane is contra-indicated with liver dysfunction (and in debilitated birds); ketamine is contraindicated with kidney dysfunction.
  • Starvation for two to six hour prior to surgery has been suggested to reduce the risk of oesophageal reflux and inhalation prior to surgery (B37.x.w1).
  • Weight should be determined accurately prior to anaesthetic for dose calculation with injectable agents. If inhalation induction is to be used, weight may be determined after induction, but is still required e.g. for fluid therapy calculations (B11.9.w20).
  • The number of contour feathers removed to provide a clean surgical site should be minimized to retain waterproofing and allow an early return to water.
  • Birds must be kept warm before, during and after anaesthetic. Cooling may lead to hypothermia, which may be fatal, and may predispose to cardiac arrhythmias, as well as increasing recovery time. Vetbed, towels or similar materials, or a heating pad, should be placed under the bird. The use of wetting agents such as surgical spirit should be minimized due to their chilling effect. Consideration should be given to the use of disposable adhesive drapes such as Opsite (Smith & Nephew) to minimize the area which must be prepared for surgery while maintaining an adequate clear surgical site. N.B. cool anaesthetic gases flowing through the respiratory tract also act to cool the bird.(B11.9.w20, B14)
  • Profuse salivation is common in anaesthetized waterfowl. The use of an anticholinergic agent such as atropine or glycopyrrolate is not recommended, as this results in thicker secretions.

(B11.9.w20, B13.39.w16, B14, B37.x.w1)

ANAESTHETIC MONITORING

Constant monitoring of depth of anaesthesia, heart, respiration and if possible oxygenation is important; temperature should also be monitored.

  • Cardiac: doppler probe under tongue, against carotid artery or on recurrent ulnar artery, or oesophageal stethoscope, or ECG (leads placed over distal lateral tarsometatarsus and carpal joint of each wing, with atraumatic clamps or silver needles); pulse - e.g. radial artery. Normal heart rate in waterfowl is quite variable, e.g. 180-230bpm in the Pekin duck (Anas platyrhynchos domesticus - Domestic duck). Changes in heart rate may give more information than absolute rate (B11.9.w20, B13.46.w1, B14).
  • Oxygenation: Oximeter may be used over tibiotarsal bone (appears most consistent and reliable), wing web, toe or tongue. Arterial oxygen saturation should be maintained well over 85%: level under 80% may be dangerous. Pulse oximeter will also give pulse rate (B13.39.w16, B14).
  • Temperature: rectal or oesophageal thermometer. Normal body temperature of waterfowl is approximately 39-41.6C. N.B. Longer recovery time with lower body temperature, as well as myocardial depression. (B11.9.w20, B13.46.w1, B14).
  • Tube patency should be checked regularly - may become blocked by secretions. Changing the tube every 20 minutes during anaesthesia has been suggested (B37.x.w1).
  • Respiration - rate and excursion should be noted; respiration should be slow and regular. Normal respiratory rate is quite variable in waterfowl - may be e.g. 13-40 breaths per minute in geese and swans, or e.g. 30-95 breaths per minute in ducks (B11.33.w1, B13.46.w1). Depth, rate and pattern of respiration should be noted, particularly any changes. Rapid, jerky respiration or hyperventilation indicate ensuing problems. Slow irregular respiration with too deep anaesthesia. Increased respiratory rate/depth may indicate lightening of anaesthetic plane, stimulation (pain), difficulty in breathing (e.g. due to a blocked tube) or elevated paCO2. Rapid, shallow or intermittent respiration may also indicate too deep anaesthesia. N.B. Drapes may make visual monitoring difficult - clear surgical drapes facilitate monitoring, as does the use of a small anaesthetic bag. Apnoea monitors may be used (e.g. Imp respiratory monitor, IMP Electronics; apALERT apnoea monitor, MBM Enterprises, Australia) but may not register respiration, especially in small birds. They can only be used if the bird is intubated, and care must be taken that the monitor does not kink the tube. Respiratory rate should not fall below 12-15 breaths per minute for large birds such as swans, or below 25-50 breaths per minute for birds weighing less than 500g (should be not less than half of normal resting rate), or hypercapnoea may develop (B11.9.w20, B13.39.w16, B14, B37.x.w1).
  • Reflexes: loss of voluntary motion, but retained palpebral, corneal and pedal reflexes in light anaesthetic plane (B13.49.w16), slow to absent pedal reflex and wing reflex in surgical plane of anaesthesia, loss of corneal reflex indicates deep anaesthesia (B13.49.w16). Wing flutter may indicate the bird is becoming light (B13.39.w16).
  • Capnography - measurement of end tidal carbon dioxide level may be useful, but is not standard at present (B14).
ORAL SEDATION
  • In certain circumstances sedation with an orally absorbed drug may be an appropriate means of waterfowl capture. This method may be used to capture an individual bird (e.g. one duck in a park situation), using a bait which can be targeted at that individual, such as a piece of bread, or a group of waterfowl, for example by using baited grain.
  • In using oral bait to sedate/anaesthetize waterfowl for capture it is particularly important to ensure that the bird(s) are watched closely with rapid intervention to prevent drowning or attack by other individuals. This method must be used with extreme caution if the possibility exists that the birds may fly away from the site between ingestion of the drug and it having its effect. Other potential hazards include a lack of control over the amount of drug consumed by each individual, variability in the responses of different individuals to a given dose (possible effects of age, sex, health status and degree of stress), and effects on non-target species consuming the bait. Additionally, there is little data on the effects of orally administered immobilizing agents on behaviour, physiology and survival. The possibility of residues must also be considered if birds may be used for human consumption.
  • Drugs which have been used or tested for use as oral immobilization agents include alpha-chloralose, methoxymol, metomidate, pentobarbital sodium, secobarbital sodium, thiopental sodium and tribromoethanol. A combination of alpha-chlorulose and tribromoethanol has also been used successfully.
  • (See: Oral Sedation of Waterfowl).

(J2.8.w1, J4.161.w1, B13.46.w1, B36.4.w4, B123).

LOCAL ANAESTHESIA
  • Frequently sufficient for superficial procedures.
  • Safe if dose is carefully calculated (B14).
  • Gross overdose may occur in small birds if dose not calculated.
  • Lidocaine (Lignocaine) hydrochloride (2%) usually safe and effective, although general depression may occur with high doses. 1-3 mL may be used in birds greater than 2kg body weight, and up to 1 mL in a 400 g bird. Solution may be diluted to give 0.5% solution. Preparations with adrenaline are recommended to limit absorption rate. Lignocaine ointment may be used around the vent following cloacal prolapse (B14). Maximum dose 4 mg/kg (B23.39.w3).
  • Use of 2% procaine, 1ml in ducks, 3ml in swans reported to provide good local anaesthesia with few problems. (B13.46.w1). Narrow safety margin, recommended dilution to produce 0.2% solution, after which 1-2 mL/kg may be used (B14).
  • Xylocaine hydrochloride has also been considered a safe local anaesthetic for use in waterfowl (B10.26.w3).
INJECTABLE ANAESTHESIA
  • Often marked inter-species and intra-species variability in response. (J13.51.w1, B13.39.w16).
  • Accurate weighing important for correct dose calculation (B11.9.w20, B14).
  • Anaesthetic dose may be difficult to control, and irreversible after administration. (J13.51.w1, B13.39.w16).
  • Predispose to intraoperative hypothermia and hypoglycaemia: further exacerbated by prolonged recovery time (J13.51.w1, B13.39.w16).
  • Injectable anaesthetic drugs and drug combinations which have been used in waterfowl include propofol (Rapinovet), alphaxolone/alphadolone (Saffan), ketamine, ketamine/xylazine, ketamine/medetomidine, ketamine/diazepam.
  • Propofol (8mg/kg) may be induction agent of choice if mask induction with isoflurane is not possible (B37.x.w1), and it has been suggested that it may be preferable to isoflurane in some field situations (J1.36.w1), although it also has been suggested that the duration of action of propofol is too short to be of practical use in birds (B11.9.w20).
  • Details of the use of individual injectable anaesthetic agents are given in:
  • N.B. Injectable anaesthetic agents may provide a health hazard for the anaesthetist and other people exposed to these anaesthetic agents. Care should be taken to avoid self-injection. Several drugs may be absorbed through the skin and/or mucous membranes and care should be taken to avoid splashing of drugs onto the skin, lips or eyes. If such contact does occur the relevant area should be irrigated with copious amounts of water (B121).
GASEOUS ANAESTHESIA
  • Precisely controllable, little species variation, little or no metabolism required.
  • Should always be used with precision vapourizer.
  • N.B. greater efficiency of avian lungs produces greater sensitivity to small changes in anaesthetic gas concentration (B11.9.w20).
  • Carrier gas flow 3-5 litres/minute.
  • Relatively high level for induction (e.g. isoflurane at 4.0 to 5.0%), allows rapid induction (may be within four to five breaths) (B11.9.w20).
  • Lower level for maintenance.
  • Intermittent positive pressure ventilation may be required to maintain adequate oxygenation (B11.9.w20, B13.39.w16)
  • Isoflurane is generally considered to be the anaesthetic agent of choice for birds for both induction and maintenance. N.B. it has been suggested that propofol may be more suitable for some field situations (J1.36.w1).
  • Always leave on oxygen for a time after turning off anaesthetic agent at the end of the procedure (B14).
  • Details of the use of individual gaseous anaesthetic agents are given in:
  • N.B. Exposure to gaseous anaesthetic agents may have health implications for the anaesthetist and other people exposed to the anaesthetic agents. It is suggested that "reasonable measures should be taken both to reduce the risk of serious contamination of the atmosphere with inhalation anaesthetics and to remind operating theatre staff of possible hazards" (B121).
  • The "reasonable measures" include filling vapourizers using proper filling apparatus or funnels, outside the operating theatre and preferably out-of-doors, turning vapourizers off when not in use, taking care when handling anaesthetic agents, using low flow systems when possible, using scavenging of waste gases/vapours, using endotracheal intubation rather than a face mask when possible and checking breathing circuits regularly for leaks (B121).

(B11.9.w20, B13.39.w16, B13.46.w1, B14, B37.x.w1).

ANAESTHETIC EQUIPMENT AND USE

Circuits:

  • Non-rebreathing circuit with small dead-space and low resistance recommended, e.g. modified Ayres T-piece or mini-Bain system.
  • Bain co-axial circuit may warm inflow gases and provide humidification using expired gases of patient.
  • IPPV (intermittent Positive Pressure Ventilation) may be provided with reservoir bag. Important to maintain high oxygen flow rate during bird anaesthesia.
  • Flow rate should be at least three times normal minute volume for the individual: minute volume of 2.5 kg chicken is 770 mL/minute, suggested flow rate 3 litres per minute for birds of this general size. Minute volume of American black duck (Anas rubripes - American black duck) at rest 815.4 mL/minute (body weight 1.026 kg).
  • Scavenging system should be used to protect operating room personnel from exposure to anaesthetic gases.

(B11.9.w20, B14, B13.39.w16).

Ambu bag:

  • Self-inflating resuscitation apparatus
  • May be used for ventilation during anaesthesia in the field.
  • Paediatric size may be suitable e.g. for ducks; neonatal size may be required for small birds.
  • Squeeze to give visible expansion of the thorax, twenty times a minute (every five seconds).

(J1.36.w1, P3.1999.w2)

Face mask:

  • Used for induction and may be used for maintenance in very short procedures (up to 10-15 minutes) (B11.9.w20, B13.39.w, B13.46.w1). Choose to fit comfortably with minimal dead space. Commercially-available transparent masks or a mask created e.g. from a 60ml syringe case may be modified using a latex glove or tape to reduce leakage.

Intubation: (Used once the bird has been induced, although may not be required for very short procedures.)

  • Reduces dead space, maintains airway.
  • Glottis is visible just behind the tongue.
  • Intubation is easier if the tongue is pulled forwards.
  • Usually soft uncuffed tube, but cuffed tube, inflated with care, may be chosen for waterfowl, to avoid inhalation of regurgitated fluids, particularly for e.g. flushing oesophagus and gizzard.
  • Care not to over-inflate due to risk of pressure necrosis (birds have complete tracheal rings and fragile tracheal mucosa).
  • Care to ensure neck extended in long-necked birds to avoid risk of trachea folding over the edge of the tube with resultant partial or complete obstruction of the airway.
  • Risk of blockage of tube with mucus (copious secretions by waterfowl during anaesthesia). Risk of blockage is greatest in small birds, due to small diameter tube. Tube should be checked regularly throughout anaesthesia. Changing the tube every 20 minutes has been suggested (B37.x.w1).
  • Allows scavenging of waste gases.
  • Facilitates artificial ventilation.

(B11.9.w20, B13.46.w1, B37.x.w1, P3.1999b.w2, P7.1.w4)

AIR SAC INTUBATION
  • Useful for surgery of the mouth/bill, or in a bird with tracheal obstruction.
  • Usually placed in an anaesthetized bird. May be placed in an unanaesthetised bird with tracheal obstruction.
  • Placement similar to normal site for endoscopic examination e.g. for surgical sexing.
  • Surgically prepare site. (Minimal skin preparation with alcohol may be used in an emergency).
  • Extend leg caudally.
  • Stab incision of skin over sternal notch, or snick incision with scissors, with points then spread to enlarge hole.
  • Thrust straight artery forceps (haemostats) through muscle into abdominal cavity, then opened slightly to allow insertion of cannula, or trochar may be used, to gain access to air sacs.
  • Place large tube of inert material (e.g. French 14G silastic tubing, or commercially available air sac cannula - Cook Instrumentation) into airsac to maximum depth of 1cm (to minimize risk of liver or spleen damage by the tube).1-2.5 inches 2.5-7cm (P3.1999.w2)
  • Suture in place using purse-string suture transfixing the tube (monofilament non-adsorbing suture material).
  • Attach end of tube to anaesthetic circuit.
  • Intermittent positive pressure ventilation recommended while anaesthetized: may stop breathing due to hypocapnoea, and not restart spontaneous breathing until airsac perfusion is stopped and blood paCO2 rises.
  • Tube may be removed postoperatively, or may be left in place e.g. if syringeal aspergillus plug or other dyspnoic problem.

(B11.9.w15, B13.39.w16, B14, P3.1999b.w2)

EFFECT OF POSITIONING
  • In dorsal recumbency, breathing amplitude may decrease by 40-50% or even more; IPPV may be useful, 20-40 per minute, with peak pressure 15-20cm water (B11.9.w20, B13.39.w16, B14).
  • In lateral recumbency, a rolled towel may be placed between the keel of the bird and the table to allow ventral movement; reduction of breathing amplitude much less marked in lateral recumbency than in dorsal recumbency (B14, P8.3.w1).
RECOVERY
  • Variable length: may be less than five minutes following isoflurane anaesthesia, but much longer with e.g. ketamine.
  • Endotracheal tube may be left in place until the bird has recovered voluntary head and neck control (B37.x.w1, P3.1999.w1, J1.36.w1).).
  • IPPV may be continued until the endotracheal tube is no longer tolerated, in severely depressed birds (B13.39.w16).
  • Should take place in a warm, dim and quiet environment.
  • Should be monitored, with the bird not left unsupervised before it is balanced and able to stand/perch unaided (B13.39.w16).
  • Wrapping the bird e.g. in a towel may be useful to prevent wing-flapping and self-induced trauma, particularly following e.g. ketamine in which the recovery period may be prolonged.
  • Give access to water only after full recovery (B10.26.w3).

(B10.26.w3, B11.9.w20, B13.39.w16, B37.x.w1, P3.1999b.w2)

POST-OPERATIVE ANALGESIA

The following combinations have been suggested for use in birds:-

  • Buprenorphine (0.01-0.05 mg/kg intramuscular) (B11.9.w20).
  • Butorphanol 1.0-4.0 mg/kg intramuscular, N.B. short duration of action (2-4 hours ) (B23.39.w3).
  • Ketoprofen (Ketofen, Rhine Merieux): 1 mg/kg intramuscular once daily for up to ten days (B11.X.w11); 5-10 mg/kg intramuscular (B11.9.w20); 1-5mg/kg intramuscular twice daily (B11.4.w17).
  • Carprofen (2 mg/kg intramuscular) (B14); 4 mg/kg subcutaneous once daily, has been used on three consecutive days (B37.x.w1); 5-10 mg/kg intramuscular (B11.9.w20).
  • Flunixin meglumine 1mg/kg subcutaneous daily, has been used on three consecutive days (B37.x.w1). 1.0-10.0 mg/kg once daily (B23.39.w3); 1-10 mg/km intramuscular (B11.4.w17)
  • Aspirin 30 mg/200 g body weight has been used (B11.4.w17).
ANAESTHETIC EMERGENCIES:
SPECIFIC IMMEDIATE RESPONSE TO RESPIRATORY ARREST
  1. Disconnect from anaesthetic gas.
  2. Increase oxygen flow rate.
  3. Press on sternum gently, 40 to 50 cycles per minute (B13.36.w16); unlikely to be effective and may cause physical damage (B14).
  4. Intubate or insert air sac tube if neither is present (see above for technique).
  5. Provide intermittent positive pressure ventilation (IPPV) through the tube, by intermittent occlusion of exhaust arm of Ayres T-piece or by using rebreathing bag, or appropriate-sized AMBU bag (neonatal or paediatric). 12-15 breaths per minute
  6. Administer reversal agent if injectable anaesthetic has been used: 
    • Naloxone for opioids, total dose 2mg, slow intravenous injection P3.1999b.w2,
    • Atipamezole for alpha-2 agonists, intravenous or intramuscular N.B. also antagonizes the analgesia provided by the alpha2 agonist P3.1999b.w2
  7. Doxapram, 5-10mg/mg (P3.1999b.w2) 7mg/kg (0.3ml/kg): dilute 1:3 and give by slow intravenous injection or intramuscularly, or in smaller birds dropped onto the tongue may help stimulate respiration.
  8. Adrenaline may be given, 0.5-1.0mg/kg intravenous, in response to cardiac arrest, anaphylactic shock or bronchial spasm (P3.1999b.w2).
  9. Dexamethasone (synthetic corticosteroid) 4mg/kg (1mg/kg in raptors) intramuscular or subcutaneous in case of shock. (P3.1999b.w2).
  10. Dextrose: administer in the case of seizures due to hypoglycaemia.

(B13.39.w16, B14, P3.1999b.w2).

EMERGENCY DRUGS
  • Administer reversal agent if injectable anaesthetic has been used:
  • Naloxone (pure opioid antagonist) for opioids, total dose 2 mg, slow intravenous injection (P3.1999b.w2).  Reverses respiratory depression caused by opioids.
  • Atipamezole for alpha-2 agonists, intravenous or intramuscular N.B. also antagonizes the analgesia provided by the alpha2 agonist. (P3.1999b.w2)
  • Doxapram: short-acting respiratory stimulant. 5-10 mg/mg (P3.1999b.w2) 7 mg/kg (0.3 mL/kg): dilute 1:3 and give by slow intravenous injection or intramuscularly, or in smaller birds dropped onto the tongue may help stimulate respiration.
  • Adrenaline may be given, 0.5-1.0 mg/kg intravenous, in response to cardiac arrest, anaphylactic shock or bronchial spasm (P3.1999b.w2).
  • Dexamethasone (synthetic corticosteroid) 4 mg/kg (1 mg/kg in raptors) intramuscular or subcutaneous in case of shock. (P3.1999b.w2).
  • Dextrose: administer in the case of seizures due to hypoglycaemia.
  • Diazepam: first-line treatment for seizures, including epileptic fits, and all other seizures except those caused by Strychnine and hypoglycaemia
  • Atropine: Anticholinergic. Initial bradycardia due to central effects, followed by tachycardia due to blockage of cardiac muscarinic receptors. Also relaxation of bronchial smooth muscle. 0.5 mg/kg intramuscular
  • Prednisolone sodium succinate (Solu-Medrone): 2-4 mg/kg intramuscular. Synthetic water-soluble corticosteroid. Treatment of shock, endotoxaemia, spinal cord compression.
  • Sodium bicarbonate 1-4mg/kg slow intravenous injection

(P3.1999b.w2).

CHECK EQUIPMENT FOR FAILURE
  • Endotracheal tube:
  • Check correct placement in trachea, not oesophagus
  • Not too long (causing bronchial intubation)
  • Too narrow
  • Obstructed
  • Kinked
  • Disconnected from anaesthetic machine
  • Vaporizer:
  • No anaesthetic agent
  • Incorrect setting on dial
  • Wrong agent in vaporizer
  • Inaccurate vaporizer calibration
  • Anaesthetic machine:
  • No gas in cylinders
  • Flow meter - incorrect siting
  • Flow meter - failed
  • Connections (vaporizer-machine, or breathing system) - leakage
  • Breathing system obstruction.

(P3.1999b.w2)

Crane Consideration

Induction inhalational anaesthesia, crane. Click here for full-page view with caption. Maintenance inhalational anaesthesia, crane.  Click here for full-page view with caption.

Gaseous anaesthesia
  • Gaseous anaesthesia with Isoflurane is generally used for both induction and maintenance. (B115.8.w4, B336.20.w20)
    • This provides rapid, smooth induction and recovery, with lower complication rates than injectable agents. (B115.8.w4, B336.20.w20)
  • A mask is used for induction. (B336.20.w20)
    • A 60 mL syringe case can be adapted to use as a face mask, with the closed end cut open and attaching to the tube from the anaesthetic machine, while the open end (lightly padded) fits over the crane's bill. (B12.56.w14, B336.20.w20, B703.10.w10)
  • A 2.5 - 6.0 uncuffed tracheal tube is used during maintenance of anaesthesia. (B115.8.w4, B336.20.w20)
  • During endoscopic retrieval of foreign bodies from the gizzard, use of a cuffed tube with about 1 - 2 mL air used for inflation has been preferred, due to the risk of leakage of gastric contents into the trachea during surgery. There has been no problem with injury to the trachea with this low volume inflation. (P87.8.w6)
  • Use 4- 5% isoflurane  in oxygen for induction. (B12.56.w14, B703.10.w10)
  • For maintenance of anaesthesia, 1-3% Isoflurane in oxygen (oxygen flow rate 1-2 litres per minute). (B12.56.w14, B703.10.w10)
  • Halothane has been used in the past. (B115.8.w4, J14.17.w2)
    • Halothane is associated with a higher rate of cardiac and respiratory problems. (B115.8.w4)
Injectable agents
General notes
  • Injectable agents are now rarely used for anaesthesia in cranes; induction and maintenance with isoflurane is generally preferred. (B115.8.w4, B336.20.w20, B703.10.w10)
  • Complication rates (both cardiac and respiratory) B115.8.w4, are generally higher with injectable anaesthetic agents than with isoflurane inhalant anaesthesia. (B336.20.w20)
  • Once injectable agents have been given, the crane must be held until it is anaesthetised, as if left alone it might injure itself when it loses its balance. (B703.10.w10)
  • The crane should be intubated even if injectable anaesthesia is being used rather than a gaseous anaesthetic agent, in order to minimise the risk of regurgitation and aspiration of gastrointestinal contents. (B703.10.w10)
  • During recovery, the crane must be held until it can stand and walk without falling, or it might injure itself. (B703.10.w10)
Injectable agents used for anaesthesia
  • Ketamine 10-15 mg/kg plus 1.0 mg/kg Xylazine, or 10-15 mg/kg Ketamine plus 0.2 - 0.5 mg/kg Diazepam, intramuscularly, can be used as a preanaesthetic, for sedation before surgery, or for a short time of manipulation. (B12.56.w14)
  • Ketamine 22 mg/kg plus Xylazine 1 mg/kg can be used for surgical anaesthesia if gaseous anaesthetic agents are not available. (B12.56.w14)
    • Yohimbine has been used to reduce reversal time following use of xylazine. (B115.8.w4)
  • Ketamine plus midazolam has been used. (B336.20.w20)
  • Propofol can be used if use of a gaseous anaesthetic agent is not practical. (B703.10.w10)
  • Alfaxalone-alphadolone can be used at 6.5-7.0 mg/kg intravenously for induction. This gives a smooth induction after about 13-26 seconds, with anaesthesia lasting 5-6 minutes. Time to standing without ataxia is about 20-33 minutes. (B703.10.w10)
    • In a study, 6.5 - 7.0 mg/kg alfaxalone-alphadolone intravenously into the metatarsal vein provided smooth, rapid induction (13-26 seconds, with good muscle relaxation and maximum effect sufficient for e.g. endoscopic sexing lasting for five to six minutes. Recovery took 12-15 minutes to head-up in four Grus grus - Common crane and 25 Grus virgo - Demoiselle crane, but 21 minutes in two Balearica regulorum - Grey crowned-crane. The crowned cranes also took longer until they were on their feet (28 minutes) versus 18-21 minutes for the other two species and to standing without ataxia: 33 minutes compared with 20-25 minutes. (J3.145.w11)
    • In a Grus antigone - Sarus crane, 2.8 mg/kg alfaxalone-alphadolone (C1341) intravenously was considered to produce good ultra-short anaesthesia, with induction sufficient to allow intubation after 20 seconds, and seven to 11 minutes of anaesthesia suitable for radiography, manipulations and removal of a poorly placed intramedullary pin in a fractured humerus. Anaesthesia was also prolonged with halothane in oxygen on one occasion. (J14.17.w2)
Sedation/tranquilisation
  • Diazepam, 0.5 - 1.0 mg/kg alone provides about 4-6 hours of tranquilisation. The dose given e.g. for shipping an agitated crane should be enough to quieten the crane, but low enough that it can still balance. (B703.10.w10)
Oral sedation
Bear Consideration

Click here for full-page view with caption Click here for full-page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full page view with caption Click here for full-page view with captionClick here for full page view with caption Click here for full page view with caption Click here for full-page view with caption Click here for full-page view with caption Click here for full-page view with caption

Anaesthesia of bears is not particularly difficult, but it is important to recognise the potential dangers and consider the safety of both personnel and the bear. (D156.w2, J213.4.w3)
  • Note: the need for anaesthesia of zoo bears for purposes such as movements, routine examinations and simple procedures can be reduced by use of positive reinforcement training. (J4.223.w2, P20.1998.w11, P82.7.w1, W643.June06.w4) See: Mammal Handling & Movement - Husbandry Training
For information on bear anaesthetic emergencies see: ANAESTHETIC CRISES IN BEARS

This section is divided into:

  • PROCEDURES FOR ANAESTHETISING BEARS
    • Potential risks to be considered when anaesthetising bears
    • Handling and physical restraint
    • Pre-anaesthetic preparation
    • Anaesthetic drug administration
    • During induction
    • Anaesthetic monitoring and support
    • Maintenance and inhalational anaesthesia
    • Local and regional anaesthesia
    • Transport of anaesthetised bears
    • Reversal of anaesthesia
    • During recovery
  • ANAESTHETIC DRUGS FOR BEARS
    • List of anaesthetic regimes (with links to individual technique pages)
    • ORAL SEDATION
    • INJECTABLE ANAESTHESIA
    • BEAR SPECIES-SPECIFIC NOTES
  • ANAESTHETIC CRISES IN BEARS
PROCEDURES FOR ANAESTHETISING BEARS 

Before anaesthetising a bear, consider whether the benefits of the immobilization outweighs the risks. (P106.2007.w5)

POTENTIAL RISKS TO BE CONSIDERED WHEN ANAESTHETIZING BEARS

Risks to the bear from the anaesthetic

  • Bears are monogastric and may vomit during induction or recovery, or regurgitate while anaesthetised. If possible, avoid anaesthetising bears which have eaten recently. (D156.w2, J213.4.w3)
  • There is a potential risk that the bear may need to be killed to protect human life if there are problems with inadequate anaesthesia. (D156.w2)
  • Note: Risks are increased when the bear is not healthy.
  • For information on bear anaesthetic emergencies see: ANAESTHETIC CRISES IN BEARS
  • Physical injury, sometimes severe or even fatal, can occur when bears are darted. (P9.2004.w4, J40.32.w1, D249.w10)

Risks to the bear from the environment

  • Consider the risks of the anaesthetised bear being attacked if other bears are nearby. (D156.w2)
  • Bears are at risk of injury if they can reach a hazard between injection of the anaesthetic drugs and the time they become recumbent. This may occur in captivity as well as in the wild. Hazards to be considered include water (ponds, streams, water troughs etc.), cliffs and trees (which the bear could climb and then fall off/out of). (D249.w10)
  • It is important to consider the risks of the bear becoming recumbent against a fixed object (e.g. a wall or door, or in the soil) in a posture that will restrict its breathing. (D249.w10)
    • It the bear's breathing is compromised so that it in danger of suffocation, but it is not sufficiently anasethetised to handle, it may be possible to reposition the bear's head from a distance using a stick. Unless it is danger of suffocation, leave it alone. (D249.w10)
  • Additionally, if a bear is in an inside area, consider whether it will be possible to get into the area with the bear if it is recumbent against the door, and whether it is possible to do so safely.

Risks to humans

  • There is always a potential risk to personnel when dealing with bears. (D156.w2)
  • Particular care must be taken when the bear appears to be anaesthetised and is first approached.
  • Note: With some drug combinations (xylazine-ketamine and medetomidine-ketamine) bears may arise suddenly with little or no warning. (D156.w2)
  • Consider the risks of other nearby bears attacking personnel. This is a particular concern if a cub is anaesthetised and its mother is nearby. (D156.w2)
HANDLING AND PHYSICAL RESTRAINT
  • Only young bears (cubs) can be manually restrained, using gloves, or for slightly older/larger bears, a snare or net. (B64.26.w5, B123.19.w19, B429.3.w3)
  • In captivity, larger bears may be restrained in a squeeze cage to allow injection of immobilising drugs. (B123.19.w19, B429.3.w3)
  • Free-living bears may be caught in a snare prior to injection of immobilising drugs. (D156.w2)
  • See also Manual Restraint information in Mammal Handling & Movement (Mammal Husbandry and Management) - Holding & Carrying.
PRE-ANAESTHETIC PREPARATION
  • Whenever possible, the bear should be moved to a safe, quiet, well-controlled situation such as an indoor den with good lighting and ventilation, to allow a quiet induction and recovery, without the risk of the bear encountering hazards such as ponds or trees (which can be climbed then fallen out of) while semi-sedated, and where there will be no interference from other animals in the enclosure. (B407.w18, D247.7.w7)
    • It is important to ensure that it will be possible to get into the den after the animal becomes recumbent. (B407.w18)
      • "Moving a 600 kg polar bear wedged against a door can be very difficult!" (B407.w18)
  • Preventing/reducing vomiting:
    • Avoid anaesthetising immediately after the bear has eaten, to reduce the risks associated with vomiting and regurgitation during induction, anaesthesia or recovery. (D156.w2)
      • Preferably starve for 24 hours before anaesthesia. (B407.w18)
      • Withhold water for eight hours and food for 24 hours before immobilization. (D247.7.w7)
      • In one study with, vomiting was seen in four of eight Ursus americanus - American black bears fed within four hours of anaesthesia, but not when fed 12 hours prior to anaesthesia. (J1.15.w11)
    • Administration of 2-30 mg metaclopramide (depending on the size of the bear), five minutes before induction or mixed with the anaesthetic agents, may prevent vomiting during induction; a second dose may be given just before recovery/reversal to prevent or reduce vomiting during recovery. (J213.4.w3)
      • If there is a suspected gastrointestinal obstruction and metaclopramide is contraindicated, acepromazine, 0.01 mg/kg given with medetomidine-tiletamine-zolazepam has been used successfully to minimise or prevent vomiting. (J213.4.w3)
ANAESTHETIC DRUG ADMINISTRATION
  • Reliable, accurate drug delivery is important. (D156.w2)
  • Hand injection (intramuscular) is sometimes possible with wild bear cubs (following anaesthesia of their dam). (J40.35.w10)
  • A pole syringe may be used to inject wild bear cubs (following anaesthesia of their dam) and for bears with a limited range of movement, e.g. during restraint in a foot snare or in a small cage. (J1.33.w17, J2.30.w6, J40.35.w10, P9.2004.w4, P20.1998.w10)
  • Remote injection systems (darting) may be used to inject immobilising drugs without any form of prior physical restraint in either captive or free-living bears. (D156.w2)
    • For polar bears in the wild, darting from a helicopter is used most commonly. (B406.37.w37)
  • An alternative to manual restraint in captive bears is to give the bear honey mixed with an appropriate amount of a potent trans-mucosally absorbed narcotic agent (carfentanil) to produce sedation/light anaesthesia sufficient for approach and manual injection. (D156.w2) see below: Anaesthetic drugs for bears - Oral sedation
  • In most circumstances, remote drug delivery systems will be used. (J1.16.w14, J1.16.w15, J1.21.w8, J1.33.w16, J1.33.w17, J4.175.w2, J40.53.w2, J59.19.w1, J345.14.w6, P504.2001.w5)
    • Low impact systems delivering drugs at a low velocity are preferable if this will not compromise the safety of the bear or personnel. (D156.w2)
    • Potent drug combinations are required, due to limited volumes. (D156.w2)
    • If a large, aggressive bear has been caught in a snare, drugs may be administered using a pneumatic pistol. (D156.w2)
    • For free-ranging bears which are not restrained, darting from a distance is advisable, using a CO2 or cartridge powered rifle. (D156.w2)
  • Oral administration of alphachloralose has been used to immobilize culvert-trapped Ursus americanus - American black bear. (N29.14.w1)

Sites for darting/intramuscular injections

  • For intramuscular injections it is important to be aware of the anatomy of bears, in particular the large quantities of subcutaneous fat which may be present over the rump and hind legs of hibernating species in late summer and winter, and polar bears at any time of year. Therefore injecting into the shoulder or neck muscles is preferable. (D156.w2)
    • Injection into the fat may be ineffective. (B16.9.w9)
    • A needle length of at least 7.5 cm (3.0 inches) is required to reach through the subcutaneous fat layer on adult bears. (B16.9.w9, B64.26.w5)
  • A preferred site is the triceps muscle area of the forelimb, dorsal to the elbow and caudal to the humerus and scapula. (B123.19.w19)
    • The hind limb should be avoided in captive bears since there may be a lot of fat present resulting in the drug being deposited in the adipose tissue rather than muscle. (B123.19.w19)
      • Fat deposits over the rump and thighs may be several inches thick. (B16.9.w9)
    • The neck and shoulder are preferred as there is less fat over the muscle. (B16.9.w9)
  • Darting into the neck gives the fastest and most predictable response when darting wild polar bears. (B406.37.w37)
  • The distal (lower) muscle masses of the hind leg may be used, aiming towards the rear of the leg to make sure the femur is not hit. (B345.2.w2)
  • The rump is not useful due to large fat deposits around this site. There may also be large fat deposits over the shoulders. (B345.2.w2)
  • In captive bears at short range, injection into the muscles of the forearm can be used, delivered by blowpipe; standard 5 cm 18 gauge or 19 gauge needles can be used. (B407.w18)
  • Note: The time to induction varies depending on the injection site. (B406.37.w37)
  • Intravenous injection of anaesthetic drugs has been used for induction. (J4.137.w2)
  • See: 
DURING INDUCTION
  • The bear should be left undisturbed during induction but monitored e.g. by looking through a peephole. Keep light, noise and movement around the bear to a minimum during induction.
  • If the initial anaesthetic dose fails to adequately immobilise the bear, a top-up dose is required. It is important not to underdose at this stage. It is suggested that if the bear is able to sit up or move substantially, a second dose should be equal to the first dose. If the bear is recumbent but reactive to stimuli a minimum 2/3 dose should be given. Generally bears can rouse extremely quickly from an apparently deep plane of anaesthesia and great care should be taken during induction and anaesthesia. (V.w6)

  • Assess the bear's depth of anaesthesia BEFORE entering the enclosure.
    • A bear may appear to be anaesthetised but may still react to noise or movement.
    • Once the bear is recumbent and an appropriate length of time has passed, prod the bear gently then more vigorously using e.g. a long broom handle, from outside the enclosure. If the bear does not respond to prodding of the body, prod the bear's ear. If the bear still does not respond, it should be safe to enter.

    (B185.37.w37, B407.w18, V.w6)

ANAESTHETIC MONITORING AND SUPPORT
  • Monitor body temperature, respiratory rate, heart rate, colour of mucous membranes, capillary refill time, jaw tone (muscle relaxation) and the palpebral reflex throughout the anaesthesia. (B185.37.w37) Use other devices such as capnography, ECG, pulse oximetry etc. as available. (P106.2007.w5)
  • Eye protection
    • Cover the eyes once the bear is unresponsive to tactile and auditory stimuli. (B185.37.w37) A blindfold reduces visual stimulation and helps to protect the eyes. (D156.w2, J1.21.w7)
      • Eye lubrication (bland ophthalmic ointment) should be used, as well as a blindfold, to protect the eyes. (D156.w2, J1.15.w11, J1.17.w12, J1.21.w7)
  • Monitor depth of anaesthesia:
    • The depth of anaesthesia should be monitored at all times. (B185.37.w37, D156.w2)
    • With tiletamine-zolazepam, lightening of anaesthesia is indicated by spontaneous blinking, then chewing movements and paw movement, followed by lifting of the head and attempts to rise on the forelimbs. (D156.w2)
      • A top-up is required (tiletamine-zolazepam, or ketamine) if head movements are significant and further work is required. Ketamine is useful for an additional 5-20 minutes of anaesthesia, tiletamine-zolazepam can be used if at least 30 further minutes are required. (D156.w2)
    • With xylazine-ketamine or medetomidine-ketamine, arousal can be very sudden. (D156.w2) 
      • Early signs of arousal include increased palpebral reflex, or nystagmus. (D156.w2)
      • Do not approach the bear if it is showing signs of very light anaesthesia - head-lifting or limb movement. (D156.w2)
      • Arousal may be stimulated by:
        • Loud noises; (D156.w2)
        • Distress vocalisation by cubs of the bear;. (D156.w2)
        • Moving the bear or changing its position; (D156.w2)
        • Painful stimuli, such as tooth extraction. (D156.w2)
    • With medetomidine-tiletamine-zolazepam or xylazine-tiletamine-zolazepam
      • Signs of anaesthetic lightening include deep breathing, sighing, licking and development of a spontaneous palpebral. (D156.w2)
      • Head lifting or paw movement may indicate imminent arousal. (D156.w2)
      • Do not approach the bear if it is lifting its head.
  • Monitor respiration and oxygenation:
    • Monitor the respiratory rate and character (depth, regularity. (B407.w18)
    • Monitor the colour of the mucous membranes. (B407.w18, D156.w2, J213.4.w3)
    • Using a pulse oximeter probe on the tongue. (D156.w2)
    • Monitor arterial blood gases if possible. (B185.37.w37, J1.31.w11, J2.32.w2)
    • Give supplemental oxygen if the bear becomes hypoxaemic (haemoglobin saturation below 85%). (D156.w2)
      • A flow rate of 5-10 L/minute will be needed for most bears. (D156.w2)
        • In the field, an ambulance-type regulator and a lightweight, portable but sturdy D-cylinder are useful. This can provide a flow rate of 10 L/minute for up to 30 minutes; an E-cylinder can provide 10 L/min for an hour. (D156.w2)
      • Oxygen can be delivered via a nasal catheter placed into one nostril and passed up the nasal chamber as far as the medial canthus of the eye. (D156.w2)
      • Monitor the efficiency of the oxygen therapy by pulse oximetry. (D156.w2)
    • Note: with the alpha-2 agonist combinations, relative arterial saturation (SpO2) measured by pulse oximetry is often below 90%, but with nasal insufflation of oxygen rises to above 90%. Although SpO2 measured by pulse oximetry may not equate to direct measurements of arterial oxygen saturation, pulse oximetry does indicate the trend of oxygenation. Supplemental oxygen should be given whenever possible. (J213.4.w3)
  • Monitor the cardiovascular system:
    • Monitor the pulse/heart rate. (B185.37.w37, B407.w18)
    • A pulse can be palpated at the femoral artery or alternatively the brachial artery. (D156.w2)
      • With tiletamine-zolazepam, heart rates are usually 70-90 bpm. (D156.w2)
      • With medetomidine-tiletamine-zolazepam or xylazine-tiletamine-zolazepam, heart rates are usually 50-70 bpm. (D156.w2)
      • With medetomidine-ketamine, heart rates are lower, often 30-40 bpm. (D156.w2)
      • Bradycardia as low as 20-24 bpm has been seen with high dose orally administered carfentanil, but with lower doses heart rates of 40-88 bpm occurred. (J1.31.w11)
    • Monitor the capillary refill time. (B185.37.w37, J213.4.w3)
    • If possible, monitor peripheral blood pressure and ECG. (B185.37.w37)
    • Blood pressure can be measured via the femoral artery. (D156.w2)
      • Use a blood pressure cuff with a width about 0.4 times the circumference of the bear's limb. (D156.w2)
  • Monitor rectal temperature:
    • Bears are prone to hyperthermia because of their thick fat layer; close monitoring of body temperature during anaesthesia is important. (B10.48.w43, B407.w18)
      • Polar bears are particularly prone to hyperthermia while anaesthetised; monitor carefully. (B345.6.w6
    • With tiletamine-zolazepam, rectal temperature often decreases over time during the anaesthetic period. (D156.w2)
    • With medetomidine-tiletamine-zolazepam or xylazine-tiletamine-zolazepam, rectal temperature often decreases over time during the anaesthetic period.
    • In high ambient temperatures, the bear's body temperature may quickly reach dangerous levels (> 41C) (D156.w2)
      • Antagonise the alpha-2 agonist as soon as possible in bears with a high body temperature. (D156.w2)
    • Bears reaching a rectal temperature of 40.0 C or greater should be given treatment to reduce their body temperature. (J1.25.w6)
      • Hyperthermia in anaesthetised bears can be fatal; the bear may apparently recover from the anaesthetic but die in the next day or two. (J1.25.w6)
  • Ensure venous access:
    • Establishing an intravenous line early during the anaesthetic ensures ensures that venous access is available in the event of an emergency and drugs and fluids can be given rapidly if required. (B185.37.w37, V.w6)
    • This is particularly recommended during prolonged surgical procedures. (V.w6)
    • Appropriate veins include the cephalic (B185.37.w37), femoral vein or the medial saphenous on the inside of the hind leg. (V.w6)
    • See: Intravenous Injection of Bears
  • Give fluids:
    • During prolonged surgical procedures, fluids should be given intravenously (see section on fluid therapy, above). (P62.18.w1)
MAINTENANCE AND INHALATIONAL ANAESTHESIA
  • Anaethesia can be maintained for longer periods by either injection of further doses of anaesthetic agents or by the use of inhalant agents. (B16.9.w9)
  • Inhalation anaesthesia is recommended when long procedures are required, such as dental or orthopaedic operations or other surgery. (B16.9.w9, P62.13.w2)
  • If not intubating the bear, consider enriching inspired air with oxygen; a portable oxygen cylinder can be set to e.g. 2.0 - 4.0 litres per minute and delivered to the externl nares via a small rubber tube. (B185.37.w37)
Mask induction 
  • Inhalational anaesthesia can be used for induction of anaesthetic in young cubs, using inhalational anaesthetic (isoflurane) given via a face mask, while the cubs are hand-held (using gloves and blanket). (B336.51.w51, P62.13.w2)
  • Mask induction can be used if necessary in immobilized older bears to provide a deeper plane of anaesthesia before intubating. (B407.w18, P77.1.w19)
Use of short-acting anaesthetics
  • A short-acting anaesthetic can be given intravenously in an immobilized bear, to deepen the plane of anaesthesia and facilitate intubation. (P77.1.w19)

Intubation

  • Once a bear (any age) is anaesthetised, an endotracheal tube can be placed and the anaesthesia maintained via an inhalation agent. (B336.51.w51)
  • Intubation is recommended for anaesthetic maintenance. (B407.w18, J213.4.w3, P62.13.w2)
  • For an adult Ursus maritimus - Polar bear, an endotracheal tube of 11-14 mm is appropriate. (D315.3.w3)
  • To intubate a bear: (V.w6)
    • Place the bear in sternal recumbency, with the fore and hind legs placed careful at the sides of the bear so it is in a symmetrical position. (V.w6)
    • Have an assistant stand astride the bear, facing its head, with his/her feet inside the bear's elbows. (V.w6)
    • Place a rope behind the bear's upper canines and have the assistant lift the bear's head using this rope so the head is stretched forwards and up. (V.w6)
    • Take hold of the bear's tongue and pull it forwards and down. The larynx should now be clearly visible. (V.w6)
    • Pass an endotracheal tube of an appropriate size through the larynx and down the trachea. (V.w6)
Inhalation agents
  • Halothane or isoflurane may be used to prolong anaesthesia following induction using an injectable anaesthetic combination. (B16.9.w9, J213.4.w3)
  • Maintenance doses following use of commonly-used immobilisation drugs are isoflurane at 2-2.5% or halothane at 1% or less; higher concentrations can be used if required for deeper anaesthesia. (P1.1990.w5, P62.13.w2)
  • Note: Bears, even elderly individuals and those in poor condition, can tolerate long periods, even several hours, of gaseous anaesthesia, particularly if isoflurane is used. (B407.w18)

Additional notes

LOCAL AND REGIONAL ANAESTHESIA
  • Lumbosacral epidural anaesthesia has been used on a bear for analgesia during femoral head and neck excision. Medetomidine 5 g/kg and bupivacaine 0.25 mg/kg were used, mixed in the same syringe. This allowed reduction in maintenance isoflurane concentration (reduced from 2.5% to 1.4-1.8% 20 minutes after the epidural injection was given), provided excellent muscle relaxation during the operation, and provided postoperative analgesia for about 10-14 hours. (J2.32.w3)
TRANSPORT OF ANAESTHETISED BEARS
  • Bears transported while anaesthetised need to be monitored. (B407.w18)
  • Anaesthetised bears can be transported in a cargo net slung under a helicopter. HOWEVER this can cause hypertension and hypoxaemia and may cause mortality. (D156.w2, J1.35.w4, P20.1998.w8)
  • Anaesthetised bears should be transported with the head and neck extended to ensure a clear airway and with the body extended in either sternal or dorsal recumbency. (D156.w2)
    • A stretcher-type sling facilitates this positioning. (D156.w2)
  • If a reversible drug or drug combination is used to get a bear into a crate for transportation, the anaesthesia must be reversed and the bear fully recovered from the anaesthetic before the journey begins. (B407.w18, D247.10.w10)
  • If a free-living bear is to be transported inside a culvert trap, the anaesthesia should be reversed before transportation starts. If the bear is still anaesthetised it may move towards the end of the culvert, its neck can become flexed at it may lose airway patency and die. (D156.w2)
REVERSAL OF ANAESTHESIA
  • Reverse whenever possible in free-ranging bears, particularly if a sow with cubs has been anaesthetised, or if there is a high concentration of bears in the area. (D156.w2)
  • Giving 10 mg metaclopromide just before recovery reduces the risk of vomiting. (J213.4.w3)
  • Atipamezole is given at 3-4 times the dose rate of medetomidine used. It is safest to give the whole reversal dose intramuscularly. It can be given half intravenously, half intramuscularly, but this may lead to very rapid reversal. In an emergency, give intravenously. (D156.w2)
  • Atipamezole given intravenously may cause very rapid recovery, hyperexcitation and hypotension. Give only one tenth of the dose intravenously, the remainder intramuscularly. (J213.4.w3)
  • For very long procedures (over two hours) consider reversing the initial immobilising agents early, so that the animal is maintained on the isoflurane (i.e. the anaesthetic maintenance regime is simplified); this may make recovery at the end of the procedure more predictable. (J213.4.w3)
DURING RECOVERY
  • The bear should be left undisturbed, preferably in a cool, dimly lit area in which it can be kept under observation.
  • The bear's mouth and airways must be clear and its respiration monitored. 
  • The bear should not have access to food, water or other animals, nor be able to climb, until it is fully recovered. 
(B407.w18, D247.7.w7)

ANAESTHETIC DRUGS FOR BEARS

The choice of anaesthetic drug or drugs to use will depend on what the bear is being anaesthetised for (e.g. physical examination, surgical procedure) and personal choice - what the person carrying out the procedure is most comfortable with. (P106.2007.w5)

Details have been provided for the following anaesthetic regimes (in alphabetic order):

ORAL SEDATION
INJECTABLE ANAESTHESIA
  • This is the most common method for initiation of anaesthesia in bears.
  • Usually drugs are used in combination, mixed in the same syringe for injection. (J213.4.w3)
  • Most commonly, an alph-2 agonist is used in combination with a dissociative agent. (J213.4.w3)
  • N.B. Weights of bears may be overestimated when they have a thick coat, resulting in relative over-dosing with anaesthetic agents. (J59.24.w1)
  • NOTE:
    • If only a partial dose has been delivered, usually it is best to re-dart with the full dose: most of the drug combinations have wide safety margins; also, having been darted once the animal probably has raised catecholamines and may therefore suppress the effects of many drugs. (J213.4.w3)
    • If the initial anaesthetic dose fails to adequately immobilise the bear, a top-up dose is required. It is important not to underdose at this stage. It is suggested that if the bear is able to sit up or move substantially, a second dose should be equal to the first dose. If the bear is recumbent but reactive to stimuli a minimum 2/3 dose should be given. Generally bears can rouse extremely quickly from an apparently deep plane of anaesthesia and great care should be taken during induction and anaesthesia. (V.w6)

    • If the animal does not respond properly, consider terminating the procedure by administering the antagonist(s). (J213.4.w3)
  • Note: During the winter, torpid bears tend to require lower doses of anaesthetic. (B16.9.w9, D249.w10)
    • The onset of anaesthesia may take longer in hIbernating bears. (D249.w10)

Drugs and drug combinations

  • Note: medetomidine doses are usually given in microgrammes per kilogram bodyweight, indicated in the text below as " g/kg". Other drug doses are usually given in milligrams per kilogram bodyweight (mg/kg).
  • See BEAR SPECIES-SPECIFIC NOTES (below) for recommendations for individual bear species.
  • The advent of reversible anaesthetics has been advantageous in two ways in research on wild bears: as a matter or routine, it allows an individual bear to be processed more quickly, and if an individual bear is becoming unduly stressed physiologically under anaesthetic, the procedure can be shortened. (B406.37.w37)
  • Medetomidine 0.03 mg/kg plus tiletamine-zolazepam 3 mg/kg gives a predictable, smooth induction, excellent immobilisation and good recovery following reversal (atipamezole, 2.5 - 5 times the medetomidine dose on a mg-to-mg basis). (J213.4.w3)
  • Tiletamine-zolazepam, 4-6 mg/kg gives rapid immobilization; recovery to sternal recumbency with the head up takes about two hours. (J213.4.w3)
  • Xylazine plus Ketamine:
    • Xylazine 1-2 mg/kg plus ketamine 5 mg/kg immobilises most bear species. Antagonising the xylazine with e.g. atipamezole at 5:1 (atipamezole at five times the zylazine dose), or yohimbine  at 0.3 mg/kg, IV or IM, gives rapid reversal, to standing usually within less than 10 minutes can be used as an alternative to yohimbine. (J213.4.w3)
    • Xylazine 2 mg/kg plus ketamine 5-8 mg/kg has also been used, with the xylazine reversed with an alpha-2 antagonist e.g. yohimbine (0.1 mg/kg) or idazoxan (0.05 mg/kg). (B407.w18)
    • Note: rapid unexpected arousal can occur. (D156.w2)
    • [Editor's note: Atipamezole, rather than yohimbine, is now normally used for reversing alpha-2 agonist drugs.]
    • See: Xylazine-Ketamine Anaesthesia in Bears (Techniques)
  • Medetomidine plus Ketamine
  • Medetomidine plus Ketamine plus Midazolam has been used in bears, reversed with .
  • Etorphine hydrochloride) (M99) 
    • No longer recommended for use; better drugs are available. (B407.w18)
    • Can be reversed using diprenorphine (Revivon): give a volume of Revivon equal to the volume of Immobilon LA used. (B407.w18)
    • See: Etorphine Anaesthesia in Bears
  • Pentobarbital sodium for intravenous induction of anaesthesia: 8-10 mg/0.5 kg can be used; lower dosages may be required in obese bears. Give half the calculated dose rapidly, then the remainder to effect. If an immobilising agent has been used, only 1/3 to 1/2 the dose of pentobarbital sodium is needed, titrated to effect. (B16.9.w9) 4-5 mg/kg. (B64.26.w5)

Older drugs (now superseded)

  • Carfentanil has been used, 12-28 g/kg in Ursus maritimus - Polar bear. (B10.48.w43)
  • Fentanyl has been used:
    •  0.34-0.68 mg/kg in captive Ursus maritimus - Polar bear, giving smooth induction in 5-8 minutes. Injection of antagonist (naloxone, 25 mg naloxone per 10 mg of fentanyl given) gave arousal in 1-11 minutes, but one bear, on its feet six minutes after injection of the antagonist, relapsed and was recumbent overnight. (J4.175.w2)
    • Fentanyl plus azaperone has been used (15 mg fentanyl plus 60 mg azaperone in a 150 kg two-year-old female American black bear and 5 mg fentanyl plus 10 mg azaperone in a 47 kg male six-month old black bear). There was severe respiratory depression. The female was revived using 1.25 mg naloxone. (P1.1976.w2)
    • Fentanyl plus etorphine also has been used in captive Ursus maritimus - Polar bear. (J4.175.w2)
  • Fentanyl-droperidol (Innovar Vet, 0.4 mg/mL fentanyl + 20 mg/mL droperidol) (B121) has been used at 1.0 mL Innovar vet per 18.2 kg bodyweight, with an induction time of 10-15 minutes. (B16.9.w9)
    • Neuroleptanalgesia with Innovar Vet produces deep sedation with profound analgesia, appropriate for minor procedures such as lancing an abscess or carrying out diagnostic procedures (e.g. endoscopic examination), but not sufficient for major surgery. (B121)
    • Fentanyl may severely depress respiration. (B407.w18)
  • Note: Fentanyl may severely depress respiration. (B407.w18)
  • Phencyclidine-promazine is an old combination, used at 1.0 mg/kg of each agent and allowed minor surgical procedures including suturing of lacerations, and castration. With additional local anaesthetic, exploratory laparotomy was sometimes carried out. (B64.26.w5)
  • Phencyclidine was used alone at 0.5-1.0 mg/kg bodyweight intramuscularly. Severe convulsions were common, also depressed respiration and hypersalivation. (B16.9.w9)
    • Other common side-effects included excitement (increased by visual or auditory stimulation), hypertonicity of skeletal muscles, hyperthermia and vomiting. (P84.1.w1)
    • There were considerable variations in the dosages required to produce immobilisation and in the time to immobilisation. Convulsions were seen in 8/31 bears in one study (J40.32.w1) and in 20% of 400 Ursus americanus - American black bear immobilised in California. (P84.1.w2)
  • Note: Succinylcholine chloride has been used as an immobilising agent in bears. (J40.32.w2, J345.3.w4) This is a depolarising neuromuscular blocking agent, not an anaesthetic or analgesic agent. Animals immobilised with this drug are unable to move, and may be unable to breath, but are fully conscious. There is also a very narrow margin between an effective dose and a fatal dose, and respiratory failure sometimes occurred even at very low dose rates. (J40.32.w2, P84.1.w1) This is no longer considered a humane and appropriate method of restraint. 
BEAR SPECIES-SPECIFIC NOTES
  • Note: Preferred/recommended regimes are in bold.
  • All immobilising agents are given intramuscularly unless stated otherwise.
  • Medetomidine doses are mainly given in microgrammes per kilogram bodyweight, indicated in the text below as " g/kg". Other drug doses are usually given in milligrams per kilogram bodyweight (mg/kg).
Ursus thibetanus - Asiatic black bear
Drug 1* Drug 2* Reversal Notes Reference
Tiletamine-zolazepam, 4.4 mg/kg -- -- Ketamine, 2.2 mg/kg as a supplemental drug if required. (B345.6.w) B345.6.w6, B336.51.w51
Tiletamine-zolazepam 2.8 - 4.4 mg/kg -- -- -- D156.w2
Medetomidine 0.01 mg/kg Tiletamine-zolazepam 1.0 mg/kg Atipamezole In captive bears. This provides Stage 2/Stage3 anaesthesia for about 30-45 minutes, allowing physical examination or minor surgical procedures such as wound treatment, skin biopsy and castration.

For longer and/or more invasive procedures, anaesthesia is prolonged with inhalant anaesthesia.

V.w90
Xylazine 2 mg/kg estimated body mass Ketamine 4 - 5 mg/kg estimated body mass   For foot snared or barrel-trapped wild bears (J46.271.w1)
Xylazine 1 mg/kg Ketamine 15 mg/kg   For culvert-trapped wild bears J345.13.w6
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 
Ursus americanus - American black bear
Drug 1* Drug 2* Reversal Notes Reference
Xylazine 2.0 mg/kg  Ketamine 4.4 mg/kg Yohimbine, 0.15 mg/kg Ketamine 2.2 mg/kg as supplemental drug if required. 
This combination is recommended when short restraint times are needed e.g. for females with cubs. (J59.24.w1)
B345.6.w6
Tiletamine-zolazepam, 7.0 mg/kg     Induction may take 20 minutes and a long time may be required for recovery.
For most situations requiring safe and effective immobilisation. (J59.24.w1)
B345.6.w6, B336.51.w51
Medetomidine 0.04 mg/kg Ketamine 1.5 mg/kg.  Atipamezole 0.2 mg/kg. -- B345.6.w6, B336.51.w51
Etorphine 0.02 mg/kg   Diprenorphine 2 mg/kg (B345.6.w6). OR Naltrexone 100 mg/mg etorphine. (B336.51.w51) May cause respiratory depression. (B345.6.w6) B345.6.w6, B336.51.w51
Xylazine 2.0-4.5mg/kg  Ketamine 4.5-9.0 mg/kg Yohimbine, 0.125 mg/kg   B336.51.w51, J1.15.w11
Tiletamine-zolazepam 4.0-6.0 mg/kg.      Recovery can be prolonged.  D156.w2
Tiletamine-zolazepam 4.7 +/- 1.9 mg/kg     Induction time 14.5 +/- 12.7 minutes, in zoo bears. J1.16.w14
Medetomidine 52 g/kg  Tiletamine-zolazepam 1.7 mg/kg. Atipamezole, 3-4 x the medetomidine dose in g/kg Give intramuscularly unless in emergency; VERY rapid recovery can occur if the atipamezole is given intravenously. 
Induction in 6.3 +/- 3.3 minutes (range 1.5-9 minutes) in wild bears injected following capture in culvert traps. (J1.33.w16)
D156.w2, J1.33.w16
Xylazine 2.0 mg/kg  Tiletamine-zolazepam 3.0 mg/kg. Yohimbine 0.1-0.2 mg/kg, or with atipamezole. Recovery is slower than with medetomidine-tiletamine-zolazepam, but faster than for tiletamine-zolazepam alone. D156.w2
Carfentanil orally, 0.0068-0.019   Naltrexone 100 mg/ml carfentanil Delivered orally for transmucosal absorption. B336.51.w51
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 

Ursus arctos - Brown bear

Drug 1* Drug 2* Reversal Notes Reference
Tiletamine-zolazepam 8.0 mg/kg -- None Ketamine 2 mg/kg as a supplemental drug if required. B345.6.w6
Tiletamine-zolazepam 7.0-9.0 mg/kg -- None -- B336.51.w51
Tiletamine-zolazepam 7-10 mg/kg -- None -- D156.w2
Medetomidine 0.06 mg/kg  Tiletamine-zolazepam 2 mg/kg. Atipamezole 0.125 mg/kg. (B345.6.w6) Atipamezole 0.3 mg/kg (B336.51.w51) -- B345.6.w6, B336.51.w51
Medetomidine 35 g/kg Tiletamine-zolazepam 4.8 mg/kg Atipamezole, 3-4 x the medetomidine dose in ug/kg. Give intramuscularly unless in emergency. VERY rapid recovery can occur if the atipamezole is given intravenously D156.w2
Medetomidine 0.01 mg/kg Tiletamine-zolazepam 1.0 mg/kg Atipamezole In captive bears. This provides Stage 2/Stage3 anaesthesia for about 30-45 minutes, allowing physical examination or minor surgical procedures such as wound treatment, skin biopsy and castration.

For longer and/or more invasive procedures, anaesthesia is prolonged with inhalant anaesthesia.

V.w90
Xylazine 11 mg/kg  Ketamine 11 mg/kg Yohimbine 0.125 mg/kg This drug combination is not currently recommended [2002]. (D156.w2) B345.6.w6
Xylazine 0.3 mg/kg  Carfentanil 0.012 mg/kg Naltrexone or naloxone 100 mg per mg carfentanil given, plus yohimbine 0.125 mg/kg -- B345.6.w6, B336.51.w51
Etorphine 0.02 mg/kg  -- Diprenorphine 2 mg per mg etorphine given -- B345.6.w6
Etorphine 0.02 - 0.06 mg/kg    Naltrexone, 100 mg per 1 ,g etorphine given.   B336.51.w51
Xylazine 2 mg/kg Tiletamine-zolazepam 3 mg/kg. Yohimbine 0.1-0.2 mg/kg OR atipamezole Recovery is slower than with medetomidine-tiletamine-zolazepam, but faster than for tiletamine-zolazepam alone D156.w2
Tiletamine-zolazepam 3.5 +/- 1.8 mg/kg     Induction time 4.0 +/- 2.0 minutes. In nine zoo bears.  J1.16.w14
Carfentanil, 8 g/kg orally slowly, in honey     Causes hypoxia. Used in captive bears to avoid darting. D156.w2
Carfentanil orally (0.006 - 0.015 mg/kg   Naltrexone 100 mg per mg carfentanil given Delivered orally for transmucosal absorption. B336.51.w51
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 
Ursus maritimus - Polar bear
Drug 1* Drug 2* Reversal Notes Reference
Tiletamine-zolazepam 8 mg/kg -- -- Ketamine 2 mg/kg as a supplemental drug if required. (B345.6.w6) B345.6.w6, B336.51.w51
Tiletamine-zolazepam 8-10 mg/kg -- - -- D156.w2
Carfentanil 0.02 mg/kg.    Naltrexone or naloxone 100 mg per mg carfentanil given Severe respiratory depression may occur. Renarcotisation may occur; it is recommended that an additional dose of the antagonist should be given subcutaneously or intramuscularly. B345.6.w6, B336.51.w51
Medetomidine 0.03 mg/kg Ketamine 2.5 mg/kg Atipamezole 0.15 mg/kg Spontaneous recovery can occur. Loud or sharp noises should be avoided, and if possible avoid cubs vocalizing while their mother is anesthetized. (B345.6.w6) B345.6.w6, D156.w2, B336.51.w51
Medetomidine 0.012-0.159 mg/kg Ketamine 3.0-4.0 mg/kg Atipamezole 0.631 mg/kg   B336.51.w51
Etorphine 0.035 mg/kg   Diprenorphine 2 mg per mg etorphine given. (B345.6.w6) OR naltrexone 100 mg/mg etorphine given. (B336.51.w51)   B345.6.w6
Xylazine 7.0 mg/kg CUBS of the year: xylazine 3.0 mg/kg Ketamine 7.0 mg/kg CUBS of the year:  ketamine 3.0 mg/kg -- Xylazine-ketamine is not recommended. (D156.w2) B345.6.w6
Medetomidine 75 g  Tiletamine-zolazepam 2.2 mg/kg -- -- D156.w2
Xylazine 2.0 mg/kg  Tiletamine-zolazepam 3.0 mg/kg.  -- -- D156.w2
Xylazine 7.0-11.0 mg/kg Ketamine 7.0-11.0 mg/kg Yohimbine 0.125 mg/kg   B336.51.w51
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 

Melursus ursinus - Sloth bear
Drug 1* Drug 2* Reversal Notes Reference
Tiletamine-zolazepam 6.0 mg/kg -- -- Ketamine 2 mg/kg as a supplemental drug if required. (B345.6.w6) B345.6.w6, B336.51.w51
Tiletamine-zolazepam 5.5 - 6.6 mg/kg -- - -- D156.w2
Xylazine 2 mg/kg  Ketamine 7.5 mg/kg. Yohimbine 0.125 mg/kg. -- B345.6.w6, B336.51.w51
Medetomidine 0.07 mg/kg.  Ketamine 3.0 mg/kg Atipamezole 0.35 mg/kg. -- B336.51.w51
Xylazine 1.4-2.4 mg/kg Ketamine 5.8-9.7 mg/kg.  Yohimbine 0.1-0.2 mg/kg, or  atipamezole Recovery is slower than with medetomidine-tiletamine-zolazepam, but faster than for tiletamine-zolazepam alone. D156.w2
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 

Tremarctos ornatus - Spectacled bear
Drug 1* Drug 2* Reversal Notes Reference
Tiletamine-zolazepam 6 mg/kg     Ketamine 2 mg/kg as a supplemental drug if required B345.6.w6, B336.51.w51
Tiletamine-zolazepam 3.2-11.1 mg/kg       D156.w2
Tiletamine-zolazepam 2.8 +/- 0.5 mg/kg     Induction time 15.0 +/- 0.8 minutes J1.16.w14
Xylazine 0.6 - 3.0 mg/kg Ketamine 4.0 - 4.0 mg/kg     P77.1.w19
Tiletamine-zolazepam 2.0 - 3.4 mg/kg       P77.1.w19
Etorphine 0.2 mg/kg       P77.1.w19
Carfentanil 0.0085 mg/kg Azaperone 0.0005 mg/kg     P77.1.w19
Ketamine 3.5 - 7.0 mg/kg     Immobilization of young bears, 10-20 kg P77.1.w19
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.

 

Helarctos malayanus - Sun bear
Drug 1* Drug 2* Reversal Notes Reference
Medetomidine 0.07 mg/kg Ketamine 3.0 mg/kg Atipamezole 0.35 mg/kg, half the dose  intravenously and half intramuscularly. Ketamine 2.0 mg/kg as a supplemental drug if required. (B345.6.w6) B345.6.w6, B336.51.w51
Tiletamine-zolazepam 5.0 mg/kg       B345.6.w6, B336.51.w51
Tiletamine-zolazepam 4.0-5.5 mg/kg       D156.w2
Tiletamine-zolazepam, 4 mg/kg estimated body weight     For immobilisation of wild bears caught in culvert traps, to allow radio-collaring. Administered by pole syringe. J17.119.w1
Tiletamine-zolazepam 4.1+/-0.9 mg/kg     In zoo bears. Induction time 8.7 +/- 3.0 minutes. J1.16.w14
Tiletamine-zolazepam 3-5 mg/kg       D255.6.w6e
Medetomidine 60-80 g/kg Ketamine 2.0-3.0 mg/kg   Moderate sedation to complete immobilisation. Based on three immobilizations. J2.21.w3
Xylazine 2.2 mg/kg Ketamine 3-4 mg/kg   Preferably pre-treat with 0.1 mg/kg (or 10 mg total dose) metaclopramide to avoid vomiting. Quickly reversible with 0.2 mg/kg yohimbine D255.6.w6e
Medetomidine 0.01 mg/kg Tiletamine-zolazepam 1.0 mg/kg Atipamezole In captive bears. This provides Stage 2/Stage3 anaesthesia for about 30-45 minutes, allowing physical examination or minor surgical procedures such as wound treatment, skin biopsy and castration.

For longer and/or more invasive procedures, anaesthesia is prolonged with inhalant anaesthesia.

V.w90
Preferred/recommended regimes in bold
* All immobilising agents given intramuscularly unless stated otherwise.
ANAESTHETIC CRISES IN BEARS
It is important always to be ready for an emergency. (P106.2007.w5) Emergencies include:
  • Insufficient sedation following administration of appropriate agents; (J213.4.w3) 
    • If the attempt to anaesthetise the animal persists, it is likely to get hyperthermic. Dart it with the appropriate antagonist(s) and leave it to calm down. (J213.4.w3)
  • Hyperthermia or other metabolic derangement; (J213.4.w3)
  • Respiratory arrest in an animal which has not been intubated and which has failed to respond to intravenous diagram hydrochloride. (J213.4.w3)
    • Administer the appropriate antagonist(s), terminating the immobilisation procedure. (J213.4.w3)
  • Seizures: (D249.w13)
    • Wrap the bear in a tarpaulin to prevent it injuring itself during the seizure., keep the bear quiet, make sure the are no sharp objects nearby. Hold the head and make sure the eyes are protected. (D249.w13)
  • NOTE: The lingual veins are preferred for "crisis access" for administration of emergency drugs, as they are large and easily visible on the ventral surface of the tongue. (B407.w18, D255.6.w6e, V.w6, V.w91, V.w92)

Lagomorph Consideration

(Information on ANAESTHETIC EMERGENCIES is at the end of these Lagomorph Considerations.)

NOTE: The majority of the information below concentrates of anaesthesia of the domestic rabbit. Special considerations for wild lagomorphs are noted as appropriate. It is important to remember that wild lagomorphs, particularly free-living lagomorphs (e.g. wildlife casualties) will probably be highly stressed by being in a human contact/veterinary hospital situation.

Particular potential problems associated with anaesthesia in rabbits include:

  • Stress - from unfamiliar surroundings, presence of strange people and predator animals, rough handling, restraint and pain, as well as stress associated with surgery. (B600.5.w5)
    • Note: stress can lead to post-anaesthetic ileus. (J15.30.w2)
    • Reduce stress by keeping rabbits in a quiet area away from predator species, gentle handling and restraint, provision of appropriate analgesia, and if possible keeping the rabbit with a familiar rabbit companion. (J15.30.w2)
  • Hypoxia - from decreased oxygen tension associated with the anaesthetic agents, respiratory depression, breath-holding, airway occlusion due to poor positioning, reduced diaphragmatic movement due to weight of the viscera on the diaphragm from incorrect positioning, pre-existing respiratory disease, or firm restraint around the chest reducing respiratory movements. (B600.5.w5)
    • Note: rabbits have a small lung capacity, only 4 - 6 mL/kg. They also have a restricted nasopharynx, and even more so in breeds with a short nose. (B600.5.w5)
Controlled drugs
  • Fentanyl, pethidine and morphine are all Schedule 2 controlled drugs. These must be kept in a locked, immovable cabinet. Purchase and supply must be recorded in a register. To obtain these drugs, a written requisition, signed by a veterinary surgeon, is required. (B600.5.w5)
  • Buprenorphine and barbiturates are Schedule 3 controlled drugs. These must be kept in a locked, immovable cabinet. To obtain these drugs, a written requisition, signed by a veterinary surgeon, is required. (B600.5.w5)
  • To obtain diazepam, a written requisition, signed by a veterinary surgeon, is required. (B600.5.w5)
SEDATION
  • Fentanyl/fluanisone (Hypnorm, Janssen).
    • 0.2 - 0.3 mL/kg intramuscularly. (B600.5.w5)
    • This provides sedation and profound analgesia. (B600.5.w5)
    • Produces vasodilatation, making blood sampling or placement of an intravenous catheter easier. (B600.5.w5, B601.16.w16, J15.30.w2)
  • Acepromazine 0.5 mg/kg plus Butorphanol (Opiate analgesic) 0.5 mg/kg subcutaneously or intramuscularly. (B600.5.w5)
    • It is possible to give the two drugs mixed in the same syringe. Causes vasodilatation. (B600.5.w5)
  • Diazepam (Sedative)
    • 1 - 2 mg/kg intravenously or intramuscularly. Does not provide analgesia. (B600.5.w5)
    • 1-3 mg/kg by intramuscular injection - light sedation. (B609.2.w2)
  • Ketamine (Anaesthetic)
    • 25 - 50 mg/kg intramuscularly. (B600.5.w5)
    • Used in combination with other agents for anaesthesia. (B600.5.w5)
  • Midazolam (Sedative)
    • Can be used alone to sedate for minor procedures. (B600.5.w5)
    • 0.5 - 2.0 mg/kg intravenously. (B600.5.w5, B609.2.w2) Note: precipitates in Hartmann's solution. (B600.5.w5)
  • Ketamine 20-25 mg/kg plus Xylazine 2 mg/kg intramuscularly. (B604.3.w3)
    • For radiography. (B604.3.w3)
  • Ketamine 20-25 mg/kg plus Acepromazine 2 mg/kg intramuscularly. (B604.3.w3)
    • For radiography. (B604.3.w3)
  • Ketamine (15-20 mg/kg by intramuscular injection) and Midazolam (0.5 mg/kg by intramuscular injection). (B609.2.w2)
    • For deeper sedation and longer procedures. (B609.2.w2)
PRE-ANAESTHETIC PREPARATION
Before any anaesthetic, the rabbit's general health and physiological status should be assessed. If time allows, stabilisation (normalisation of body temperature, correction of dehydration and electrolyte imbalance, syringe feeding of anorectic animals) should be carried out. (J15.30.w2)

Feeding

  • Lagomorphs are unable to vomit, therefore pre-anaesthetic fasting is not required. (B538.59.w59, B600.5.w5)
  • Fasting may have a negative impact on gastro-intestinal function, leading to ileus, and should be actively avoided. (B538.59.w59, B600.5.w5, B601.16.w16)
  • Withholding food for one or two hours before anaesthesia ensures there is no food in the mouth, and that the stomach is not excessively full. (B600.5.w5, B601.16.w16) 30 minutes is sufficient to ensure no food is in the mout. (B539.1.w1)
Housing
  • If rabbits are in the veterinary hospital for any length of time prior to anaesthesia, they should be placed in a quiet area, on familiar bedding (usually hay), away from the sight, sound and smell of predators, including dogs, cats, ferrets, birds of prey etc. (B600.5.w5)
  • For wild lagomorphs, it is important that they should also be protected from human activities and noises as much as possible.
Pre-anaesthetic assessment
  • Assessment should include consideration of the rabbit's history to help identify risk factors predisposing to anaesthetic problems. (J15.30.w2)
  • Carry out a full clinical examination. (B539.1.w1, B601.16.w16, J15.30.w2) See: Physical Examination of Mammals
    • Note respiratory rate and character in the undisturbed rabbit. (B601.16.w16)
      • There should be only minimal movement of the thoracic wall; rate should be 30 - 60 per minute. (B539.1.w1)
      • Auscultate the thorax - use a paediatric stethescope. (B539.1.w1)
    • Remember that respiratory and heart rate are likely to be elevated when the rabbit is being handled and examined. (B601.16.w16)
    • Check the patency of the nares and nasopharynx, particularly if inhalation anaesthesia via a face mask will be used. (B538.59.w59)
    • Check the forelegs for staining indicating nasal discharge. (B539.1.w1)
    • Check the rabbit's body temperature (should be 38.5 - 40 C). (B539.1.w1)
    • Check the rabbit's hydration status. (B539.1.w1)
  • Consider what pre-existing disease conditions may affect the rabbit's physiological status. (B600.5.w5, B601.16.w16) For example: (B600.5.w5)
    • Dental or oral disease causing pain, malnutrition and often excessive salivation (therefore possibly dehydration and electrolyte imbalances). (B600.5.w5)
      • Dental problems such as spurs should be treated before other surgery is carried out, to avoid anorexia after surgery due to the tooth problems. (B539.1.w1)
    • Gastrointestinal disease which may cause dehydration and electrolyte imbalances. (B600.5.w5)
    • Respiratory disease - if nasal discharge is present there is likely to be an increased anaesthetic risk. (B601.16.w16)
    • Radiography may be needed if the are obvious respiratory problems. (B601.16.w16)
  • Weigh on accurate digital scales to ensure that injectable drug rates can be calculated accurately. (B539.1.w1, J15.13.w7, J15.30.w2)
Fluid and electrolyte balance
  • If possible, correct fluid and electrolyte imbalances before anaesthetising the rabbit. (B601.16.w16)
  • Give intravenous glucose if the rabbit is hypoglycaemic. (J15.30.w2)
  • Give a blood transfusion if considered necessary due to anaemia from acute blood loss. (J15.30.w2)
  • Place an intravenous catheter (see: Intravenous Injection and Catheterisation of Rabbits) or if this is not possible, an intraosseous catheter (see: Intraosseous Catheterisation and Administration of Medication in Rabbits). (B601.16.w16)
    • Intravenous access should always be secured before anaesthesia in individuals which are physiologically unstable, those where a prolonged anaesthetic period is expected, and when an operation which may be expected to produce significant haemorrhage is to be performed. (B602.33.w33)
  • Consider giving fluids e.g. Hartmann's solution (lactated Ringer's solution) 10 mL/kg. (B545.8.w8) See Fluid Therapy section above.
Pre-medication
  • This is advisable to reduce stress during restraint and induction. (B601.16.w16)
  • Pre-medication agents which may be used include:
    • Acepromazine maleate:
      • Sedative; potentiates the effects of other anaesthetic agents and facilitates a smooth recovery. Hypotensive. No analgesic action. (B600.5.w5)
      • 0.1 - 0.5 mg/kg intramuscularly. Provides moderate to mild sedation. Causes peripheral vasodilatation; use with care in individuals which are dehydrated or have cardiovascular disturbances. (B601.16.w16)
      • 0.5 - 1.0 mg/kg intramuscularly or subcutaneously. Note: no analgesic action. (B600.5.w5)
      • A phenothiazine tranquilizer. These cause marked peripheral vasodilatation and thereby hypotension. They have a long duration of action. (J34.23.w1)
      • 0.1 mg/kg subcutaneously or intramuscularly. (J34.23.w1)
    • Butorphanol
      • 1 mg/kg intramuscularly plus acepromazine maleate 0.5 mg/kg intramuscularly. (B601.16.w16)
      • Provides moderate sedation and some analgesia. Causes peripheral vasodilatation; use with care in individuals which are dehydrated or have cardiovascular disturbances. (B601.16.w16)
    • Diazepam
      • 1-2 mg/kg intramuscularly or intravenously. (B601.16.w16)
      • Provides moderate to deep sedation. If using intravenously, use the emulsion formulation to minimise the risk of thrombophlebiasis. (B601.16.w16)
      • A benzodiazepine sedative. These decrease anxiety. Used in combination with dissociative anaesthetics to increase duration and improve muscle relaxation, but they do not have any analgesic effect. (J34.23.w1)
    • Fentanyl/fluanisone (Hypnorm):
      • 0.2 - 0.5 mL/kg intramuscularly. (B601.16.w16)
      • Provides mild to profound sedation and moderate to marked analgesia; may provide sufficient analgesia for minor surgery. At high dose rates, can cause marked respiratory depression. (B601.16.w16)
    • Fentanyl/droperidol (Innovar vet):
      • 0.22 mL/kg intramuscularly. (B601.16.w16)
      • Provides mild to profound sedation and moderate to marked analgesia; may provide sufficient analgesia for minor surgery. At high dose rates, can cause marked respiratory depression. (B601.16.w16)
    • Ketamine:
      • 15 - 30 mg/kg intramuscularly. (B601.16.w16)
      • Provides moderate to heavy sedation and some analgesia. (B601.16.w16)
      • Dissociative anaesthetic. Poor muscle relaxant. Does not abolish ocular, laryngeal or swallowing reflexes. Sympathomimetic effect - raised heart rate, blood pressure and cardiac output when used alone. For surgical anaesthesia it is usually used in combination with Medetomidine or Xylazine. (B600.5.w5)
      • Note: recovery is prolonged in individuals with renal impairment, due to renal excretion of ketamine and its metabolites. (J204.47.w1)
    • Medetomidine:
      • 0.1 - 0.5 mg/kg intramuscularly or subcutaneously. (B601.16.w16)
      • Provides mild to profound sedation. Causes peripheral vasoconsriction which may make intravenous access difficult. May cause respiratory and cardiovascular depression; preferably avoid use in individuals in poor health. (B601.16.w16)
      • Can be reversed with Atipamezole. (B600.5.w5)
    • Midazolam:
      • 2 mg/kg intravenously, intramuscularly or by intraperitoneal injection. (B601.16.w16)
      • Provides moderate to deep sedation. Has the advantage (compared with diazepam) of being water soluble). (B601.16.w16)
      • A benzodiazepine sedative. These decrease anxiety. Used in combination with dissociative anaesthetics to increase duration and improve muscle relaxation, but they do not have any analgesic effect. (J34.23.w1)
    • Xylazine:
      • 5 mg/kg intramuscularly. (B601.16.w16)
      • Provides mild to profound sedation. Causes peripheral vasoconstriction which may make intravenous access difficult. May cause respiratory and cardiovascular depression; preferably avoid use in individuals in poor health. (B601.16.w16)
      • Usually used in combination with ketamine. (B600.5.w5, J34.23.w1)
      • Can be reversed with Atipamezole. (B600.5.w5)
Anticholinergics:
  • These are given to reduce salivation and bronchial secretions, and for prevention of vagally-mediated bradycardia. (B601.16.w16, J15.30.w2)
  • Atropine:
    • 0.05 mg/kg, subcutaneously or intramuscularly. (B600.5.w5, B601.16.w16)
    • Note: ineffective in many rabbits, due to the presence of endogenous atropinase. (B601.16.w16, J15.30.w2)
  • Glycopyrrolate:
    • 0.01 mg/kg intravenously or 0.1 mg/kg subcutaneously or intramuscularly. (B600.5.w5, B601.16.w16); 0.01 - 0.02 mg/kg subcutaneously. (J15.30.w2)
    • Glycopyrrolate has a slower onset of action but longer duration of effect than atropine. (J34.23.w1)
    • Note: does not cross the blood-brain barrier. (B600.5.w5)
Pre-emptive analgesia
  • Advisable prior to painful surgery. (J15.30.w2)
    • Use of pre-emptive analgesia may reduce the anaesthetic requirements of patients undergoing surgery. (J4.219.w4)
  • Opioids should be considered as part of the pre-medication protocol prior to surgery if opioids will not be part of the main anaesthetic protocol. (B601.16.w16)
  • NSAIDs should be given pre-operatively if these are used to provide analgesia. (B601.16.w16)
    • Avoid giving NSAIDs pre-operatively if adequate blood pressure and therefore renal perfusion cannot necessarily be maintained during anaesthesia. (B601.16.w16)
HANDLING AND RESTRAINT FOR ANAESTHETIC INDUCTION
  • Ensure than handling is always gentle and quiet to minimise stress. (B600.5.w5, J15.30.w2)
  • Minimise physical restraint of wild (free-living) lagomorphs. (B538.59.w59)
  • Note: an unsedated rabbit exposed to volatile anaesthetic agents will struggle. It is important that the handler be aware of the need for firm restraint. This situation (exposing an unsedated rabbit to vapours which it finds noxious) should be avoided if possible. (B601.16.w16)
  • For free-living lagomorphs, handling cn be minimised by placing the lagomorph into an induction chamber for anaesthesia with a volatile agent, or by placing the live trap, containing the individual, into the induction chamber. (B538.59.w59)
  • For further information see:
ANAESTHETIC MONITORING AND SUPPORT
  • Constant monitoring of depth of anaesthesia, heart, respiration and if possible oxygenation is important; temperature should also be monitored. (B601.16.w16)
  • For treatment of respiratory and cardiac arrest see ANAESTHETIC EMERGENCIES

Eye protection
  • Protect the eyes, which are prominent and prone to abrasion and drying during anaesthesia: apply an ophthalmic ointment or artificial tears, or tape the eyelids shut with micropore tape. (B601.16.w16, J15.30.w2)
    • It is particularly important to protect the eyes when ketamine is used, since the eyes will remain open. (J15.30.w2)
    • When a face mask is used, drying of eyes is increased and they must be protected. (J15.30.w2)
  • Note: the rabbit can maintain the palpebral reflex even at dangerously deep levels of anaesthesia, therefore ocular reflexes are of little use in monitoring rabbit anaesthetic depth. (B601.16.w16)
Monitoring anaesthetic depth
  • At an anaesthetic depth sufficient for major surgery, the ear pinch response is abolished and the hindlimb pedal withdrawal is absent or nearly absent. (B601.16.w16)
  • Loss of the forelimb withdrawal reflex indicates deeper anaesthesia; this level of anaesthesia is not generally required. (B601.16.w16)
  • In adequate anaesthetic depth and/or inadequate analgesia is indicated if a stimulus results in sudden tachycardia, hypertension or tachypnoea. (B602.33.w33)
Respiration
  • It may be difficult to monitor respiration by observation of chest movements since these may be very shallow and the rabbit may be obscured by drapes. (B601.16.w16)
  • Nasal movements associated with respiration do not indicate respiratory depth and are not reliable. (B601.16.w16)
  • A stethoscope or oesophageal can be used. (J15.30.w2)
  • Pulse oximetry: 
    • This is a simple, effective means of monitoring respiratory function, indicating oxygenation; it also indicates heart rate. (B600.5.w5, B601.16.w16, B602.33.w33, J290.32.w4)
    • It is important to ensure that the instrument is capable of giving an accurate oxygen saturation reading at high heart rates (> 250 bpm). (B601.16.w16)
    • Sites
      • The sensor can be placed on the pinna (ear), buccal mucosa or tongue to measure saturated oxygen. (J213.1.w1)
      • The tongue is the preferred site, if available. (B600.5.w5)
        • An angled reflectance probe, lodged between the tongue and the hard palate, gave consistent data. (J290.32.w4)
      • The pinna (ear) can be used. (B600.5.w5)
        • Readings from the pinna may be lower than those in the mouth. (J290.32.w4)
      • The ventral base of the tail can be used. (B600.5.w5); this should be clipped. (B600.5.w5)
      • A digit can be used in white laboratory rabbits, but is less reliable in rabbits with pigmented skin and in small individuals. (J290.32.w4)
      • A rectal probe can be used. (B600.5.w5)
    • Note: Peripheral vasoconstriction associated with ketamine/medetomidine or ketamine/xylazine anaesthesia (Medetomidine - Ketamine Anaesthesia in Rabbits) may cause failure of pulse oximetry, particularly in smaller individuals, or result in abnormally low oxygen saturations being recorded. (B600.5.w5, B601.16.w16, J290.32.w4)
  • A respiratory monitor using a thermistor sensor can be used with rabbits larger than 500 g but are not reliable in smaller rabbits or if respiration is depressed. (B601.16.w16)
  • Capnography can be used and provides an indirect indication of oxygenation. (J15.30.w2)
  • A fall in respiratory rate to less than 40% of the resting/pre-anaesthetic rate of the patient suggests impending respiratory failure. (J15.13.w7)
  • A rise in respiratory rate in response to surgical stimulus indicates inadequate anaesthetic depth or inadequate analgesia. (J15.13.w7)
  • Treatment of respiratory depression: 
    • Usually there is a reduction in respiratory rate or depth before respiratory arrest; however, some animals develop apnoea without much warning. (B601.16.w16)
    • Manual assistance of respiration - gently squeeze the reservoir bag (preferred), or intermittently occlude the outflow from a T-piece or Bain's circuit. (B601.16.w16)
    • Use a mechanical ventilator if available. (B601.16.w16)
    • If the rabbit is not intubated, be aware that assisted intermittent positive pressure ventilation may cause stomach inflation and further problems; as an alternative, use chest compression: place the rabbit on its back, head and neck extended, and gently squeeze the chest at 40-60 times per minute. (B601.16.w16) Additionally, the rabbit can be rocked to and fro so that the abdominal contents give intermittent compression of the diaphragm. (B601.16.w16)
  • Treatment of respiratory arrest: SEE ANAESTHETIC EMERGENCIES

Pulse/Heart:
  • Palpate over the central auricular artery in the ear (gently). (B600.5.w5)
  • Directly auscultate the chest (B600.5.w5); a good quality paediatric stethoscope is appropriate. (B602.33.w33, B538.59.w59)
  • An oesophageal stethoscope can be used. (B602.33.w33, B538.59.w59)
  • The heart rate is typically 240-280 bpm; it may be lower (120-160 bpm) when medetomidine is used. (B600.5.w5, (B601.16.w16)
  • Pulse oximetry can be used (B600.5.w5, B601.16.w16, B602.33.w33) - see above for details. 
    • Note: this does not indicate whether peripheral perfusion is adequate. (B602.33.w33)
  • Doppler flow probe
    • A Doppler flow probe over a peripheral artery may be most useful in smaller rabbits. (B601.16.w16)
    • This can be placed over the ventral tail base, carotid, femoral or auricular arteies, or directly over the heart. B538.59.w59
  • Mucous membranes
    • Assess the colour of the mucous membranes of the nose, lips or tongue. (B600.5.w5)
    • Capillary refill time can be used for clinical assessment of circulatory function. (B601.16.w16)
  • ECG:
    • This can be used to monitor the heart. (B600.5.w5, B602.33.w33
    • It is necessary to use an ECG instrument able to detect low signal strength and high frequencies. (B601.16.w16)
    • Use standard lead positions. B538.59.w59
    • The electrocardiograph must be able to record at 100 mm/second and amplify the signal to at least 1 mV equal to 1 cm; preferably a multichannel oscilloscope with non-fade tracing and freeze capabilities is used. (B538.59.w59)
  • Arterial blood pressure:

    • Normal systolic pressure in adults 110 mmHg (range 90 - 130 mmHg); diastolic pressure in adults 80 mmHg (range 60 - 91 mmHg). (B611.3.w3)

    • Direct measurement of arterial blood pressure can be carried out using a catheter placed in the central artery of the ear. (B601.16.w16) See: Arterial Catheterization of Rabbits (Techniques)

  • Indirect measurement of blood pressure can be carried out: (J15.27.w1)

    • Place a neonatal size 1 or 2 blood pressure cuff, attached to a manometer, on the rabbit's foreleg, just proximal to the elbow.

      • The cuff's width should be equal to 50 per cent of the circumference of the leg at this point.

    • Apply coupling gel to the palmar surface of the leg, just proximal to the carpus. Usually it is not necessary to clip the fur.

    • Place a Doppler ultrasound transducer over the palmar common digital arteries; make sure sufficient coupling gel is present.

    • Inflate the blood pressure cuff to occlude the vessels in the forelimb

    • Slowly deflate the cuff; read the blood pressure indicated on the manometer at the point when the Doppler ultrasound transducer detects resumption of blood flow (which occurs when the pressure in the cuff drops to systolic pressure).

    • Repeat several times for a mean value.

    (J15.27.w1)

    • Alternative sites are the hind leg, tail or ear. (B538.59.w59)
  • Treatment of bradycardia
    • If bradycardia which may be vagally-induced occurs during abdominal or cervical surgery, glycopyrrolate, 0.01 mg/kg intravenously or 0.1 mg/kg intramuscularly or subcutaneously can be given. If this is not available, atropine can be given; due to the individually-variable production of atropinase in rabbits, give 0.05 mg/kg subcutaneously or intramuscularly initially, assess the effect and increase the dose if required. (B601.16.w16, J15.30.w2)

  • Treatment of cardiac arrest: SEE ANAESTHETIC EMERGENCIES

Temperature:
  • It is important to monitor temperature. (J15.13.w7, J15.23.w6)
  • Anaesthetised animals are unable to use muscle activity to generate heat; small animals (under 5 kg, i.e. most rabbits) have a large surface area to volume ration therefore are more prone to hypothermia. (J15.23.w6)
  • A standard rectal thermometer can be used. (B600.5.w5, J15.30.w2)
    • Either a mercury-in-glass or a digital thermometer can be used. Mercury thermometers are relatively fragile. Digital thermometers have a relatively slow response. (J15.23.w6)
    • It may be difficult to access the rectum during surgery. (J15.23.w6)
    • Falsely low reading may occur due to faeces or gas in the rectum. (J15.23.w6)
  • If available, a digital thermometer with remote sensor can be used. Lubricate the remote sensor and carefully place it in the rectum to allow continuous monitoring. (B600.5.w5)
  • Thermistor and thermocouple probes may be used in the rectum, oesophagus or nasopharynx. (J15.23.w6)
  • Electronic monitors attached to a probe and with settable upper and lower temperature alarms can be used. (J15.13.w7)
  • Or if using an oesophageal stethoscope, a temperature probe may be attached to this. (B602.33.w33)
  • If two probes are available, one can be used to measure core body temperature and the other attached to a distal limb to measure peripheral temperature; this provides additional information on peripheral blood flow and circulation. (J15.23.w6)
  • Hypothermia is a particular risk with small rabbits (B601.16.w16), if internal organs are exposed for long periods, and if fluids are used for abdominal lavage without first being warmed to body temperature.
  • Avoid hypothermia by placing the anaesthetised rabbit on a safe heating pad. (B601.16.w16)
    • Electrical heat pads, microwavable heat pads, hot water bottles and latex gloves filled with hot water ("hot hands") can be used; all should be wrapped in a towel to avoid the possibility of contact burns. (J15.23.w6, J15.30.w2)
    • Water-circulating blankets can be used, but may restrict surgical access. (J15.23.w6)
    • Microwavable hot packs (e.g. wheat-filled) can be used. (J15.23.w6)
    • Latex or nitrile gloves filled with hot water can be placed alongside the rabbit. (J15.30.w2)
    • Warm-air circulating blankets can be used but may restrict access to the patient. (J15.23.w6, J15.30.w2)
    • Heat lamps can be used, with care to avoid overheating or burning. (J15.23.w6)
    • Maintain body heat using e.g. blankets, or bubble wrap. (J15.23.w6)
    • Avoid excess clipping of hair for surgery. (J15.23.w6)
  • Maintain body temperature at 39 C during anaesthesia. (B601.16.w16)
  • Note: adverse effects of hypothermia include:
    • general depressive effect; this decreases the anaesthetic required; recovery from anaesthesia may be prolonged if hypothermia develops. (J15.23.w6)
    • predisposition to cardiac arrhythmias. (J15.23.w6) increased clotting time. (J15.23.w6)
  • Consider monitoring the environmental temperature; provide a room temperature of about 24 C while the rabbit is anaesthetised. (J15.30.w2)
  • Fluids:

    • If fluid loss is expected, an intravenous catheter should have been placed prior to anaesthesia. (B601.16.w16)

    • Consider giving body-temperature dextrose saline at the end of surgery, subcutaneously or intraperitoneally, to ensure the rabbit does not become dehydrated in the immediate post-operative period before it resumes normal water intake. (B601.16.w16)

  • Blood loss:

    • If blood loss is anticipated during surgery, initiate infusion of 10 - 15 mL/kg/hr warmed lactated Ringer's solution once the rabbit is anaesthetised. (B601.16.w16)

    • Monitor blood loss by weighing swabs. (B601.16.w16)

    • If significant blood loss occurs, a whole blood transfusion may be needed (usually an initial transfusion from a single donor provides only a low risk of adverse reaction. (B601.16.w16)

    • If whole blood is not available. give a synthetic haemoglobin glutamer (Oxyglobin). (B601.16.w16)

LOCAL ANAESTHESIA
  • Local anaesthetics can be used as an adjunct to general anaesthetics (B601.16.w16) to reduce the depth of anaesthesia required during painful surgical procedures.
  • Note: It is important, particularly with smaller individuals, to carefully calculate the dose of any local anaesthetic drug used to ensure that the total used does not reach a toxic dose - give maximum 2 mg/kg bupivacaine, or 10 mg/kg Lidocaine (Lignocaine). (B601.16.w16)
  • EMLA cream (ASTRA Pharmaceuticals Limited, King's Langley, England) is a local anaesthetic cream that contains lidocaine (lignocaine) and prilocaine and can produce full thickness skin anaesthesia. (B600.3.w3, J15.20.w2, J83.24.w1)
    • It has been recommended for use on the marginal ear vein venepuncture site (B600.3.w3, B601.2.w2, J83.24.w1) and the lateral saphenous site. (B601.2.w2)
    • Apply over the site 30 minutes (B601.16.w16, J83.31.w2) 45 to 60 minutes (B600.3.w3, J15.20.w2, J83.24.w1) prior to venepuncture (after clipping) and cover with an occlusive dressing or cling film. (B600.3.w3, B601.16.w16, J15.20.w2, J83.24.w1)
    • Prior to venepuncture, clean the skin with a cotton swab (J83.24.w1) and wipe the site with 70% isopropyl alcohol. (B601.2.w2)
Anaesthetic Induction
  • Injectable anaesthetic agents should be used whenever possible to avoid the problem of breath-holding which is common when rabbits smell anaesthetic vapours. (B600.5.w5, B601.16.w16, J83.30.w2)
  • If induction with a gaseous anaesthetic agent is used, the rabbit should be premedicated with an appropriate sedative. (B600.5.w5, B601.16.w16, J15.30.w2)

Anaesthetic induction in wild lagomorphs

  • For wild lagomorphs, an injectable agent or combination is appropriate (B284.10.w10, B538.59.w59) and can be given intramuscularly. (B538.59.w59)
  • Induction with inhalant anaesthetics is appropriate when rapid induction and rapid recovery is required, for free-living lagomorphs. (B538.59.w59) 
    • However as with domestic rabbits this is stressful and causes breath-holding. Therefore if this is used, the animal should be sedated beforehand using e.g. fentanyl/fluanisone, or acepromazine. (B284.10.w10)
    • Gaseous induction may be appropriate when very rapid recovery is essential. (V.w5)
INJECTABLE ANAESTHESIA

Balanced anaesthesia, using an appropriate combination of agents, is generally recommended in rabbits, rather than use of a single agent. (B601.16.w16)

  • Details of the use of individual injectable anaesthetic agents are given in:
  • Alternatives:
    • Fentanyl/Fluanisone - Diazepam:
    • Ketamine 50 mg/kg plus Acepromazine 1 mg/kg, or ketamine 25 mg/kg plus midazolam 5 mg/kg or ketamine 25 mg/kg plus diazepam 5 mg/kg. All given intramuscularly. (B601.16.w16)
      • Provide light to moderate anaesthesia for 20 -40 minutes; may not provide surgical anaesthesia. (B601.16.w16)
      • Useful for non-painful procedures such as radiography. (B601.16.w16)
      • Produce less respiratory depression than other combinations producing deeper anaesthesia; supplemental oxygen should still be given. (B601.16.w16)
      • Recovery time usually 2 - 3 hours. (B601.16.w16)
    • Diazepam 0.2 - 0.5 mg/kg intravenously (through an intravenous catheter) followed by Ketamine 10 - 15 mg/kg. (J34.23.w1)
    • Xylazine 1 - 5 mg/kg plus Ketamine 20 - 40 mg/kg, intramuscularly, then mask with isoflurane. (J34.23.w1)
      • Note: myocardial necrosis and fibrosis has been seen following multiple ketamine-xylazine anaesthetics (using 50 mg/kg ketamine and 10 mg/kg xylazine intramuscularly) in Dutch belted rabbits. (J495.49.w1)
    • Thiopentol 30 mg/kg intravenously using a 1.25% solution (B601.16.w16); 50 mg/kg (J204.47.w1) or methohexital 10 - 15 mg/kg intravenously, using a 1% solution. (B601.16.w16)
      • For induction followed by endotracheal intubation and maintenance with a volatile anaesthetic agent. (B601.16.w16)
      • Provides 5 - 10 minutes light to moderate anaesthesia if no other agent is used. (B601.16.w16)
        • 5 - 15 minutes with 0 mg/kg thiopentol. (J204.47.w1)
      • Recovery is rapid. (J204.47.w1)
      • Often excitement on recovery. (B601.16.w16)
      • Excitement can be reduced by pre-medication with a sedative (e.g. acepromazine maleate). (B601.16.w16)
      • Severe thrombophlebitis may occur if thiopentone is given extravascularly. (B601.16.w16)
      • (J204.47.w1)
    • Alfaxalone alone (Alfaxan, Vetoquinol UK) has been used in rabbits. 
      • 6.0 - 9.0 mg/kg intravenously, or 9.0 mg/kg intramuscularly. (B546)
      • In a study, following premedication with 0.03 mg/kg buprenorphine, alfaxanone was given intravenously at 2.0 or 3.0 mg/kg over a period of 60 seconds. Rabbits showed apnoea for about 45 seconds (range 10 to 120 seconds) following administration of the alfaxalone. All rabbits could be intubated (blind method). Isoflurane at about 3.0 % produced a surgical plane of anaesthesia following alfaxanone anaesthetic induction. Basic cardiopulmonary parameters were noted to remain stable and within the normal range. Recovery was rapid in some rabbits (e.g. lifting head within 3-5 minutes; mean was about 15 minutes). Rabbits were standing by about 35-40 minutes (mean; range 10 - 69 minutes) after anaesthesia. It was noted that very rapid recoveries could be avoided by use of premedication with a greater sedative effect, for example an alpha-2 adrenergic agonist. (J3.163.w1)
    • Alfaxalone plus medetomidine.
      • In a study in wild rabbits (Oryctolagus cuniculus - European rabbit), rabbits were give medetomidine 0.5 mg/kg subcutaneously and alfaxalone was given (5 mg/kg) intramuscularly for induction. Anaesthesia was maintained with isoflurane at 1.5 - 3.0% in oxygen (as required to maintain a surgical plane of anaesthesia). Following surgery (abdominal implantation of temperature loggers), the medetomidine was reversed with 5 mg/kg Atipamezole. Recovery took 27.8 +/- 15.7 minutes. Anaesthetic nduction was considered to be smooth, and recovery uneventful; the combination was considered to be safe and effective. (J3.164.w1)
  • Anaesthetics not recommended
    • Alfaxalone-Alphadolone (Saffan) 12 mg/kg intravenously provides rapid induction (within seconds) and light to medium anaesthesia for eight to 10 minutes. Muscle relaxation is good but analgesia is not always adequate. Full recovery requires 2.0 - 2.5 hours. (J83.12.w1)
      • Note: this combination has a poor safety margin in rabbits; when given at higher dose rates it can cause sudden apnoea which may be rapidly followed by cardiac arrest. (B601.16.w16, J83.27.w2)
      • This is not recommended for use in rabbits. (B600.5.w5, B601.16.w16)
    • Tiletamine/Zolazepam 5 - 25 mg/kg intramuscularly or intravenously. (B538.59.w59, B602.33.w33)
  • Reversal agents:
    • Atipamezole
      • 1 mg/kg, subcutaneously, intravenously or intramuscularly. For the reversal of medetomidine. (B600.5.w5)
    • Naloxone
      • 10 - 100 ug/kg intramuscularly, intravenously or by intraperitoneal injection. Reversal of fentanyl (and other narcotic analgesics). (B600.5.w5)
    • Buprenorphine (Opiate analgesic)
      • 0.01 - 0.05 mg/kg intravenously or subcutaneosuly. For reversal of fentanyl but with continued provision of analgesia (for 6 - 12 hours). (B600.5.w5)
    • Butorphanol (Opiate analgesic)
      • 0.1 - 0.5 mg/kg intravenously, intramuscularly or subcutaneosuly. For reversal of fentanyl but with continued provision of analgesia (for 2 - 4 hours). (B600.5.w5)
    • Doxapram (CNS Stimulant). 5 mg/kg intramuscularly or intravenously. To reverse the respiratory depressant effects of anaesthetic drugs. Effects last about 15 minutes, after which injection can be repeated. (B600.5.w5)
GASEOUS ANAESTHESIA
Induction of anaesthesia
  • Induction of anaesthesia using gaseous agents should be avoided in rabbits. With either halothane (all concentrations) or isoflurane (concentrations >0.5%), rabbits showed an aversion to the agent, attempting to avoid it, struggling, and showing periods of apnoea of 30-120 seconds at a time (sometimes repeatedly) until loss of consciousness occurred. Pronounced bradycardia (55 - 82% decrease in heart rate), hypercapnoea and acidosis occurred during the periods of apnoea. Hypoxia did not occur, but arterial pO2 remained low despite use of 100% oxygen as the carrier gas. (J83.20.w1)
  • Many rabbits exposed to halothane pawed at their face, tried to remove the mask or tried to escape from the anaesthetic induction chamber; it was considered probable that they would have would have had increased catecholamine levels, which could increase the risk of cardiac arrhythmia. (J83.20.w1)
  • If a volatile anaesthetic agent is to be used for induction, preferably pre-medicate with e.g. midazolam or fentanyl/fluanisone. (J15.30.w2)
  • Induction can be speeded up by using higher concentrations of the anaesthetic, but it is important to remember that these agents "cause a dose-dependent cardiovascular and respiratory depression" so overdose can be fatal.(J15.30.w3)
  • Nitrous oxide can be used (at 50:50 with oxygen) to smooth induction with a volatile anaesthetic agent. Once the rabbit is fully anaesthetised, the nitrous oxide should be switched off and 100% oxygen used as the carrier gas. If nitrous oxide has been used pure oxygen should be given for at least 10 minutes before the end of anaesthesia. (B600.5.w5)
  • For further information on induction see: Inhalational Anaesthesia Induction in Rabbits (Techniques)
Maintenance of anaesthesia
  • Volatile anaesthetic agents can be used for maintenance (prolongation) or deepening of anaesthesia following induction with injectable agents. (J15.30.w2)
  • It is important to remember that "All the commonly used inhalation anaesthetic agents cause a dose-dependent cardiovascular and respiratory depression." (J15.30.w3) Therefore overdose can be fatal. (J15.30.w3)
  • Halothane
    • Provides moderate muscle relaxation. N.B. dose-dependent cardiopulmonary depression, and sensitises the myocardium to catecholamines. (J34.23.w1)
    • Maintenance at 1 - 2%. (B538.59.w59, J34.23.w1)
    • Now superseded by other agents, particularly isoflurane. (B600.5.w5, J15.30.w3)
  • Isoflurane (J34.23.w1)
    • Can be used in converted halothane vapourisers (e.g. Fluotech) or in Isotec vapourisers. Care should be taken when using in converted halothane vapourisers as the setting goes higher (8%) than required with isoflurane. (J15.30.w3)
    • Causes greater respiratory depression than halothane. (J15.30.w3)
    • Licensed for use in rabbits in the UK. (J15.30.w3, W713.Oct08.w1)
    • Maintenance at 2 - 3% (B538.59.w59)
  • Sevoflurane
    • In the UK, presently [2008] licensed for dogs only. (J15.30.w3)
    • Needs its own vapouriser (vapour pressure is markedly different from halothane or isoflurane), with concentration settings up to 8%. (J15.30.w3)
    • May produce more stable anaesthesia and faster , more complete recovery than isoflurane.
    • Maintenance at 3-4%. (B538.59.w59)
  • Desflurane
    • In the UK, presently [2008] not licensed for use in animals. (J15.30.w3)
    • Rapid complete recovery. (J15.30.w3)
    • Requires a special vapouriser which needs electric power to work, since it involves heating the liquid to gas. Gives an output of up to 14%, twice the minimum alveolar concentration (MAC) for most species. (J15.30.w3)
  • Use of 100% oxygen as the carrier gas is preferred since respiratory disease is common in rabbits. (B601.16.w16)
  • Nitrous oxide
    • Use of nitrous oxide in rabbit anaesthesia is not recommended. (B601.16.w16)
    • Nitrous oxide has only a low potency in rabbits, minimally reducing the concentration of volatile agents used for anaesthetic maintenance. (B601.16.w16)
    • Nitrous oxide may diffuse into the stomach (J204.47.w1), caecum or other gas-filled spaces. (B600.5.w5)
    • Nitrous oxide can cause gastric stasis. (B600.5.w5)

Health implications and scavenging

  • N.B. Exposure to gaseous anaesthetic agents may have health implications for the anaesthetist and other people exposed to the anaesthetic agents. It is suggested that "reasonable measures should be taken both to reduce the risk of serious contamination of the atmosphere with inhalation anaesthetics and to remind operating theatre staff of possible hazards." (B121)
  • The "reasonable measures" include filling vapourizers using proper filling apparatus or funnels, outside the operating theatre and preferably out-of-doors, turning vapourizers off when not in use, taking care when handling anaesthetic agents, using low flow systems when possible, using scavenging of waste gases/vapours, using endotracheal intubation rather than a face mask when possible and checking breathing circuits regularly for leaks. (B121)
    • Active scavenging should be used when volatile anaesthetic agents are given through a face mask. (J15.13.w7, J15.30.w2)
    • Note: nitrous oxide is not removed by activated charcoal in anaesthetic gas scavenging systems. (B601.16.w16)
ANAESTHETIC EQUIPMENT AND USE

Circuits:

  • Use a circuit with a low dead space; remember that rabbit have a tidal volume of only 4-6 mL/kg. (B600.5.w5)
  • A T-piece circuit on a normal small animal anaesthetic machine is appropriate. (J15.30.w2)
  • A Bain circuit can be used. B600.5.w5
  • Use paediatric connectors. B600.5.w5

Face mask:

  • This can be used to deliver oxygen and anaesthetic gases to an anaesthetised rabbit. (J290.32.w3)
  • Use of a face mask for anaesthetic induction (i.e. delivering anaesthetic gases to a conscious rabbit) is not a preferred technique. If it must be used, the rabbit should be sedated prior to induction to reduce struggling (and the risk of related physical injury); this will not however prevent breath holding and hypercapnoea. (B601.16.w16) 
    • Note: breath holding may not be detected when a rabbit is being physically restrained for mask induction. (J83.20.w1)
  • Small, close-fitting face masks should be used. (J15.30.w2)
  • Clear masks are preferred as they permit better patient monitoring. (J15.30.w2)
  • A small mask can be used over the nares if access to the mouth is needed. (J15.30.w2, J513.2.w2)

Endotracheal tubes for intubation: 

  • Intubation reduces dead space compared to a face mask and helps maintain the airway.
  • Use uncuffed tubes in most rabbits to allow as large an internal tube diameter as possible. (B601.16.w16)
  • For rabbits of 1.0 - 3.0 kg, uncuffed tubes of 1.5 - 3.0 mm are appropriate. (J15.30.w2)
  • In large rabbits (rabbits over 5 kg), a cuffed tube can be used (internal diameter at least 4 mm). (B601.16.w16)
  • It may be necessary to shorten the tube to reduce the functional dead space within the anaesthetic circuit. (J15.30.w2)

Laryngoscope, otoscope or endoscope:

Laryngeal mask:

  • This can be placed through the rabbit's mouth and positioned over the larynx. (B601.16.w16)
  • It may be easier for an inexperienced anaesthetist to place a laryngeal mask than to intubate a rabbit. (B601.16.w16)
  • In a study, this provided better oxygenation than a face mask, and avoided the hypercapnoea seen with a face mask. However, use of intermittent positive pressure ventilation (IPPV) through the laryngeal mask produced gastric tympany in four of six rabbits (leading to gastric reflux in one rabbit). (J290.32.w3)

Nasal catheter:

  • Useful during oral surgery to deliver oxygen without obstructing access to the oral cavity. (B601.16.w16)
  • Lubricate a catheter of suitable size (e.g. in a 2.0 kg rabbit, 2.5 mm catheter) with lidocaine gel. (B601.16.w16)
  • Gently insert the catheter into the nares. (B601.16.w16)
  • Ensure entrance to the ventral nasal meatus by lifting the muscular nasal fold and directing the tube ventrally and medially as much as possible. (B601.16.w16)
  • The tube can be passed through the nasal passages, through the pharynx and into the trachea. (B600.5.w5, B601.16.w16)
    • 1 mm tube for rabbits under 1 kg bodyweight, 2 mm for larger rabbits. (J213.10.w2)

Mechanical ventilator:

  • To provide intermittent positive pressure ventilation. (J15.30.w2)
    • e.g. SAV03 Small Animal Ventilator, Vetronic Services Ltd., Newton Abbott, UK. (J15.30.w2)
EFFECT OF POSITIONING
  • Ensure that the rabbit's head is not flexed too far and obstructing the airway.
  • Avoid positioning the rabbit so that the contents of the abdomen push against the diaphragm; rabbits have a small thorax and large abdomen, and respiratory movements of the rabbit are mainly diaphragmatic, not costal. Pressure from the abdominal contents can reduce respiration. If possible, particularly if the rabbit is in dorsal recumbency, elevate the head and thorax relative to the abdomen, reducing pressure on the diaphragm from the abdominal contents (B539.1.w1, J15.30.w2)
RECOVERY
Place the patient in a quiet, dimly-lit area and monitor until fully recovered. (B545.8.w8)
  • Oxygen
    • Continue to provide supplemental oxygen via a facemask or in an oxygen chamber until the rabbit shows normal respiratory rate and pattern. (J15.30.w2)
    • If the rabbit is known or suspected to have developed low arterial oxygen saturation during the anaesthetic, oxygen supplementation should be continued for a longer time. (J15.30.w2)
  • Temperature
    • Provide supplemental heat until the rabbit is sufficiently recovered to resume normal thermoregulation. (B600.5.w5, J15.30.w2)
      • Take care not to overheat the animal, remembering that rabbits are susceptible to hyperthermia. (B600.5.w5, J15.30.w2)
      • Remember that rabbits can chew through electrical wires. (B600.5.w5)
  • Fluids:
    • Provide as required. (J15.30.w2)
  • Feeding:
    • Syringe feed if needed until self-feeding occurs. (J15.30.w2)
    • Consider using prokinetics to reduce the risk of post-anaesthetic ileus developing. (J15.30.w2)
      • Metoclopramide 0.5 mg/kg subcutaneously every 8 -12 hours. (J15.30.w2)
      • Ranitidine 2 mg/kg orally or subcutaneously every 12 hours. (J15.30.w2)
      • Cisapride 0.5 mg/kg orally every 8 - 12 hours. (J15.30.w2)
    • Offer tempting foods: as well as making hay available, offer fresh grass, dandelions, fresh vegetables and any prefered foods of the individual rabbit. (B600.5.w5)
  • Bedding:
    • Hay provides security, if familiar, and can be eaten as a source of indigestible fibre. (B600.5.w5)
  • Note: recovery may be prolonged in individuals with renal or hepatic disease in which drug metabolism may be slower than normal. (J15.30.w2)
  • Monitor for signs of pain and give appropriate treatment if required - see Analgesia section above
POST-OPERATIVE ANALGESIA
  • Post-operative analgesia is very important to restore appetite and gastro-intestinal motility as well as to reduce pain and stress. (B600.5.w5)
  • Careful observation is needed to detect subtle signs of pain. (B600.5.w5)
  • For further information on post-operative analgesia see section above: Analgesia
ANAESTHETIC EMERGENCIES:

The goals of cardiopulmonary resuscitation, as in other animals, are to provide the rabbit with ventilation and circulatory support until spontaneous cardiovascular function returns. (J213.1.w1)

SPECIFIC IMMEDIATE RESPONSE TO RESPIRATORY ARREST
  • Check the plane of anaesthesia - the rabbit may be breath-holding in response to the smell of inhalational anaesthetic agents, if it is lightly anaesthetised.
    • Check the anaesthetic is not too deep. (B601.16.w16)
  • Check there is a clear airway.
    • Clear the airway/endotracheal tube if this is blocked.
    • If the rabbit is not intubated, extend the head and neck and pull the rabbit's tongue forwards. (J15.13.w7)
    • If the airway is obstructed and cannot be cleared, consider tracheotomy with a large-bore hypodermic needle inserted in to the trachea.
    • Check there is no physical interference with respiratory movements. (B601.16.w16) Note: rabbit respiratory movements are mainly diaphragmatic; positioning with the caudal part of the body raised can increase pressure on the diaphragm from abdominal organs. 
  • Check that the circuit is patent and that oxygen is still being supplied. (B545.8.w8)
  • Check the heart and pulse - respiratory arrest can be followed rapidly by cardiac arrest. (B601.16.w16)
  • Gently compress the chest between thumb and a finger (one compression per second) to move air into and out of the lungs; this may stimulate breathing.
  • Give 100% oxygen via the endotracheal tube or a face mask (without any volatile anaesthetic agent).
  • If the rabbit is deeply anaesthetised and not intubated, attempt intubation.
  • If the rabbit is intubated, start IPPV (intermittent positive pressure ventilation).
  • Give Doxapram, 5 mg/kg (Dopram-V, 0.25 mL/kg, intravenously or intramuscularly). Note: drops of doxapram can be placed on the oral or nasal membranes for mucosal absorption.
    • The effects of doxapram last about 10 - 15 minutes, after which the dose may need to be repeated. (B601.16.w16, B545.8.w8)
  • If respiratory depression/arrest is due to an opioid (fentanyl) give an antagonist.
    • Note: consider the implications of reversing anaesthesia during surgery. (J15.13.w7)
  • If the rabbit is not intubated and an endotracheal tube cannot be passed, consider tracheotomy with a large-bore hypodermic needle inserted in to the trachea.
    • If an endotracheal tube cannot be passed, consider retrograde guided intubation: puncture the trachea just below the larynx using a 17-gauge needle catheter. Pass a guide wire or catheter up through the trachea and larynx and into the mouth, and then pass an endotracheal tube down through the larynx over the guide wire/catheter. (B538.59.w59, J83.35.w2)

(B121, B600.5.w5, J15.13.w7)

SPECIFIC IMMEDIATE RESPONSE TO CARDIAC ARREST

Note: this can follow rapidly after respiratory arrest. (B601.16.w16)

  1. Check the rabbit's airway is unobstructed.
  2. Give 100% oxygen - by endotracheal tube if placed, otherwise by facemask
  3. If the rabbit is not already intubated, try to place an endotracheal tube.
  4. Start IPPV (intermittent positive pressure ventilation).
  5. Give external cardiac massage, 70-90 per minute.
  6. If anaesthetic overdose is suspected, give the specific antagonist. (J15.13.w7)
    • Give Atipamezole intravenously if possible, otherwise intramuscularly or subcutaneously, if an alpha-2 agonist has been used in the anaesthetic protocol.
    • Note: consider the implications of reversing anaesthesia during surgery. (J15.13.w7)
  7. Give adrenaline, 0.2 mL/kg of a 1: 10,000 solution: intravenously, subcutaneously or squirted into the trachea. Note: solutions are provided at a 1:1,000 dilution; dilute this 1:9 with sterile water to give a 1:10,000 dilution.
    • Lignocaine 1.0 - 2.0 mg/kg if ventricular tachycardia. (J15.13.w7)
  8. Consider percentage blood loss and give fluid therapy as required to support the circulation. (B601.16.w16, J15.13.w7)
    • 50 mL/kg over one hour in the treatment of hypovolaemia, otherwise 10 - 15 mL/kg/hr. (B545.8.w8)
    • See: Fluid Therapy section above for more details on fluid therapy.

(B545.8.w8, B600.5.w5, B601.16.w16, J15.13.w7)

EMERGENCY DRUGS
  • Administer reversal agent if injectable anaesthetic has been used:
  • Naloxone (pure opioid antagonist) for opioids, total dose 2 mg, slow intravenous injection.  Reverses respiratory depression caused by opioids. (J83.23.w2)
  • Atipamezole for alpha-2 agonists, intravenously, intramuscularly or subcutaneously N.B. also antagonizes the analgesia provided by the alpha2 agonist.
  • Doxapram (CNS Stimulant): short-acting respiratory stimulant. 5-10 mg/mg (P3.1999b.w2) 7 mg/kg (0.3 mL/kg): dilute 1:3 and give by slow intravenous injection or intramuscularly, or in smaller birds dropped onto the tongue may help stimulate respiration.
  • Adrenaline may be given, 0.5-1.0 mg/kg intravenous, in response to cardiac arrest, anaphylactic shock or bronchial spasm (P3.1999b.w2).
  • Dexamethasone (Corticosteroid) 4 mg/kg (1 mg/kg in raptors) intramuscular or subcutaneous in case of shock. (P3.1999b.w2).
  • Dextrose: administer in the case of seizures due to hypoglycaemia.
  • Diazepam (Sedative): first-line treatment for seizures, including epileptic fits, and all other seizures except those caused by Strychnine and hypoglycaemia
  • Atropine: Anticholinergic. Initial bradycardia due to central effects, followed by tachycardia due to blockage of cardiac muscarinic receptors. Also relaxation of bronchial smooth muscle. 0.5 mg/kg intramuscular
  • Prednisolone sodium succinate (Solu-Medrone): 2-4 mg/kg intramuscular. Synthetic water-soluble corticosteroid. Treatment of shock, endotoxaemia, spinal cord compression.
  • Sodium bicarbonate 1-4mg/kg slow intravenous injection

(P3.1999b.w2).

CHECK EQUIPMENT FOR FAILURE
  • Endotracheal tube:
  • Correct placement in trachea, not oesophagus?
  • Too long or placed too deep (causing bronchial intubation)/
  • Too narrow?
  • Obstructed?
  • Kinked?
  • Disconnected from anaesthetic machine?
  • Vaporizer:
  • No anaesthetic agent?
  • Incorrect setting on dial?
  • Wrong agent in vaporizer?
  • Inaccurate vaporizer calibration?
  • Anaesthetic machine:
  • No oxygen in cylinders?
  • Flow meter - incorrect siting?
  • Flow meter - failed?
  • Connections (vaporizer-machine, or breathing system) - leakage?
  • Breathing system obstruction?

(B121, P3.1999b.w2, J15.13.w7)

Ferret Consideration
Pre-anaesthetic preparation
  • The ferret should be given a pre-anaesthetic physical examination, including body weight measurement, thoracic auscultation and abdominal palpation, with particular attention to the cardiovascular and respiratory systems. (B232.18.w18, B602.33.w33, J15.24.w5, J29.7.w1); their clinical history should be reviewed. (B602.33.w33)
    • A complete blood count and serum biochemistry should be carried out; assess hepatic and renal function. (B232.18.w18, B629.13.w13)
  • Obtain an accurate body weight for dosing. Note that in winter there may ebe a higher percentage of body fat, so more anaesthetic may be required. (J15.24.w5)
  • Ferrets should be fasted for 4-6 hours before induction of anaesthesia, as they may vomit during induction. (B232.18.w18, J29.6.w3)
    • Do not fast for longer periods; they may become agitated if hungry, and seriously hypoglycaemic if they have an insulinoma. (J29.6.w3)
    • Gut transit time is short and six hours easily empties the gastrointestinal tract. (B602.1.w1)
    • Fasting for up to four hours is adequate. (B602.33.w33, J15.24.w5)
    • The ferret may be fasted for 1-4 hours pre-surgery; ferrets with islet cell neoplasia (Insulinoma in Ferrets) should be fasted for a maximum of two hours. (B631.22.w22)
    • Give free access to water untill immediately before anaesthesia. (B232.18.w18)
  • If time allows, the ferret should be stabilised before anaesthesia: dehydration, electrolyte imbalances, hypoglycaemia etc. should be corrected. (B602.33.w33)
  • Ferrets stressed by transport or poor handling should be given time to calm down, as high circulating catecholamines increase anaesthetic doses needed and thereby increase the risks of side-effects. (B232.18.w18)
  • To minimise heat loss and the risk of hypothermia (Chilling - Hypothermia (with special reference to Waterfowl, Hedgehogs, Bears, Lagomorphs and Ferrets)), the area of fur clipped for surgery should be minimised and alcohol rinses avoided during aseptic preparation. (B631.23.w23)
Premedication

Premedications are useful to reduce fear and anxiety (sedatives), ensuring the ferret is calm at the time of anaesthesia, also to reduce the dose of the induction agent, reduce the vagal reflex (which is strong in ferrets), produce a smoother recovery, reduce bronchial secretions and provide post-operative analgesia. Premedication used should be chosen depending on the procedure to be carried out - e.g. sedation only for a short, non-painful procedure, or sedation plus opiate analgesia plus an antimuscarinic agent before major abdominal surgery. (B232.18.w18, B631.22.w22, J15.24.w5)

  • Atropine Sulphate should be given prior to anaesthesia (0.05 mg/kg subcutaneously, intravenously or intramuscularly) to reduce salivation and bradycardia. (B631.22.w22) The airways of ferrets are small and easily blocked, therefore use of a drying agent is recommended before anaesthesia. (J15.24.w5)
    • Atropine sulphate can be given 20 minutes prior to anaesthesia, to reduce the gag reflex and production of saliva, making intubation easier. (P120.2006.w7)
  • Famotidine (H2 blocker) and diphenhydramine (antihistamine) are useful prior to surgery alongside atropine, to reduce stress-related oesophageal reflux, gastric irritation and subsequent histamine release. (B631.22.w22)
  • Diazepam 0.5 mg/kg intravenously. (J513.7.w3)
  • Midazolam 0.25 mg/kg intramuscularly or intravenously. (J513.7.w3)
  • Midazolam 0.2 mg/kg plus Ketamine 10 mg/kg, intramuscularly. These can be given mixed in the same syringe. This dose provides short-lasting sedation and relaxation and is useful for e.g. radiography. (B232.18.w18)
  • Midazolam 2 mg/kg to reduce anxiety and produce relaxation. (B232.18.w18)
  • Diazepam 2 mg/kg to reduce anxiety and produce relaxation. (B232.18.w18)
  • Medetomidine
    • 1 - 2 g/kg intramuscularly or intravenously. (J513.7.w3)
    • 100 g/kg intravenously, intramuscularly or subcutaneously, as premedication, also alone for minor non-painful procedures. (B232.18.w18)
  • Acepromazine 0.2 - 0.5 mg/kg subcutaneously or intramuscularly. Note this is hypotensive. (B232.18.w18)
    • Note: Use of acepromazine should be avoided in ferrets, due to vasodilatation. (B631.22.w22)
  • Xylazine20 - 30 mg/kg intravenously, intramuscularly or subcutaneously. Note: this is hypotensive. (B232.18.w18)
  • Prior to gaseous anaesthetic induction: butorphanol plus midazolam or diazepam:
    • Butorphanol 0.2 mg/kg subcutaneously, 20-30 minutes before induction, as an analgesic. (B631.22.w22) 
      • 0.2 - 0.8 mg/kg subcutaneously, intramuscularly or intravenously. (J513.7.w3)
    • Midazolam 0.25 - 0.3 mg/kg intramuscularly or intravenously, (B631.22.w22, J513.7.w3) 15-20 minutes before induction OR Diazepam 1-3 mg/kg intramuscularly or intravenously, to reduce anxiety and improve muscle relaxation. (B631.22.w22)
  • Fentanyl/Fluanisone 0.5 mL/kg intramuscularly; this produces neuroleptanalgesia but muscle relaxation is poor. (B232.18.w18)
Restraint
  • Usually, anaesthetic premedication drugs can be given intramuscularly or subcutaneously in the ferret under manual restraint. (B232.18.w18)
  • Ferrets can be scruffed, held on a table in lateral recumbency with one hand at the sruff, the other at the hips, or scruffed and then wrapped in a towel to allow injection of drugs. (J29.7.w1) 
  • If a ferret is agitated, frightened or aggressive, it may be restrained using thick gloves if necessary and given 0.5 mg//kg Midazolam plus 0.4 mg/kg Butorphanol for sedation to allow safe handling for induction. (B631.22.w22)
Anaesthetic monitoring & support

Good monitoring is very important during ferret anaesthesia. (B232.18.w18, J29.14.w2) Body temperature and the cardiovascular and respiratory systems should be monitored carefully. (B232.18.w18)

  • Always provide oxygen. Many anaesthetics cause respiratory depression, and the ferret may be in a poor state of health. (J15.24.w5)
  • Cardiac rate and rhythm should be monitored using:
    • Direct auscultation. (B629.13.w13)
      • An oesophageal stethoscope can be used if the ferret has been intubated. (J29.7.w1)
    • Doppler flow detector (B629.13.w13, J29.7.w1); this can be placed on a hind foot. (J29.14.w2)
    • ECG. (B629.13.w13, B631.22.w22) This should be capable of measuring heart rates over 400 bpm. (J29.14.w2)
  • Pulse oximetry can be used (B629.13.w13, B631.22.w22, J29.14.w2) if a sufficiently small sensor is available for attachment to the ferret's ear, cheek or tongue. (B631.22.w22) The sensor can be attached to a paw or the tail, but the contact area may need to be shaved for a good reading. (B629.13.w13)
    • Pulse oxymetry can be useful for monitoring both heart rate and blood oxygenation. (B629.13.w13)
    • If an ear-lob-type clip is used, reposition the clip periodically to avoid local ischaemia and associated inaccurate readings. (J29.7.w1)
  • Capnography can be used to monitor respiration once the ferret has been intubated. (B631.22.w22, J29.7.w1, J29.14.w2)
  • Blood pressure monitoring; this can be carried out via a cuff on the front leg or tail. (B631.22.w22, J29.7.w1)
  • The depth of anaesthesia should be monitored using reflexes, muscle tone and responses to surgical stimulation: (B232.18.w18, B629.13.w13, B631.22.w22)
    • Palpebral reflex. (B629.13.w13, B631.22.w22)
    • Toe pinch. (B629.13.w13, B631.22.w22)
    • Corneal reflex. (B631.22.w22)
    • Swallowing. (B631.22.w22)
    • Rectal tone. (B631.22.w22)
    • Changes in heart rate or respiratory rate in response to surgical stimulation also indicate anaesthetic depth. (B232.18.w18)
    • The degree of muscle relaxation should be monitored. (B631.22.w22)

Temperature

  • Hypothermia (Chilling - Hypothermia (with special reference to Waterfowl, Hedgehogs, Bears, Lagomorphs and Ferrets)) is probably the most common problem during ferret anaesthesia. (J15.24.w5, J29.7.w1)
  • Core body temperature should be monitored during surgery. (B629.13.w13, B631.22.w22, B631.23.w23, J29.14.w2)
    • An oesophageal or rectal probe can be used. (J29.7.w1)
  • During anaesthesia, external heat sources should be available. (B232.18.w18, B629.13.w13, B631.23.w23, J15.24.w5, J29.6.w3, J29.14.w2) Possible heat sources include:
    • A forced-air blanket system. (B629.13.w13, B631.22.w22, J29.14.w2)
    • Circulating warm water pad or blanket. (B629.13.w13, B631.22.w22, J29.7.w1)
    • Heated gel discs can be used. (B631.22.w22)
    • Hot water bottle ( e.g. warmed intravenous fluid bags, wrapped in towels). (B629.13.w13, )
    • An overhead heat lamp. (B629.13.w13, J29.6.w3)
  • Intravenous fluids and and fluids used for flushing (e.g. in the abdomen) should be warmed prior to use. (B629.13.w13, B631.22.w22, J29.6.w3, J29.7.w1, J29.14.w2)

Fluids

  • Monitor fluid losses during surgery (B629.13.w13) and maintain fluid balance, allowing for fluid/blood losses during surgery as well as maintenance requirements. (B232.18.w18)
  • Fluids should be warmed before administration. (B232.18.w18)
  • During short surgical procedures, 30 minutes or less, or if only minimal blood loss is expected, fluids can be given subcutaneously. (B631.22.w22)
  • It is recommended that an intravenous catheter should be placed when a ferret is to undergo surgery. (B629.13.w13, J29.14.w2) 
  • Intravenous fluid therapy should be used for longer procedures or when there is the possibility of significant blood loss. (B631.22.w22)
    • Give fluids as indicated based on monitoring. (B629.13.w13)
    • Usually, lactated Ringer's solution is given, at 10 mL/kg/hour. (J29.7.w1)
      • Saline, lactate Ringer's solution (Hartmann's solution) or dextrose salive can be given. (B232.18.w18)
    • Ferrets with Insulinoma (or with hypoglycaemia for any other reason) should be given fluids containing 2.5% or 5% dextrose during anaesthesia. (B629.13.w13, J29.7.w1)
  • For further information see section above on Fluid Therapy
Injectable general anaesthesia

Induction

  • Propofol (B631.22.w22, B339.9.w9, J29.7.w1, J513.7.w3)
    • 1.0 - 3.0 mg/kg intravenously via a cephalic vein catheter, after premedication with medetomidine 100 g/kg. After induction, intubate and maintain on isoflurane. (B339.9.w9)
    • 5 mg/kg intravenously, given to effect. (B631.22.w22)
    • 2 - 5 mg/kg intravenously. (J29.7.w1)
    • 4 - 6 mg/kg intravenously. (J513.7.w3)
    • Note: propofol commonly produces apnoea, particularly if administered rapidly. (J513.7.w3)
  • Etomidate 
    • 1 mg/kg intravenously, following premedication with Diazepam (0.5 mg/kg intravenously), for compromised ferrets. (J513.7.w3)
    • 1 mg/kg intravenously about 15 - 20 minutes after 0.25 - 0.3 mg/kg Midazolam for induction. This is usually sufficient for intubation, does not produce depression of cardiopulmonary parameters and is excellent for ill, critical animals. (B631.22.w22)
    • Cardiovascular and respiratory depression are minimal and the margin of safety is wide; this is useful for compromised ferrets. (J513.7.w3)
  • Ketamine is rarely used alone; it is generally given in combination with a benzodiazepam sedative or an alpha-2 antagonist.. (B631.22.w22)
    • Ketamine 10 - 20 mg/kg plus Diazepam 1 - 2 mg/kg intramuscularly provides light anaesthesia and only poor analgesia. (B631.22.w22)
      • Ketamine 25 mg/kg plus diazepam 2 mg/kg, intramuscularly. (B232.18.w18)
      • Note: Ketamine/diazepam results in paddling during recovery. (J29.7.w1)
    • Ketamine 5 - 8 mg/kg plus Medetomidine 0.08 - 0.1 mg/kg intramuscularly provides light anaesthesia with analgesia, hypotension and respiratory depression. Oxygen should be available. Reverse the medetomidine with Atipamezole at the end of the procedure. (B631.22.w22)
    • Ketamine 5 mg/kg plus Medetomidine 0.8 mg/kg plus Butorphanol 0.1 mg/kg intramuscularly. (J29.7.w1)
      • This allows intubation. (J29.7.w1)
      • Reversal of the medetomidine with Atipamezole produces a return to mobility within 10 minutes. (J29.7.w1)
    • Ketamine 5 - 10 mg/kg ten minutes after 0.25 mg/kg Midazolam provides heavy sedation/induction; inhalant anaesthetic is then required. (B631.22.w22)
      • Ketamine15 mg/kg plus Midazolam 0.4 mg/kg intramuscularly provides good sedation for intravenous catheterisation. (J29.7.w1)
    • Ketamine 25 mg/kg plus Acepromazine 0.25 mg/kg intramuscularly, to provide about 30 minutes of surgical anaesthesia. (B232.18.w18)
      • Note: Use of acepromazine should be avoided in ferrets, due to vasodilatation. (B631.22.w22)
    • Ketamine 10-20 mg/kg plus Xylazine 0.5 - 1.0 mg/kg subcutaneously or intramuscularly. (B631.22.w22) 
      • Ketamine 25 mg/kg plus xylazine 1-4 mg/kg intramuscularly. (B232.18.w18)
      • Respiratory depression is greater than with a medetomidine-ketamine combination. (B232.18.w18) 
      • Note: xylazine is not recommended for use in ferrets; it may produce hypotension, bradycardia and arrhythmias. (B631.22.w22) 
  • Thiopental can be used for induction, given intravenously at 8 - 12 mg/kg, to effect. (B631.22.w22, J29.7.w1)
  • Tiletamine-Zolazepam is rarely used in ferrets; recovery is prolonged with higher doses. (B631.22.w22)
    • The dose rate is 12-22 mg/kg intramuscularly. (B631.22.w22, J29.7.w1)
    • Analgesia is poor, although immobilisation is good. (J29.7.w1)
  • Fentanyl/Fluanisone, 0.3 mg/kg intramuscularly. (J29.7.w1)
Gaseous anaesthesia

Volatile agents can be used for induction and maintenance, or can be used for maintenance following induction using an injectable anaesthetic agent. (B232.18.w18, J15.24.w5)

Induction

  • Gaseous induction is generally preferred to injectable anaesthesia, especially in ill ferrets. (B629.13.w13) This method is used commonly in ferrets. (J29.7.w1)
  • Gaseous induction without premedication should be avoided, since this may result in anxiety and stress-related abnormalities such as increased heart rate, blood pressure etc. (B631.22.w22)
  • Induction may take place via mask or in an induction chamber until the ferret is sufficiently anaesthetised to allow endotracheal intubation. (B232.18.w18, B629.13.w13, B631.22.w22)
    • Mask induction can be carried out with the ferret restrained in a towel or held at the scruff and hips. (J29.7.w1)
    • Midazolam premedication should be given to reduce anxiety and associated cardiovascular changes, even if an induction chamber is used. (B631.22.w22)
  • An oxygen flow rate of 2 litres per minute and an isoflurane concentration of 4-5% should produce anaesthesia in 2-5 minutes. (B631.22.w22)
  • Without premedication, ferrets can be induced at 5% in 2 L/minute oxygen and will become anaesthetised in about two minutes. (J29.6.w3)
  • Isoflurane induction rate 5% (B629.13.w13) 3-4 % (B232.18.w18)
  • Sevoflurane induction rate 7%. (B629.13.w13)
  • Note: a ferret which has been anaesthetised with isoflurane previously may react to the small by salivating copiously. This ceases once the ferret is anaesthetised. (J29.6.w3)
  • Note: haematological parameters including rbc count, haemoglobin concentration, wbc count, PCV and plasma proteins decrease by 20-36% within 15 minutes of induction of isoflurane anaesthesia. (J13.55.w2,  J29.7.w1)
  • Note: Use of the same induction chamber for ferrets and for prey species such as rodents should be avoided, since the prey species will be distressed by the smell of the ferrets. (B232.18.w18, J15.24.w5)
  • A low-resistance circuit should be used, e.g. a T-piece. (B232.18.w18, J15.24.w5)

Endotracheal intubation

  • Except for very minor procedures, the ferret should be intubated. (B631.22.w2, J15.24.w52)
    • Intubation ensures a patent airway. (J29.14.w2)
    • Intubation generally is not difficult in ferrets. (B629.13.w13)
    • To intubate: With the ferret in sternal recumbency, have an assistant pull the ferret's head upwards by placing the forefinger and thumb in the corners of the ferrets mouth. Pull the ferret's tongue forward over the lower incisors, depressing the mandible. Advance the lubricated ET tube through the mouth and gently through the glottis. (B232.18.w18, J15.24.w5)
    • Use of a laryngoscope may assist in visualising the larynx, and facilitate intubation. (B232.18.w18, B629.13.w13, J15.24.w5, J29.13.w2)
      • A number one straight blade should be used. (J29.14.w2)
  • If laryngospasm occurs, Lidocaine (Lignocaine) may be applied to the larynx. (B629.13.w13, B631.22.w22)
    • 0.1 mL of Lidocaine (Lignocaine). (B631.22.w22) 0.05 mL of 2% lidocaine. (J29.13.w2)
    • One drop of 2% Lidocaine (Lignocaine) or a spray of topical cetacaine (Cetacain, Cetylite Industries Inc, Pennsauben, NJ, USA) can be used on the larynx to decrease spasm. (J29.14.w2)
  • Ferrets vary in size; 1.5 - 4.5 French endotracheal tubes should be available. (B631.22.w22)
    • 2 - 3.5 mm endotracheal tube. (B629.13.w13) 2.5 - 4 mm tube. (J15.24.w5)
    • A cuffed tube is recommended if available. (B631.22.w22)
    • For ferrets under 800 g bodyweight, an endotracheal tube  internal diameter 2.0 - 2.5 mm should be appropriate, but larger ferrets may need a 3.5 mm internal diameter uncuffed or 3.0 mm internal diameter cuffed tube. (J29.14.w2)
    • A 2.5 - 3.5 mm cuffed or uncuffed endotracheal tube may be used. (J29.13.w2)
  • Endotracheal tubes can be placed in the refrigerator to cool them so they become more rigid, or a stylet may be used. (B631.22.w22)
    • Pre-measure the stylet to fit the tube. (J29.14.w2)
  • Ferrets may be placed in sternal, dorsal or lateral recumbency for intubation. (J29.14.w2)
  • Once the endotracheal tube has been inserted, it can be secured by tying a length of gauze around the tube just outside the mouth, with the ends of the gauze then tied around the front leg on either side (if an intravenous catheter has been placed, tie the gauze distal to this). (B631.22.w22)
    • Alternatively, the gauze can be cross-tied under the chin then around the back of the head. (J29.7.w1)
  • Note: in a critically ill ferret, a 3.5 French red rubber catheter can be passed intranasally for oxygen supplementation. This should be premeasured. Prior to passing the catheter, instil a few drops of local anaesthetic solution into the nostril. Once the catahter is in place, suture to the skin of the head. (J29.13.w2)

Maintenance

  • For short procedures, gaseous anaesthesia can be provided via a face mask. (P120.2006.w7)
  • Once the ferret has been intubated, an oxygen flow rate of 0.6 - 1.0 L/min can be used, in a non-rebreathing circuit. (B631.22.w22)
    • Isoflurane 3% or Sevoflurane 5% in 1 L/minute oxygen, using a non-rebreathing semi-open circuit. (B629.13.w13)
    • Isoflurane can be used with nitrous oxide to provide additional analgesia. (J15.24.w5)
  • Isoflurane maintenance concentration is 1-3 %. (B631.22.w22); 1.5 - 3%. (B232.18.w18)
  • If apnoea occurs, Doxapram can be given 2-5 mg/kg intravenously or intramuscularly. (B631.22.w22)
  • Ventilate at 30-40 breaths per minute. (J29.14.w2)
  • If mechanical ventilation is used it should be set to give 40-70 breaths per minute, at the estimated lung volume. Note: the percentage of inhalant anaesthetic agent required is reduced with mechanical ventilation. (B631.22.w22)
Recovery
  • Discontinue the gaseous anaesthetic agent (isoflurane or sevoflurane) when the surgery has been completed. (B631.22.w22)
  • Maintain on oxygen until the ferret is breathing spontaneously (if mechanical ventilation has been used) with normal heart rate, respiratory rate and blood pressure. (B631.22.w22)
  • Note: alternative forms of analgesia (opiate, NSAID) should be provided before the vapouriser is turned off and the analgesia associated with the gaseous anaesthetic agent is removed. (B629.13.w13) See section above: Analgesia
  • As the ferret starts to move or gag, remove the endotracheal tube and disconnect the ferret from monitoring equipment such as ECG. (B631.22.w22)
  • Continue monitoring body temperature and reflexes until the ferret starts to move around. (B631.22.w22)
  • Continue providing supplemental heat as required until the ferret regains consciousness. (B232.18.w18)
  • Once recovered, ferrets generally curl up and sleep; ensure appropriate bedding is available. (B631.22.w22)
  • Specific reversal agents:
    • Atipamezole 0.4 - 1.0 mg/kg subcutaneously, intramuscularly, intraperitoneally or intravenously for the reversal of Medetomidine. (B631.22.w22)
    • Yohimbine 0.2 mg/kg intravenously or 0.5 mg/kg intramuscularly for the reversal of Xylazine. (B631.22.w22)
    • Naloxone 0.02 - 0.04 mg/kg subcutaneously, intramuscularly or intravenously, for the reversal of opiates. (B631.22.w22)
Local and Regional Anaesthesia 

Local anaesthetic agents can be useful as a ring block, or at major nerves as they exit from bony foraminae. (B631.22.w22) 

  • Lidocaine (Lignocaine) and Bupivacaine can be used in ferrets, either for local infiltration prior to incision of the surgical site, or for nerve blocks. (J513.7.w3)
    • Care must be taken not to use a toxic dose in this small species. (J29.14.w1)
    • Lidocaine (Lignocaine) 
      • 2 - 4 mg/kg local infiltration at the surgical site. (J513.7.w3)
      • Up to 2 mg/kg can be given subcutaneously. (J29.14.w1)
      • 1-2 mg/kg total given as a ring block or local infiltration. 1% or 2% solution can be used. The analgesia lasts about 15-30 minutes. (B631.22.w22)
    • Bupivacaine 
      • 1 -  3 mg/kg local infiltration at the surgical site. (J513.7.w3)
      • Less than 1.5 mg/kg subcutaneously. (J29.14.w1)
Dental nerve blocks
  • Dental nerve blocks should be carried out using a 25-gauge or 27-gauge needle. Infiltrate a few millimetres deep, lateral to the bone, in the area of the nerve. (B631.22.w22)
    • The technique should be practiced on a cadaver, so that the landmarks become familiar, using India ink as a marker. (B631.22.w22)
  • The infraorbital and zygomatic nerves, which provide sensory fibres to the maxillary incisors and canines, as well as the upper lip and adjacent tissues, exit the skull through the infraorbital foramen. This is on the lateral aspect of the face, just anterior to the zygomatic arch and rostral to the orbit, in the region of the second and third maxillary premolars. (B631.22.w22)
  • The mandibular nerve, which provides sensory fibres to the mandibular premolars and molars, and adjacent soft tissue, is found on the medial aspect of the mandible, about midway from the molar surface to the ventral surface of the mandible. Approaching from inside the mouth, the local anaesthetic is injected while "walking" the needle along the medial aspect of the mandible. (B631.22.w22)
  • The  mental nerve, which supplies sensory fibres to the lateral and ventral aspects of the mandible, together with the mandibular incisors and canines, and the lip, exits via the mental foramen on the lateral aspect of the mandible, about 2-4 mm rostral to the second or third mandibular premolar. (B631.22.w22)
  • The maxillary nerve, is located in the infraorbital canal; it is accessed from inside the mouth, but this may be difficult in small ferrets. It is imprtant to aspirate before injecting, to ensure the needle is not in a blood vessel. Local anaesthetic is infused slowly, while digital pressure is placed rostral to the end of the canal; if it is not possible to insert the needle into the infra orbital canal, then the local anaesthetic is infused at the rostral end of the canal, with firm digital pressure applied in the area and the injection taking place just caudal to the finger. (B631.22.w22)
  • Note: mild seizures have been observed in ferrets given bupivacaine at 1 mg/kg to block the innervation of the maxillary teeth or mandibular canine teeth. (B631.22.w22)
Epidural analgesia

This route provides analgesia without the systemic effects seen when analgesics are administered intravenously or intramuscularly. (J513.7.w3)

  • Lidocaine (Lignocaine)
    • Doses may be as high as 6 mg/kg. After injection, analgesia develops within 10-15 minutes and lasts for 60 - 90 minutes. (J513.7.w3)
    • 4.4 mg/kg epidurally. (B631.22.w22, J29.14.w1)
      • Lidocaine can be used together with epidural morphine, producing a more complete and immediate analgesia. (B631.22.w22)
  • Bupivacaine 
    • Can provide surgical analgesia for up to six hours. (J513.7.w3)
    • 1.1 mg/kg epidurally. Note: this may lead to hind limb motor weakness for up to 12 hours. (J29.14.w1)
  • Morphine 
    • 0.1 mg/kg once, as an epidural for surgical analgesia; the effects last 12-24 hours. (B631.22.w22)
    • Can be used with lidocaine (4.4 mg/kg, 2% formulation) or bupivacaine (1.1 mg/kg). (B631.22.w22)
  • Anaesthetise the ferret. (B631.22.w22)
  • Place the ferret in sternal recumbency with the lower back flexed. (B631.22.w22)
    • This opens the lumbosacral space. (B631.22.w22)
  • Identify the dorsal spines of the last lumbar vertebra (usually L6; sometimes L5 or L7) and S1, and the wings of the lieum. (B631.18.w18, B631.22.w22)
  • Clip and aseptically prepare the skin over the lumbosacral area. (B631.18.w18, B631.22.w22)
  • Slowly insert a 25-gauge needle almost perpendicularly, in the midline, between the vertebrae, and at the level of the wings of the ileum. (B631.18.w18, B631.22.w22)
    • Note: unlike in dogs and cats, it is unlikely that any "pop" will be noticed as the intervertebral ligaments are punctured. (B631.22.w22)
  • A small amount of fluid may be seen at the hub of the needle. (B631.22.w22)
    • One or two drops of CSF may be collected from this site. (B631.18.w18)
    • DO not aspirate fluid from this site, due to the risks of spinal cord damage. (B631.18.w18)
  • Inject the local anaesthetic and/or morphine. (B631.22.w22)
Anaesthetic Emergencies in Ferrets

As with all species, the basic "airway, breathing, circulation" protocols should be followed. (B602.33.w33, J29.14.w2)

Airway
  • Check that the mouth is clear. Extend the tongue, assess the colour of the mucous membranes, and assess breathing. (J29.14.w2)
Breathing
  • Stop any inhalation anaesthetic, ventilate with oxygen. (J29.14.w2)
  • Ventilate by:
    • Intubating. An uncuffed 2.0 mm internal diameter endotracheal tube can be placed in most ferrets. (J29.14.w2)
    • Place a small, tight-fitting mask over the ferret's muzzle and ventilate using oxygen. (J29.14.w2)
      •  It is possible to provide ventilation by blowing into the end of the mask, if no anaesthetic machine is available. (J29.14.w2)
    • If no other method is available, place your mouth over the ferret's muzzle and breath into it. (J29.14.w2) 
      • Afterwards, rinse your mouth with a disinfectant to minimise the risk of zoonotic disease transmission. (J29.14.w2)
  • Confirm that chest movement (i.e. lung ventilation) is occurring with ventilation. (J29.14.w2)
  • In respiratory arrest, administer a respiratory stimulant: Doxapram 5 - 10 mg/kg by intravenous, intratracheal or intraperitoneal injection. (J29.14.w2)
    • Repeat after 2-5 minutes if necessary. (J29.14.w2)
Circulation
  • In ferrets with cardiac arrest, perform external cardiac chest massage: remember that the heart in ferrets is quite far caudal in the chest, cup the heart with the thumb over the chest and the fingers underneath, and gently squeeze 15 times at a rate of 100 beats per minute or higher. (J29.14.w2)
  • If spontaneous cardiac contraction does not resume after 30-60 seconds of cardiac massage, administer 0.02 mg/kg adrenaline (epinephrine) via intracardiac, intravenous or intraosseous injection. (J29.14.w2)
    • Repeat after 2-5 minutes if necessary
  • If the heart is contracting but with bradycardia, administer 0.05 mg/kg atropine intravenously or 0.1 mg/kg intratracheally. (J29.14.w2)
Emergency drug administration
  • The intraperitoneal, intratracheal or intracardiac routes can be used for emergency administration of medication such as doxapram, diphenhydramine or adrenaline (epinephrine), if intravenous access is not available. (J29.14.w2)
  • Note: Response times are reduced if emergency drugs are pre-calculated and drawn into labelled syringes. (B602.33.w33)
Bonobo Consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
Anaesthesia of great apes
  • The large size, physical strength, agility and intelligence of the great apes can make anaesthesia challenging. (B538.33.w33)
  • Changes to their normal environment can make great apes excited or aggressive, which in turn can make induction of anaesthesia distressing. Additionally, disease conditions such as cardiomyopathy (Myocardial Fibrosis in Great Apes) can complicate anaesthesia. (B538.33.w33)
  • Human anaesthetic equipment (face masks etc.) often is used for great ape anaesthesia, due to the close anatomical similarities. (B538.33.w33)
  • A team of experienced personnel is very important for great ape anaesthesia. (B538.33.w33)
  • All personnel coming into close proximity with the great ape should wear appropriate personal protective equipment (e.g. face mask, gloves) to minimise risks of zoonotic disease transfer.
  • Protocols for escape and for bite wounds to humans preferably should be in place before the procedure. A first aid kit suitable for dealing with bite wounds should be available. (B538.33.w33)
  • Note: A survey of great ape peri-anaesthetic deaths in great apes showed that sick and elderly (over 30 years of age) are at much greater risk of peri-anaesthetic mortality. (J290.34.w1)
Pre-anaesthetic preparation
  • Prior to anaesthesia, 24 hours of fasting (including no water) is recommended for great apes. (B336.39.w39) 12-24 hours of fasting. (B538.33.w33, D409.6.w6)
    • Note that some individuals will eat bedding such as straw, if fasted and others will eat their own faeces (coprophagy). (B336.39.w39)
    • The fasting period may indicate to the ape that an anaesthetic procedure is imminent, with resultant agitation and aggression. Keeping the routine as close as possible to normal may minimise this distress. (B538.33.w33)
  • For elective anaesthesia it is preferable that any air sac infection (Laryngeal Air Sacculitis in Bonobos) is treated beforehand, to reduce the risk of pneumonia following from aspiration of purulent material from the infected air sacs. (B538.33.w33)
  • Individuals with a history of cardiac disease, and older individuals, should be carefully evaluated prior to anaesthesia. (B538.33.w33)
Premedication
  • Diazepam has been given orally to reduce anxiety/provide some sedation prior to anaesthesia. (B538.33.w33, P30.1.w12)
    • Diazepam 0.2 mg/kg was given orally about 90 -120 minutes before anaesthesia to reduce anxiety. (P30.1.w12)
    • Diazepam was given at 5 mg total dose orally in juvenile gorillas (four years old) before ketamine anaesthesia. (P504.2001.w7)
  • Metoclopramide (0.4 mg/kg, orally) has been given 90-120 minutes before oral administration of anaesthetic agents, to prevent vomiting. (P30.1.w12)
Injectable general anaesthesia
  • Ideally, great apes are trained to present a large muscle mass for injection. (B336.39.w39, B538.33.w33, D409.6.w6)
    • This is the safest and least stressful option. (D409.6.w6)
  • If necessary, the anaesthetic agent(s) can be delivered via a lightweight dart using a blowpipe or dart gun. (B336.39.w39, B538.33.w33)
    • Use the least traumatic delivery system available. (B538.33.w33)
    • Every effort should be made to dart the ape without it becoming aware that this is to occur, since once aware the animal (and others in the group) will be agitated and mobile, and it is much harder to hit the animal with the dart. (B538.33.w33)
    • It can be particularly difficult to dart one individual in a social group, and separation prior to darting may be impossible or greatly increase the stress of the animal. It is essential, particularly in the event of an anaesthetic emergency,  to have a mechanism by which the anaesthetised individual can be isolated and retrieved. (B538.33.w33)
    • If possible it may be useful to remove furnishings to prevent the ape from hiding behind these to avoid being darted. (B538.33.w33)
    • Take care not to hit the sexual swelling of females as this is very vascular and friable, and severe haemorrhage could result. (B538.33.w33)
    • Note that apes can grab at loose clothing etc. and at the dart gun, from longer distances than might be expected. (B538.33.w33)
    • Preferably use potent drugs in a small volume to assist with full injection of the anaesthetic dose before the age pulls out the dart, which can happen very quickly.
    • Note: chimpanzees are known to throw materials, including faeces, to spit water, and to return darts to the darter with some force. (B538.33.w33)
  • Injectable agents used in great apes include (B336.39.w39)
    • Ketamine, 6.0 - 8.0 mg/kg intramuscularly or intravenously. (B336.39.w39)
      • Note: doses of up to 15 mg/kg have been reported. (B336.39.w39)
      • In Pan troglodytes - Chimpanzee, doses of 2-5 mg/kg or 8-10 mg/kg are used by AZA-accredited institutions. (D409.6.w6)
      • Has been used alone in chimpanzees, gorillas and orang utans. (B538.33.w33)
      • Reported to produce smooth, rapid induction, good analgesia, adequatemuscle relaxation and minimal cardiopulmonary effects. (B538.33.w33)
      • Recovery occurs 40-60 minutes after injection and is calm, with minimal ataxia and anxiety. (B538.33.w33)
      • Ketamine is not reversible, the duration of action is short, sudden unexpected recoveries can occur and, with repeated use, tolerance develops. (B538.33.w33)
      • Usually used in combination with other chemical restraint agents. (B336.39.w39)
      • Laryngospasm and hypersalivation are known problems with ketamine anaesthesia in great apes. (B538.33.w33)
    • Tiletamine/zolazepam, 4.0 - 6.0 mg/kg intramuscularly. (B336.39.w39)
      • Induction is rapid, but recovery can be rough. (B336.39.w39)
      • Recovery time appears to be dose-dependent; low loses are suggested for short procedures. (B538.33.w33)
      • In Pan troglodytes - Chimpanzee, doses of 1.5-3.0 mg/kg and of 3-6 mg/kg have been used by AZA-accredited institutions. (D409.6.w6)
    • Diazepam, 0.5 - 1.0 mg/kg. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • Can be reversed with flumazenil, 0.02 - 0.1 mg/kg intravenously. (B336.39.w39)
    • Midazolam, 0.05 - 0.15 mg/kg intramuscularly or intravenously. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • If given intravenously, transient bradycardia may occur. (B538.33.w33)
      • Can be reversed with flumazenil, 0.02 - 0.1 mg/kg intravenously. (B336.39.w39)
    • Butorphanol, 0.1 - 0.2 mg/kg intramuscularly or intravenously. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • Can be reversed with Naloxone, 0.02 mg/kg intramuscularly or intravenously. (B336.39.w39)
    • Buprenorphine, 0.01 - 0.02 mg/kg intramuscularly or intravenously. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • Can be reversed with Naloxone, 0.02 mg/kg intramuscularly or intravenously. (B336.39.w39)
    • Xylazine, 0.5 - 2.0 mg/kg intramuscularly. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • Can be reversed with atipamezole 0.25 - 0.5 mg/kg mg/kg intramuscularly or intravenously. (B336.39.w39)
    • Medetomidine, 0.02 - 0.05 mg/kg intramuscularly. (B336.39.w39)
      • Used in combination with other chemical restraint agents. (B336.39.w39)
      • Can be reversed with atipamezole 0.1 - 0.25 mg/kg mg/kg intramuscularly or intravenously. (B336.39.w39)
      • Note: can allow a reduction in ketamine dose as much as five-fold. (B336.39.w39)
    • Ketamine 10-20 mg/kg plus xylzine 1 mg/kg (chimpanzee); ketamine 5-7 mg/kg plus xylazine 1.0 - 1.4 mg/kg (orang utan)
      • Induction time is similar to ketamine alone, but the depth of anaesthesia is greater, with better muscle relaxation, analgesia and cardiopulmonary stability. (B538.33.w33)
      • Recovery uneventful. (B538.33.w33)
    • Ketamine 1.0 mg/kg plus xylazine 0.25 mg/kg plus tilatamine-zolazepam 1.25 mg/kg (B336.39.w39)
      • Provide oxygen. (B336.39.w39)
      • Monitor closely. (B336.39.w39)
      • Xylazine component can be reversed with atipamezole or yohimbine. (B336.39.w39)
    • Ketamine 2.0 mg/kg plus medetomidine 0.03 - 0.04 mg/kg. (B336.39.w39)
      • An adult female bonobo was anaesthetised using 100 mg ketamine and 1 mg medetomidine. (J543.38.w3)
      • Note: medetomidine can allow a five-fold reduction in ketamine dose. (B336.39.w39, B538.33.w33)
      • Rapid, safe induction within 3 - 15 minutes. (B538.33.w33)
      • Note: it is important to leave the animal undisturbed for the firat 10 minutes after immobilisation. Trying to move or manipulate the animal earlier than this can result in rapid arousal. (B538.33.w33)
      • Cardiovascular effects are minimal; there is a modest rise in blood pressure soon after induction. (B538.33.w33)
      • Medetomidine can be reversed with atipamezole. (B336.39.w39) at five times the dose of medetomidine (mg/kg). (B538.33.w33)
        • This produces a smooth recovery, complete within six minutes (intravenous) or 10-13 minutes (intramuscular). (B538.33.w33)
      • Note: even without use of a reversal agent, sudden recoveries can occur, producing a dangerous situaton. (B538.33.w33)
      • In Pan troglodytes - Chimpanzee, doses which have been used by AZA-accredited institutions include: (D409.6.w6)
        • Ketamine 2 mg/kg plus medetomidine 0.015-0.025 mg/kg. (D409.6.w6)
        • Ketamine 2-5 mg/kg plus medetomidine 0.03-0.04 mg/kg. (D409.6.w6)
        • Ketamine 5-7 mg/kg plus medetomidine 0.025-0.07 mg/kg. (D409.6.w6)
    • Ketamine 2.0 - 3.0 mg/kg plus medetomidine 0.02 - 0.04 mg/kg plus butorphanol 0.2 - 0.4 mg/kg (B336.39.w39)
      • Provide oxygen. (B336.39.w39)
      • Monitor closely. (B336.39.w39)
      • Medetomidine can be reversed with atipamezole. (B336.39.w39)
      • Butorphanol can be reversed with naloxone. (B336.39.w39)
      • In Pan troglodytes - Chimpanzee, ketamine 1.5 mg/kg plus medetomidine 0.015 mg/kg plus butorphanol 0.15-0.3 mg/kg. (D409.6.w6)
    • Ketamine 3.0 mg/kg plus butorphanol 0.4 mg/kg plus midazolam 0.3 mg/kg (B336.39.w39)
      • Provide oxygen. (B336.39.w39)
      • Monitor closely. (B336.39.w39)
      • Butorphanol can be reversed with naloxone. (B336.39.w39)
      • Midazolam can be reversed with flumazenil. (B336.39.w39)
    • Ketamine plus midazolam
    • Tiletamine-zolazepam
      • 1.25 mg/kg plus medetomidine 0.03 - 0.04 mg/kg (B336.39.w39)
      • Medetomidine can be reversed with atipamezole. (B336.39.w39)
      • Induction smooth and rapid (1-7 minutes in Pan troglodytes - Chimpanzee and Pongo pygmaeus - Orang-utan) with stable cardiovascular parents. Lower blood pressure in chimpanzees anaesthetised using this combination and maintained on isoflurane compared with those anaesthetised with ketamine/medetomidine and maintained with isoflurane. (B538.33.w33)
      • Recovery was prolonged in Pan troglodytes - Chimpanzee, and use of flumazenil only transiently increased alertness. (B538.33.w33)
      • In Pan troglodytes - Chimpanzee, doses of 2 mg/kg tiletamine-zolazepam plus 0.02-0.03 mg/kg medetomidine have been used by AZA-accredited institutions. (D409.6.w6)
    • Tiletamine/zolazepam plus ketamine
    • Medetomidine plus butorphanol plus midazolam
      • Medetomidine 0.015 mg/kg plus butorphanol 0.085 mg/kg plus midazolam 0.06 mg/kg. (D409.6.w6)
    • ADDITIONAL DOSES of injectable anaesthetics which have been used for maintenance of anaesthesia  in Pan troglodytes - Chimpanzee in AZAZ-accredited zoos include: (D409.6.w6)
      • Tiletamine-zolazepam 1-2 mg/kg. (D409.6.w6)
      • Ketamine 1-2 mg/kg or less commonly 3-4 mg/kg. (D409.6.w6)
      • Propofol 1-2 mg/kg. (D409.6.w6)
      • Diazepam 0.1 mg/kg. (D409.6.w6)
      • Midazolam 0.1 mg/kg. (D409.6.w6)
Oral administration of anaesthetic drugs
  • This can be a useful alternative to darting; it appears to be less stressful to the individual being anaesthetised and to the rest of the social group. (P30.1.w12)
  • In gorillas, ketamine 9.4 mg/kg plus detomidine 0.3 mg/kg resulted in lateral recumbency in 17 minutes; lesser effects occurred in individuals with partial dosing; in all cases, administration of additional anaesthetic drugs (by darting) was possible with only mild to moderate responses and absence of screaming or charging. (P30.1.w12)
  • There is a potential risk of aspiration associated with the consumption of food or drink immediately before anaesthesia, although this has not been a problem in practice. (B538.33.w33)
    • One of six gorillas given oral ketamine plus detomidine showed regurgitation. (P30.1.w12)
  • Metoclopramide (0.4 mg/kg, orally) was given 90-120 minutes before the anaesthetic agents, to prevent vomiting. (P30.1.w12)
  • Diazepam 0.2 mg/kg was given orally about 90 -120 minutes before anaesthesia to reduce anxiety. (P30.1.w12)
  • Note: Results can be variable; in one trial using Tiletamine-Zolazepam (General anaesthetic) (Chemical Page), two Pan troglodytes - Chimpanzees showed effects within 30 minutes, while two others showed no signs of sedation until they were fed, five hours later, then developed a surgical plane of anaesthesia. (B16.1.w1)
Intubation
  • Intubation should be carried out for all except the shortest, most minor anaesthetic procedures. (B336.39.w39)
  • Intubation should not be attempted until an adequate depth of anaesthesia has been obtained., or laryngospasm is likely to occur. (B336.39.w39)
    • An inhalant anaesthetic can be delivered by face mask if required initially to obtain the correct depth of anaesthesia. (B336.39.w39)
    • Laryngospasm is a particular problem with ketamine anaesthesia. (B538.33.w33)
  • To reduce laryngospasm, lignocaine can be applied to the glottis; this should be carried out several minutes before intubation is attempted. (B538.33.w33)
  • Intubation of great apes is not difficult; it is most easily achieved with the ape in dorsal recumbency, the head being extended over the edge of the table. (B336.39.w39, B538.33.w33)
  • The tongue can be pulled forwards to make intubation easier. (B538.33.w33)
  • Intubation is easier if a Macintosh laryngoscope blade is used to push the tongue down. (B336.39.w39, B538.33.w33)
  • A cuffed endotracheal tube should be used. (B336.39.w39)
    • Cuffed Murphy tubes of 6-12 mm diameter are used for great apes. (B538.33.w33)
  • It is important to use a cuffed endotracheal tube when anaesthetising a great ape with air sacculitis (Laryngeal Air Sacculitis in Bonobos). (B538.33.w33)
  • Care should be taken not to introduce the tube too deeply as the trachea is too short and there is a risk of the tube extending into one of the primary bronchi [therefore only that side is ventilated]. (B336.39.w39, B538.33.w33)
    • After placing the endotracheal tube, auscultate both sides of the chest and confirm air sounds on both sides. (B538.33.w33)
Monitoring
  • Careful, close monitoring is recommended throughout great ape anaesthetics. (B538.33.w33, D409.6.w6)
  • Both manual monitoring and instrumental monitoring should be used throughout the anaesthetic. (B336.39.w39) 
  • Particular care should be taken to closely monitor older individuals and those with a history of cardiac disease. (B538.33.w33)

Monitoring should include:

  • Core temperature. (B336.39.w39, D409.6.w6)
  • Cardiac rate and rhythm. (B336.39.w39, D409.6.w6)
    • Heart rates of 60-200 bpm have been measure in anaesthetised Pan troglodytes - Chimpanzees. (B538.33.w33)
    • ECG should be used if possible. (B336.39.w39) This is important as it shows cardiac electrical activity. (B538.33.w33)
      • This should be used particularly in longer procedures. (D409.6.w6)
  • Pulse quality. (D409.6.w6)
  • Capillary refill time. (D409.6.w6)
  • Ventilation (B336.39.w39)
    • Respiratory rate and depth should be monitored as a minimum. (D409.6.w6)
    • Capnography is suggested to measure end-tidal CO2 (B336.39.w39, B538.33.w33)
    • Measurement of arterial blood gases can be used. (B336.39.w39)
    • Respiratory rates of 20-60 breaths per minute have been measured in anasethetised Pan troglodytes - Chimpanzees. (B538.33.w33)
  • Oxygenation (oxygen haemolobin saturation). (B336.39.w39)
    • Pulse oximetry is recommended. (B336.39.w39, B538.33.w33, D409.6.w6) particularly for longer procedures. ()
    • Measurement of arterial blood gases can be use. (B336.39.w39, D409.6.w6) and is recommended for longer procedures. (D409.6.w6)
  • Blood pressure, measured directly or indirectly. (B336.39.w39)

(B336.39.w39)

Inhalant anaesthesia
  • Usually, gaseous anaesthetic agents are used for maintenance of anaesthesia. These should normally be delivered via an endotracheal tube, although delivery via  a tightly-fitting face mask may be needed until the depth of anaesthesia is sufficient to allow intubation. (B538.33.w33)
  • Isoflurane is commonly used. (B538.33.w33)
    • Vasodilatation can occur with resultant sevre hypotension. (B538.33.w33)
    • The MAC of isoflurane should be reduced as appropriate depending on the induction agent(s) and dose given. (B538.33.w33)
  • Sevoflurane can be used. (D409.6.w6)
Reversal
  • It is important to remember that reversal can occur quite suddenly after use of reversal agents. (B538.33.w33)
  • The great ape should be in a secure location before the reversal agent is given. (B538.33.w33)
  • Great apes should be allowed to recover from imobilisation in isolation, to prevent any attacks by other members of their social group. (B538.33.w33)
  • Place the anaesthetised individual in the recovery position to minimise the risks of aspiration which may occur with regurgitation, vomiting or heavy salivation. (D409.6.w6)
    • Hay or blankets can be used to assist maintence of this position, with the head slightly elevated but the mouth directed downwards. (D409.6.w6)
  • Note: A survey of great ape peri-anasthetic deaths in great apes showed that the recovery period and the first 24 hours post-anaesthetic were the greatest risk periods. (J290.34.w1)
Associated techniques linked from Wildpro Waterfowl

Cranes

Bears

Lagomorphs

Ferrets

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Surgery

GENERAL PRINCIPLES
  • Surgery should not be carried out until the patient has been stabilized. Life-threatening problems should be addressed immediately.
  • Peri-operative pain management is always important.
  • Consideration of the size of the patient is important when choosing appropriate sizes of instruments and of materials such as suture materials.
    • It is also important to consider the abilities of the patient to remove sutures post-operatively.
  • In small patients, minimising blood loss, and accurate calculation of the volume lost is extremely important, as small absolute losses may represent a large percentage loss of circulating blood volume.
  • The use of perioperative antibiotics, with parenteral antibiotics administered from 1-2 hours pre-operatively and a therapeutic level maintained for 8-16 hours post-operatively should be considered; post-operative antibiosis should be continued for contaminated wounds, or if the surgical field was contaminated intra-operatively.

(V.w5, V.w6)

FOR BIRDS
GENERAL PRINCIPLES
  • Surgery should not be instigated prior to stabilization of the patient. Life-threatening problems should be addressed immediately.
  • Birds have skin which is thin relative to the skin of mammals and is also relatively inelastic.
  • The dermis is attached to the underlying muscle fascia with little subcutaneous tissue.
  • In feathered areas the skin of the patria (areas between the feather tracts) is stronger than that of the puerile (feather tracts)
  • Birds have light bones, many being pneumotised.
  • Birds heal relatively quickly.
  • In treating wing injuries, the subsequent ability of the bird to fly is of primary concern. This is particularly important for birds which are to be returned to the wild and which rely on their flying ability.
  • The loss of even a few drops of blood from a small bird may represent the loss of a significant proportion of its total blood volume.
  • Birds rely to a great extent on their feathers for insulation. Removal of feathers in preparing a surgical site and the use of antiseptic solutions in surgical site preparation may both result in considerable heat loss and the risk of hypothermia. This is particularly important in small birds. Minimizing removal of feathers is also important for water birds which rely on their feathers for buoyancy and waterproofing.
  • The surgical site may be cleared by plucking small feathers. Removal of the large flight feathers (primaries and secondary) should be avoided if possible: these are attached to the eriostemon of the underlying bones. If plucking of these feathers is essential, each feather should be removed individually, with the surrounding skin held firmly with one hand while the feather is grasped at its base (artery forceps may be used to grip the feather firmly) and pulled in the direction of feather growth.
  • Where possible, feathers may be kept from the surgical site using e.g. masking tape, or (particularly for flight feathers) covered with self-adhesive bandage such as Vetrap, or with a sterile stockinette.
  • Plucked feathers will usually re-grow within a few weeks, whereas cut feathers will not be replaced until the next moult (B14).
  • Solutions which may be used for cleaning and sterilization of the surgical site include quaternary ammonium solutions, chlorhexidine diacetate (0.05%), chlorhexidine gluconate (4%) rinsed with saline or alcohol, benzalkonium chloride and povidone iodine (1%).
  • Saline rather than alcohol rinsing is preferred as alcohol results in greater heat loss.
  • Electrical or water-circulated heat pads or hot water bottles, suitably padded to avoid burns, may be used to reduce heat loss from the bird during surgery.
  • Clear plastic sterile drapes are particularly useful to allow visual monitoring of the bird during the operation.
  • The use of perioperative antibiotics, with parenteral antibiotics administered from 1-2 hours pre-operatively and a therapeutic level maintained for 8-16 hours post-operatively should be considered.
  • Post-operative antibiosis should be continued for contaminated wounds, or if the surgical field was contaminated intra-operatively.
  • Consideration of the size of the patient is important when choosing appropriate sizes of instruments and materials such as suture materials.

(J2.23.w2, B13.40.w13, B13.41.w14, B14)

FRACTURE MANAGEMENT
  • Well-aligned stable avian fractures heal rapidly, often within three weeks. Healing may be delayed by fracture instability, infection, metal objects placed in the fracture site which impair the formation of endosteal callus. The method chosen for fracture repair will depend on a variety of factors including: type of fracture, bone involved, age and size of bird and the required degree of post-operative function. N.B. avian bone tends to crack and shatter more readily than mammalian bone, due to the higher calcium content. Fractures are therefore more often comminuted. Some fractures such as simple wing fractures and some lower leg fractures may heal well with simple bandaging. In general, results are better using techniques allowing immediate weight-bearing and normal joint function, such as external fixation and intramedullary pinning.
  • Initial examination should include haemostasis, shock therapy and temporary support (bandaging and splinting) for any fractures, and be as atraumatic as possible, with minimal handling of the fracture area. Haemorrhage at the wound site should be controlled by a pressure bandage or by vessel ligation. Antibiotics or corticosteroids or both can be administered at this time, if indicated. Birds with fractures that are several days old may be hypoglycaemic and should receive intravenous or oral glucose solutions to meet their immediate metabolic needs (1 to 2ml of 20% dextrose per kg). After initial examination and treatment for life-threatening problems the patient should be placed in a warm, darkened environment for several hours to allow stabilization before a more detailed examination to assess the fracture is performed. Surgery to repair the fracture may be delayed for several days until the patient stabilizes.
  • Wounds associated with open fractures should be cleaned and necrotic soft tissue and bone should be debrided: osteomyelitis is probably the most significant threat to fracture healing and can be devastating. Aseptic preparation is essential prior to surgery. Caseous material must be debrided from chronic infected fractures prior to any attempt to stabilize the fracture, and tissues must be handled gently to avoid damaging blood supply and promoting the development of adhesions, osteomyelitis or non-union.

(J2.23.w2, B10.20.w16, B11.36.w4)

Waterfowl Consideration

  • In waterfowl it is particularly important to minimize the removal of feathers, as waterfowl rely on an intact layer of contour feathers (body feathers) for maintenance of buoyancy and waterproofing as well as insulation.
  • Restoration of flying ability is often less vital than for many other birds. Waterfowl in a sheltered situation (e.g. in a collection or on a park lake with an island available for roosting) may be able to cope very well with reduced flight capability or even the loss of a wing. However the loss of a leg is less likely to be tolerated well, particularly in larger, heavier species.
  • Provision of water for swimming is important for convalescent waterfowl and contamination of surgical incisions is a considerable risk with dirty water. Sealing skin wounds with e.g. OpSite Spray (Smith and Nephew) may be used to reduce the risk of infection.

(B11.23.12, B11.36.w4, V.w5).

Crane Consideration
Fracture management
Ventricular foreign body removal
Endoscopic examination
Bear Consideration Surgery can be carried out using the same general principles as for dogs, with allowance for the larger size of adult bears.
Fracture management
  • Internal fixation is preferred, generally using compression-plating techniques, particularly for complicated fractures and in larger bears. (B16.9.w9, B64.26.w5)
  • Note: The temperament of the individual bear will affect healing. (J428.34.w1)
  • Challenges to bear fracture treatment include:
    • External coaption devices may be destroyed by the bear. (J428.34.w1)
    • The bear may be stressed by confinement and isolation. (J428.34.w1)
    • Administering medication such as post-operative antibiotics. (J428.34.w1)
    • Repeated general anaesthesia is required for cast monitoring, cast changes and radiographic monitoring of healing. (J428.34.w1)
    • Social reintegration when the bear is returned to its enclosure. (J428.34.w1)

Lagomorph Consideration

General surgical information
  • Good illumination is required.
  • Compared to cats and dogs, rabbits have thin, delicate, friable tissues.
  • Fine surgical instruments are required and kits designed for rabbit surgery have been developed. (B600.15.w15)
  • An optical loupe or operating microscope may be useful.
  • Aseptic technique is important to prevent subclinical or clinical wound infections. This includes standard preparation of the surgeon and surgical site, use of gowns, masks and caps, and sterile drapes, gloves and instruments. (B615.8.w8)
  • Transparent plastic drapes are useful, making it easier for the anaesthetist to monitor the patient. (B600.15.w15)
  • Take care to avoid injuring the caecum when entering the abdomen. (B615.8.w8)
  • If the abdominal or thoracic contents are exposed for any length of time, steps must be taken to prevent development of hypothermia and dehydration. (J34.17.w1)
    • Heat may be provided by circulating hot water blankets, hot water bottles or heat lamps. (J34.17.w1)
    • During abdominal surgery, periodic irrigation of the abdominal cavity with warm sterile isotonic solution may be helpful. (J34.17.w1)
  • Whenever possible, manipulate organs via adjacent adipose tissue rather than handling the tissue directly. This can reduce the production of microhaemorrhages and therefore of possible adhesion sites. (B534.43.w43f)
  • Post-operative pain may be reduced by careful technique and gentle tissue handling. (J213.4.w5)
Minimising adhesions
  • Development of adhesions after surgery is common in rabbits. 
    • Minimise tissue handling and ensure gentle technique to minimise adhesions.
    • Adhesions may be induced by foreign material such as lint from gauze swabs or talc from gloves.
      • Once the surgeon is gowned and gloved, powder and talk should be removed from the gloves using a sterile, saline-soaked gauze swab.
      • Swabs with frayed ends should not be used in rabbits in case they become separated, remain in the abdomen and act as a focus for formation of adhesions. (B534.43.w43f)
    • While it is generally recommended that viscera be omentalised, this may not be possible because rabbits have a small omentum. (B600.15.w15)
    • Development of adhesions may be prevented by use of calcium channel blockers, e.g. verapamil, 200 micrograms per kg subcutaneously, every eight hours for nine doses. This is useful following e.g. large intestinal surgery, ruptured pyometra or removal of abdominal abscesses. (B600.15.w15, B602.22.w22)
  • Rabbits readily develop fat necrosis, especially in the broad ligament. Breakdown of fat into fatty acids and glycerol, which combine with ions (sodium, potassium, calcium) is associated with trauma. (B600.15.w15)
Pre-operative preparation
General
  • Pre-operative fasting is not required. (B602.22.w22, B534.43.w43f)
  • Prior to surgical treatment in rabbits with either localised (e.g. abscess) or generalised bacterial infection (e.g. "snuffles" due to Pasteurellosis in Lagomorphs), start antibacterial therapy. (B602.22.w22)
    • Also give prophylactic antibiotics if there is a significant risk of bacterial contamination during surgery. (B602.22.w22)
    • Prophylactic antibiotics may be given from the day before surgery (if elective) or at the time of anaesthetic induction, and continued for three days. (B534.43.w43f)
  • If vascular access or supportive fluid therapy will be needed, place a 20 - to 26-gauge catheter into the cephalic vein or lateral saphenous vein. (B602.22.w22) or the marginal ear vein (B534.43.w43f) See: Intravenous Injection and Catheterisation of Rabbits
    • Slow administration of crystalline fluids during the operation will maintain patent intravenous access. (B534.43.w43f)
  • A complete pre-surgical workup, including history, physical examination, complete blood count and urinalysis, should be carried out to assess the rabbit's general health status before surgery. (B534.43.w43f)
  • Pre-operative analgesia may reduce the degree of pain post-operatively (B601.16.w16, J15.20.w2) and the risk of post-operative gastrointestinal stasis. (J213.5.w3)
  • Preferably correct dehydration before starting surgery. (P113.2005.w3)
Clipping
  • Place sterile lubricant in the wound before clipping the fur to avoid further contamination. (J213.7.w2)
  • Gently shave the fur, taking care to avoiding damage to the skin. (J213.7.w2)
    • Rabbits have thin, delicate skin which is easily damaged. (B600.15.w15, B602.22.w22, J34.17.w1, J213.7.w2)
    • The fine, dense fur easily clogs clipper blades. (B600.15.w15)
    • Use good-quality, robust clippers, and clip slowly to prevent fur catching. (B600.15.w15, B602.22.w22)
    • Keep the skin spread flat in front of the clippers. (B602.22.w22)
    • Note: Rabbits may show postoperative irritation, pain and even self-mutilation due to iatrogenic damage to the skin from clippers.
    • Vacuum off loose hair from the site. (B534.43.w43f)
  • Depilatory creams can be used but are messy and difficult to clean off properly. (B600.15.w15)
  • Note: fur may grow back in a patchy manner, due to variations in rabbit hair growth cycles. (B534.43.w43f)
  • Avoid excess clipping, as this may predispose to hypothermia. (J15.23.w6)
Cleansing
  • Adequate cleansing of the skin is important. (J34.17.w1)
  • A single application of chlorhexidine in spirit can be used and does not require scrubbing of the skin. (B600.15.w15)
  • Avoid using large quantities of spirit as this may cause excessive heat loss. (B600.15.w15)
  • Three cycles of povidone-iodine (iodophors) soap and alcohol or sterile saline has been suggested, working out from the surgical site, and followed by spraying on povidone-iodine solution and allowing it to dry. (B534.43.w43f)
  • Avoid excessive scrubbing of the skin as this may lead to postoperative irritation, pain and even self-mutilation by the rabbit, (B600.15.w15)
Suture materials
  • Suture materials removed by hydrolytic degradation are preferable in rabbits to reduce development of abscesses around suture materials. (B602.22.w22)
  • Exuberant granulation tissue as well as fistulous tracts may develop around braided materials. monofolament absorbable suture materials such as polydioxanone are preferred for internal sutures, and monofilament non-absorbable for skin sutures - nylon is appropriate, also skin staples. (J213.5.w3)
  • Use of fine suture materials is highly recommended to reduce tissue reaction leading to formation of adhesions. (B600.15.w15, B602.22.w22)
    • Use swaged-on 3/0, 4/0 or 5/0 suture materials. (B600.15.w15)
    • Generally, use polydioxanone (PDS II) or poliglecaprone (Monocryl, Ethicon). (B600.15.w15)
      • These break down by enzymatic hydrolysis and promote minimal adverse tissue reactions in rabbits. (B534.43.w43f)
    • 3/0 or 4/0 catgut can be used for tying off blood vessels and ligaments. (B600.15.w15)
  • For closing skin incisions which are not under tension, absorbable suture materials are suitable, for example Polyglactin 910 (Vicryl Rapide, Ethicon). Rabbits have only a mild inflammatory response to polyglactic acid. (B600.15.w15)
    • Skin staples are well tolerated by rabbits and are reliable. (B602.22.w22)
    • Tissue glue can be used; occasionally a rabbit will remove this. (B602.22.w22)
  • If a contaminated wound needs to be closed, e.g. following placement of antibiotic impregnated PMMA beads, use a fine monofilament suture material and small knots to minimise the risk of secondary abscess formation. Appropriate suture materials include polydioxanone (PDS, Ethicon), or poliglecaprone (Monocryl, Ethicon). (B600.15.w15)
    • Alternatively, an absorbable material such as catgut can be used which will be removed, allong with any associated bacteria, by macrophages. (B600.15.w15)
  • For repair of hollow abdominal organs use fine absorbable monofilament e.g. polydioxanone (PDS, Ethicon), or poliglecaprone (Monocryl, Ethicon). (B600.15.w15)
    • Poliglecaprone (Monocryl, Ethicon) is good to handle and knot, and causes only minimal inflammatory reaction. (B600.15.w15)
    • Monofilament polyglyconate (Maxon, Davis & Geck, Manati, PR) can be used. (B602.22.w22)
    • Note: catgut is not suitable for closure of the stomach, due to the acidic environment. (B600.15.w15)
  • Stainless steel or tantalum clips (Hemoclips, Weck, Research Triangle Park, NC) can be used for ligation of vessels and small pedicles, and result in minimal formation of adhesions. (B602.22.w22)
Suture patterns

Abdominal incision (B600.15.w15)

  • Following a midline linea alba approach, 4/0 polydioxanone (high tensile strength, degrades slowly) or 4/0 poliglecaprone (Monocryl, Ethicon). (B600.15.w15)
  • Repair the abdominal fascia in a single layer using either:
    • A row of simple interrupted sutures. OR
    • A continuous suture, with an extra four throws at the start and six throws at the end. The first throws need to draw the edges of the fascia together without crushing the tissue. (B600.15.w15)
  • Close the skin with:
    • Continuous subcuticular suture with a buried Aberdeen knot. (B600.15.w15)
      • If required, also use additional skin sutures or tissue glue. (B600.15.w15)
      • A subcuticular suture is relatively time-consuming to place. (B602.22.w22)
      • Subcuticular sutures rather than standard skin sutures are preferable for skin closure. (B615.8.w8)
    • OR Surgical staples (B600.15.w15, B602.22.w22)
      • These are quick to use. (B600.15.w15)
      • Patients rarely manage to remove these. (B600.15.w15)
    • Tissue glue can be used (B602.22.w22, J34.17.w1) but is occasionally removed by the rabbit. (B602.22.w22)
      • Preferably use tissue glue to reinforce a suture layer rather than alone. (J34.17.w1)
  • Avoid the use of Elizabethan collars on rabbits; these generally cause stress to the rabbit, as well as preventing normal caecotrophy. (B600.15.w15, B602.22.w22)

Hollow abdominal organs

  • Single interrupted sutures should be placed 2 - 3 mm apart and 2 - 3 mm from the cut edge. (B600.15.w15)
  • A modified Gamgee suture is suitable. (B600.15.w15)
  • Bring sutures through the submucosal layer, which has abundant collagen and is important in wound healing. (B600.15.w15)
  • Bring the wound edges into apposition. Take care not to crush the tissue or evert the mucosa. (B600.15.w15)
  • It is often necessary to penetrate into the lumen. (B600.15.w15)
    • Avoid penetrating the lumen when suturing the bladder, as this presents a risk of calculus formation using the suture material as a nidus. (B600.15.w15)
  • Use a single, not a double layer closure, to avoid excessive narrowing of the lumen diameter. (B600.15.w15)
  • Avoid inverting sutures, since these induce stenosis. (B600.15.w15)
Minimising blood loss
  • Note: the blood volume of a rabbit is 55-65 mL/kg (B600.15.w15) 50 to 60 mL/kg (J213.11.w2), in contrast to 90 mL/kg in the dog. Losses up to 10% of total blood volume may not cause ill effects, but hypovolaemic shock will occur with losses above 20 - 25%. (B600.15.w15)
    • Adequate haemostasis is important. (J34.17.w1)
    • Bipolar electrocautery may be useful to minimise blood loss. (B615.8.w8, J34.17.w1)
    • Electrosurgical instruments can be used either for haemostasis following cutting of the skin with a scalpel blade, or used alone for cutting an coagulation (blended current mode). 
    • Note: the blood volume of a rabbit is 55-65 mL/kg (B600.15.w15) 50 to 60 mL/kg (J213.11.w2), in contrast to 90 mL/kg in the dog. Losses up to 10% of total blood volume may not cause ill effects, but hypovolaemic shock will occur with losses above 20 - 25%. (B600.15.w15)
      • Adequate haemostasis is important. (J34.17.w1)
      • Bipolar electrocautery may be useful to minimise blood loss. (B615.8.w8, J34.17.w1)
    • (B534.43.w43f)
Post-operative care
  • Keep the rabbit away from predators such as cats and dogs; preferably have a separate area. (B601.16.w16)
  • If a cage has previously been used for a predatory species such as a cat, make sure it has been thoroughly cleaned and deodorised before it is used for a rabbit. (B601.16.w16)
  • Prepare a recovery area in the pre-operative period, making sure it is at an appropriate temperature for when it is needed. (B601.16.w16)
  • Consider hospitalising the rabbit's companion rabbit as well (in the same cage) to reduce stress and encourage feeding. (B601.16.w16)
  • If a limb has been injured, ensure good immobilisation. (B600.5.w5)
  • Minimise disturbance including observation and handling. (J15.13.w7)
  • Temperature
    • Initially provide an environmental temperature of 35 C, reducing to 26 - 28 C as the rabbit resumes normal activity. (B601.16.w16)
    • Provide supplemental heat until the rabbit is sufficiently recovered to resume normal thermoregulation. (J15.30.w2, B600.5.w5)
  • Fluids:
    • Provide water, but take particular care that the rabbit cannot spill a water bowl and get wet and chilled. (B601.16.w16, P113.2005.w3)
    • Give supplemental fluids as required. (J15.30.w2, P113.2005.w3)
  • Feeding:
    • Encourage the rabbit to eat. (B601.16.w16, P113.2005.w3)
    • Offer tempting foods: as well as making hay available, offer fresh grass, dandelions, fresh vegetables and any preferred foods of the individual rabbit. (B600.5.w5)
      • Offer sweet foods - rabbits have a sweet tooth and sugar-rich foods may tempt an anorectic rabbit to eat. (B539.1.w1)
    • If the incisors have been removed, offer food which is soft or grated. (B600.5.w5)
    • Syringe feed if needed until self-feeding occurs. (J15.30.w2)
      • Liquidised vegetables can be given. (J15.13.w7)
      • Give 10 - 20 mL every 8 -12 hours. (P113.2005.w3); give 3 - 15 mL four to six times a day. (B539.1.w1)
    • Consider using a motility stimulant to reduce the risk of post-anaesthetic ileus developing, particularly following gastrointestinal surgery. (B601.3.w3, B601.16.w16, J15.30.w2)
      • Metoclopramide 0.5 mg/kg subcutaneously every 8 -12 hours. (J15.30.w2)
      • Ranitidine 2 mg/kg orally or subcutaneously every 12 hours. (J15.30.w2); 2 - 5 mg/kg every 12 - 24 hours. (B601.3.w3,)
      • Cisapride 0.5 mg/kg orally every 8 - 12 hours. (J15.30.w2)
        • This has been withdrawn from the market in the UK (B601.3.w3) and in many other countries. (V.w5)
    • If large amounts of gas are present in the gastro-intestinal tract, consider giving simethicone, 20 - 40 mg/kg orally. (P113.2005.w3)
    • Encourage exercise to stimulate gastro-intestinal motility. (P113.2005.w3)
    • See: Food and Feeding for Mammals - Convalescent diets / Nutritional support
  • Bedding:
    • Provide warm, comfortable bedding such as veterinary fleece or towels until the rabbit is fully conscious and active, then good-quality hay or straw. (B601.16.w16, P113.2005.w3)
    • Hay provides reassurance (it is familiar, and can be burrowed into) as well as insulation, and also acts as a source of high-fibre food. (B600.5.w5, B601.16.w16, P113.2005.w3)

Monitoring, pain assessment and post-operative analgesia

  • Monitor general physical status for the first 24 hours - e.g. body temperature, auscultation of the chest, pulse rate, water and food intake, production of faecal pellets. (B534.43.w43f)
    • If the rabbit is sent home, instruct the owners to make sure it is eating and passing hard faeces within 24-48 hours. The rabbit should be examined if it is not eating within 24 hours. (B600.5.w5)
    • It may be preferable to keep the rabbit hospitalised until it is eating voluntarily and passing faecal pellets. (P113.2005.w3)
  • Assess for pain - see Physical Examination of Mammals - Observation
    • e.g. reluctance to move, hunched posture, anorexia, teeth grinding, self-trauma, raised body temperature, increased respiratory rate, excessive or lack of drinking, head elevation or extension, pushing the abdomen onto the floor, unusual aggression, an anxious facial expression, reduced faecal output, decreased drinking and occasionally vocalisation may all be signs of pain in rabbits. (B601.3.w3, B534.43.w43f, P113.2005.w3)
    • Careful observation is needed to detect subtle signs of pain. (B600.5.w5)
  • Give post-operative analgesia as indicated by assessment of the injury or surgery. (B602.22.w22, J15.13.w7)
    • e.g. buprenorphine, 0.05 mg/kg subcutaneously twice daily. (B534.43.w43f)
    • A NSAID may be given as well as buprenorphine for extensive procedures and may be sufficient alone following a relatively simple procedure. (P113.2005.w3)
    • Note: Post-operative analgesia is very important to restore appetite and gastro-intestinal motility as well as to reduce pain and stress. (B600.5.w5, P113.2005.w3)
    • Assume that pain is present and give analgesia following surgery. (B601.3.w3)
    • For further information on post-operative analgesia see section above: Analgesia
Ferret Consideration
  • In general, surgery in ferrets can be treated similarly to surgery in cats, (B117.w11, B631.23.w23) but allowing for their smaller size, so that smaller instruments, suture material etc. may be required, and magnification facilities (2.5X  - 5X), for example a magnifying loupe, are sometimes useful. (B631.23.w23)
    • No 11 and No. 15 scalpel blades are useful. (B631.23.w23)
    • Suture material in the 2.0 - 0.4 metric (3/0 - 8/0 USP) is needed. (B631.23.w23)
  • To minimise blood loss, electrosurgery units (1.0-1.3 MHz) or preferably radiosurgery units (4.0 MHz), which produce less heat, are useful. (B631.23.w23)
  • For control of bleeding during adrenal and pancreatic surgeries, and during splenic or hepatic biopsies, gelatine sponge haemostatic material is very useful. (B631.23.w23)
  • Owners should be informed that hair on areas shaved for surgery may not regrow for up to four months (depending on seasonal hair growth patterns). Additionally, the skin in shaved areas may appear blue before hair regrows (Blue Ferret Syndrome). (J29.6.w3)
  • When a small skin incision has been made, subcuticular sutures can be used without an additional layer of skin sutures. (J29.6.w3)
  • Suture materials should be 4-0 or smaller sizes. (J29.6.w3)
  • Gentle handling of tissues minimises tissue damage and reduces the pain associated with surgical procedures. (J29.14.w1)
  • Note: ferrets will chew sutures if the sutures are uncomfortable. (J15.24.w5)
  • Preferably close the skin using a subcutcular layer of sutures and a fine, absorbable suture materil (e.g. polyglacton 910 (CCoated Vicryl, Ethicon). (J15.24.w5)
Presurgical preparation
  • As with all patients, aseptic techniques are important. Shaving should be carried out carefully to avoid skin damage. (B631.23.w23)
  • There is a risk of heat loss leading to hypothermia (Chilling - Hypothermia (with special reference to Waterfowl, Hedgehogs, Bears, Lagomorphs and Ferrets)) during long procedures. To reduce these losses, the area clipped should be minimised, alcohol rinses avoided during aseptic preparation, and external heat sources should be available. (B631.23.w23)
  • To improve anaesthetic monitoring and surgical visualisation, transparent drapes are recommended. (B631.23.w23)
  • During surgery, a heat source should be used under the ferret and if necessary also an overhead heat lamp, to avoid hypothermia. (J29.6.w3)
  • Intravenous fluids and and fluids used for flushing (e.g. in the abdomen) should be warmed prior to use. (J29.6.w3)
Monitoring
  • Core body temperature and blood pressure should be monitored. Capnography, ECG, pulse oximetry etc. are useful if available. (B631.23.w23)
    • Monitoring body temperature with a rectal thermometer enables decision-making regarding whether or not post-operative heat is required. (J29.6.w3)
    • In the post-operative period, if supplementary heat is provided, close monitoring for hyperthermia is important. (J29.6.w3)
Post-surgery
  • Provide adequate analgesia (preferably starting before surgery or before the ferret regains consciousness. (B232.18.w18)
    • Note: ferrets are stoic animals and often mask signs of pain. Assume that surgery is painful, and provide analgesia accordingly. (J15.24.w5)
    • See section above: Analgesia
  • If the ferret is anorectic, assisted feeding may be needed. (B232.18.w18) 
  • If prolonged antibiotic treatment is required, give orally or subcutaneously not intramuscularly, because of the small muscle mass of ferrets. (B232.18.w18) 
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.

General

  • In all primates, an intradermal pattern of skin sutures is recommended to minimise the risk of the self-trauma post-operatively. (D425.3.21.w3u)
  • If surgery is contaminated, e.g. if the gastro-intestinal or urinary tracts are entered, the surgical site should be flushed well to minimise contamination, and intra-operative and post-operative prophylactic antibiotics should be given for up to 24-28 hours post-surgery. (B671.13D.w13d)

Notes for surgery of specific organs

  • The pyriform uterus of nonhuman primates is normally found within the pelvic cavity. (B671.13D.w13d)
  • The wall of the gravid uterus can be 2-3 cm thick . If a Caesarean section is carrier out, the edges should be oversewn with a simple continuous suture, followed by inverting sutures to close the incision. (B671.13D.w13d)
  • A midline approach is recommended for renal surgery in nonhuman primates. (B671.13D.w13d)
  • A left subcostal incision is recommended for splenic surgery in great apes, because the spleen is firmly attached to the diaphragm and left body wall. (B671.13D.w13d)
  • There is a vermiform appendix in great apes. (B671.13D.w13d)
Associated techniques linked from Wildpro

Surgical techniques in Birds

Surgical techniques in Mammals

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Euthanasia

"Euthanasia" is a term used to describe an animal killed in a humane manner.
  • The use of euthanasia may be appropriate for example to prevent unnecessary suffering in a terminally ill animal, or in a wild animal in which treatment will not allow a return to the wild and for which there is no appropriate place in a conservation breeding or educational programme. In flock or herd medicine (the treatment of large numbers of birds or mammals), a few ill individuals may be euthanased and necropsied to provide accurate data allowing effective treatment of the remaining individuals. Humane methods of killing (i.e. euthanasia) should also be used if pest species are live-caught and cannot be appropriately relocated.
  • The method used will vary depending on the size and type of animal concerned and the availability of different physical and chemical means of euthanasia. Both physical and chemical methods of euthanasia may be extremely effective and humane in the right circumstances, and ineffective or inhumane if used incorrectly. In some countries the use of chemical methods of euthanasia may be restricted to e.g. veterinarians, due to regulations on the handling and storage of the required drugs.
  • If euthanasia is the most appropriate option for wildlife patient, the decision to euthanase should be made as soon as possible. This decision may be made at the time of initial assessment. During treatment, a daily review should include deciding whether treatment should be continued, the animal is fit for release, or the animal should be euthanased. (B545.8.w8)

(B13.14.w19, B32.1.w34, B36.5.w5, B545.8.w8)

Waterfowl Consideration

  • Cervical dislocation may be employed as a humane method of euthanasia of waterfowl if carried out by trained personnel.
  • Chemical euthanasia with a standard euthanasia solution (e.g. pentobarbitone sodium) may also be used. Euthanasia solutions are usually designed for intravenous use. N.B. Before use the manufacturer's data sheet on the agent or agents concerned should be consulted, taking particular note of any contra-indications and operator warnings.

(B36.5.w5)

Crane Consideration
  • Euthanasia may be required e.g. following leg fracture where the distal portion of the limb is devitalised and use of a prosthesis is inappropriate or unsuccessful, or when both legs are affected. (J312.26.w1)
  • Euthanasia should be carried out by intravenous injection (Venipuncture in Cranes) of barbiturate overdose; the crane must be held by someone experienced at restraining these birds and it may be preferable to anaesthetise with isoflurane first. (D281)
Bear Consideration
  • Euthanasia may be required for bears suffering from terminal illness or in severe pain. (J311.27.w1)
  • Further information on euthanasia of bears is provided in: Wildlife Casualty Euthanasia - Mammal Considerations

Lagomorph Consideration

Whatever technique is used, the aim should be to provide a humane death, painless for the rabbit, causing as little apprehension and fear as possible, and a rapid death. (B611.4.w4)

  • As far as possible after the concerns regarding the welfare of the animal have been met, euthanasia should cause as little as possible emotional distress to observers. (B611.4.w4)
  • This is particularly important when euthanasing pet rabbits in the presence of their owners. (V.w5)

Methods of euthanasia

  • The most common method of euthanasia in a veterinary setting is by intravenous injection of pentobarbital, 150 mg/kg. (B611.4.w4, B614.5.w5)
  • In research settings, carbon dioxide at concentrations above 40% has been found to produce rapid, painless onset of anaesthesia. Carbon monoxide is also rapid and painless, so long as a pure form is used (not e.g. car exhaust fumes, which contain contaminants which may be noxious to the animal), but is less safe for humans to use. (B611.4.w4, B614.5.w5)
  • Further information on euthanasia methods, including details of physical methods which may be used in the field in the absence of chemical agents, is provided in Wildlife Casualty Euthanasia (with special reference to UK Wildlife) - Mammal Considerations
Domestic rabbits
  • The aim is to minimise the stress on the rabbit (allowing for both species and the individual rabbit) and provide a peaceful death. (B601.2.w2)
  • Stressors include strange surroundings and strange people. If there is a member of staff the rabbit is familiar with, or who has a rapport with rabbits, they should play a role. (B601.2.w2)
  • A non-slip surface should be provided. (B601.2.w2)

For aggressive or fearful rabbits

  • Tranquillization or sedation should be used; this "will relax the rabbit and give the owner a few minutes to 'say goodbye' before the injection of the euthanasia agent." (B601.2.w2)
    • Acepromazine maleate, 0.7 - 1.5 mg/kg subcutaneously is useful. 
    • Medetomidine, 0.25 mg/kc subcutaneously can be given for marked sedation as well as moderate anaesthesia, however the peripheral vasoconstriction will make it more difficult to place an intravenous catheter for injection of the euthanasia agent.
    • Acepromazine 0.5 mg/kg plus butorphanol 0.1 mg/kg subcutaneously provides marked sedation and some analgesia, with peripheral vasodilatation.
    • Fentanyl/fluanisone (Hypnorm), 0.3 mL/kg subcutaneously provides greater analgesia.

    (B601.2.w2)

Wild lagomorphs
  • Casualty wild lagomorphs with disabling injuries likely to compromise their survival in the wild should be euthanased unless an appropriate permanent home is available. (B284.10.w10)
    • This includes e.g. limb injuries (reducing ability to run away), eye or ear injuries reducing the ability to detect predators and broken/abnormal teeth. (B284.10.w10)
  • Wild lagomorphs can be euthanased with intravenous pentobarbital; a sedative may be given firs, or the drug may be given by intraperitoneal injection. (B284.10.w10)
  • Further information on the euthanasia of wild lagomorph casualties, including physical methods which may be used in the field in the absence of chemical agents, is provided in Wildlife Casualty Euthanasia (with special reference to UK Wildlife) - Mammal Considerations
Ferret Consideration Note: Bonds between an owner and a pet ferret may be strong. (B631.18.w18)
  • Euthanasia may be carried out by intravenous or intracardiac administration of barbiturate euthanasia solution. (B631.18.w18)
    • Euthanasia solutions can be given at the same dose rates as in cats or dogs. (J29.13.w2)
    • Consider anaesthetising the ferret and placing an intravenous catheter, then using this route for administration of the the euthanasia solution in the owners presence; this may be less distressing for the owner than watching an intracardiac injection. (B631.18.w18)
      • Either an over-the needle catheter or a butterfly catheter may be used. (J29.13.w2)
      • Consider first administering thiopental (2.2 mL 20% thiopental per kg) before giving the euthanasia solution. (J29.13.w2)
    • Note: In a collapsed ferret, or after sedation, the veins may be difficult to access. (B631.18.w18)
    • Intracardiac injection should not be carried out in a conscious ferret; if this route is to be used, the ferret must first be anaesthetised. (B339.9.w9, B631.18.w18, J29.13.w2)
      • Sedation can be carried out using 20-40 mg/kg ketamine given intramuscularly prior to the intracardiac injection. (J29.13.w2)
      • Remember that the heart is placed relatively caudally in the chest, between the 7th and 10th intracostal spaces. (J29.13.w2)
      • Explain the procedure to the owner before carrying out intracardiac injection. (B631.18.w18)
    • Health and safety of the owner must be considered if the ferret is given gaseous anaesthesia (ensure any anaesthetic gases are properly scavenged, for example using a scavenging face mask). (B631.18.w18)
  • Intraperitoneal injection may be used if there is no need for post mortem examination of the peritoneal cavity. (B339.9.w9)
Bonobo consideration Note: There is very little published information available on veterinary care specifically in bonobos. In general, treatment and care of bonobos is the same as treatment and care of Pan troglodytes - Chimpanzee in particular and of the other great apes and other primates. Great ape treatment and care is commonly based on the treatment for their close relatives, Homo sapiens - Humans.
Associated techniques linked from Wildpro

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Authors & Referees

Authors

Debra Bourne MA VetMB PhD MRCVS (V.w5); Bridget Fry BSc, RVN (V.w143)

Referees

Suzanne I Boardman BVMS MRCVS (V.w6); Frances Harcourt-Brown BVSc FRCVS (V.w140); Marla Lichtenberger DVM, DACVECC (V.w124)

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