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- 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. Vetrap 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, B14, B116.30.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 2-4cm 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, B14, B116.30.w3) |
| 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 cm² and another 18 cm² 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
(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)
- Puncture wounds (Lacerations & Punctures, including bite
wounds), which may result in an abscess (Abscessation in Lagomorphs and Ferrets).
(B651.9.w9,
B652.10.w10,
J213.7.w5)
- Lumpy jaw, from bacteria entering the oral tissues (Actinomyces infection in Lagomorphs and Ferrets.
(J213.7.w5)
- Possible fungal infections, which may cause erupted skin. (J213.7.w5)
- Electrical injuries, which can cause burn wounds (Burns and Smoke Inhalation (with special reference to Waterfowl, Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)). (J213.7.w5)
- Lacerations (Lacerations & Punctures, including bite
wounds). (B651.9.w9)
- Incised wounds (Lacerations & Punctures, including bite
wounds). (B651.9.w9)
- Gun shot wounds (Lacerations & Punctures, including bite
wounds). (B651.9.w9)
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 |
|
|
|
|
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 98°F (36.7°C), 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).
|
| 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)
- Use the intravenous route if an intravenous catheter can be
placed, otherwise via an intraosseous catheter. (B602.2.w2,
P120.2006.w6)
See:
- At catheter 22-gauge or larger should be used to avoid red blood
cell lysis. (J513.2.w3)
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
|
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:
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|
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(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 39-41.6°C. 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).
SPECIFIC IMMEDIATE RESPONSE TO
RESPIRATORY ARREST
Disconnect from anaesthetic gas.
Increase oxygen flow rate.
Press on sternum gently, 40 to 50
cycles per minute (B13.36.w16);
unlikely to be effective and may cause physical damage (B14).
Intubate or insert air
sac tube if neither is present (see above for technique).
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
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
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.
Adrenaline may be given,
0.5-1.0mg/kg intravenous, in response to cardiac arrest, anaphylactic shock or bronchial
spasm (P3.1999b.w2).
Dexamethasone (synthetic corticosteroid) 4mg/kg (1mg/kg in raptors) intramuscular or subcutaneous in case of shock.
(P3.1999b.w2).
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)
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
- Check correct placement in trachea, not oesophagus
- Not too long (causing bronchial intubation)
- Too narrow
- Obstructed
- Kinked
- Disconnected from anaesthetic machine
- No anaesthetic agent
- Incorrect setting on dial
- Wrong agent in vaporizer
- Inaccurate vaporizer calibration
- No gas in cylinders
- Flow meter - incorrect siting
- Flow meter - failed
- Connections (vaporizer-machine, or breathing system) - leakage
- Breathing system obstruction.
(P3.1999b.w2) |
| Bear Consideration
 
 
|
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
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 (> 41°C) (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 µg/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)
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
- Oral sedation/light anaesthesia has been carried out in captive
bears by allowing controlled licking of a mixture of honey and carfentanil. (B336.51.w51,
J1.31.w11,
J2.32.w2, J4.217.w3)
- This has been alone for handling, blood sampling etc., and prior to
hand injection or darting with further anaesthetic drugs. (B336.51.w51,
J1.31.w11,
J2.32.w2, J4.217.w3,
P2.1999.w2)
- Oral ketamine,
100 mg, was used in a weak, injured wild Ursus americanus - American black bear
cub (approx. 6-9 kg bodyweight) to facilitate handling and
radiography. (P62.9.w1)
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 " µg/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 µg/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.
- 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 " µg/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 µg/kg |
Tiletamine-zolazepam 1.7 mg/kg. |
Atipamezole, 3-4 x the medetomidine dose
in µg/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 µg/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 µg/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 µg |
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 µg/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. |
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)
|
|
|
(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
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
Pulse/Heart:
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
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)
- Check the rabbit's airway is unobstructed.
- Give 100% oxygen - by endotracheal tube if placed, otherwise
by facemask
- If the rabbit is not already intubated, try to place an
endotracheal tube.
- Start IPPV (intermittent positive pressure ventilation).
- Give external cardiac massage, 70-90 per minute.
- 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)
- 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)
- 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.
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
- Correct placement in trachea, not oesophagus?
- Too long or placed too deep (causing bronchial intubation)/
- Too narrow?
- Obstructed?
- Kinked?
- Disconnected from anaesthetic machine?
- No anaesthetic agent?
- Incorrect setting on dial?
- Wrong agent in vaporizer?
- Inaccurate vaporizer calibration?
- 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 µg/kg intramuscularly or intravenously. (J513.7.w3)
- 100 µg/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 µg/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)
- 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)
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
Bears
Lagomorphs
Ferrets
|
|
|
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). |
| Bear Consideration |
Surgery can be carried out using the same general principles as for dogs,
with allowance for the larger size of adult bears.
- The commonest surgical procedures performed on bears are repairs of
traumatic injuries. (B16.9.w9)
- In bears rescued from bile farms, severe gall bladder pathology
occurs and cholecystectomy (gall bladder removal) is required. (P3.2006b.w1,
P503.1.w7)
- Dental treatment is required not uncommonly. See:
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
- Bears
- Hedgehogs
- Rabbits
- Ferrets
|
Rabbits:
