U5 O2 - Small Mammal, Bird and Reptile Emergency and Critical Care Flashcards
Small Mammal, Bird and Reptile Emergency and Critical Care
What is the ‘Coelomic cavity’ - With reference to birds and reptiles?
‘Coelomic cavity’ - With reference to birds and reptiles. This is the common single body cavity made up of what would have been the thorax and abdomen in mammalsneither birds nor reptiles have a diaphragm
Define ‘Ectothermic’ - With reference to reptiles.?
‘Ectothermic’ - With reference to reptiles. This means that they are dependent on their environment to maintain their body temperature
What is the preferred body temperature with reference to reptiles?
‘Preferred Body Temperature (PBT)’ - With reference to reptiles. This is the temperature at which the reptile’s body functions operate optimally
What is ‘Preferred Optimum Temperature Zone’ - With reference to reptiles?
‘Preferred Optimum Temperature Zone’ - With reference to reptiles. This is the environmental temperature zone in which a reptile needs to be kept in order for it to maintain its PBT
Define ‘Uric acid’ - With reference to birds and reptiles?
‘Uric acid’ - With reference to birds and reptiles. This is the main excretory waste product of protein metabolism- as opposed to urea in mammals. It forms the ‘whites’ of the droppings.
What is the normal heart rate for a domestic rabbit?
130-325
larger breeds lower rates
What is the normal heart rate for mice?
500-600
What is the normal heart rate for a rats?
260-450
What is the normal heart rate for a gerbils?
300-400
What is the normal heart rate for syrian hamsters?
280-412
What is the normal heart rate for Russian hamsters?
300-460
What is the normal heart rate for a Guinea pigs?
190-300
What is the normal heart rate for a Chinchilla?
120-160
What is the normal heart rate for a Chipmunk?
150-280
What is the normal heart rate for a ferret?
200-250
What is the normal respiratory rate for a domestic rabbit?
30-60
What is the normal respiratory rate for mice?
100-250
What is the normal respiratory rate for rats?
70-150
What is the normal respiratory rate for syrian hamsters?
33-127
What is the normal respiratory rate for Russian hamsters?
300-460
What is the normal respiratory rate for chinchillas?
50-60
What is the normal respiratory rate for chipmunks?
60-90
What is the normal respiratory rate for ferrets?
33-36
What is the normal respiratory rate for guinea pigs?
90-150
What is the normal respiratory rate for gerbils?
90-140
What is the normal body temperature for domestic rabbits?
38.5-40
What is the normal body temperature for mice?
37-38
What is the normal body temperature for rats?
37.6-38.6
What is the normal body temperature for gerbils?
37-38.5
What is the normal body temperature for syrian hamsters?
36.2-37.5
What is the normal body temperature for Russian hamsters?
36-38
What is the normal body temperature for guinea pigs?
37.2-39.5
What is the normal body temperature for chinchillas?
37-38
What is the normal body temperature for chipmunks?
37.8-39.6
What is the normal body temperature for ferrets?
37.8-40
What is the average weight for domestic rabbits?
1.5-10kg
Netherland dwarfs – Belgian Hares
What is the average weight for mice?
25-50g
What is the average weight for rats?
400-1000g
What is the average weight for gerbils?
50-60g
What is the average weight for syrian hamsters?
90-150g
males larger
What is the average weight for russian hamsters?
30-60g
What is the average weight for guinea pigs?
600-1200g
What is the average weight for chinchillas?
400-550g
What is the average weight for chipmunks?
55-150g
What is the average weight for ferret?
- 5-1kg
females) 1-2kg (males
What is the average life span in years for a domestic rabbit?
6-10
What is the average life span in years for a mouse?
2-3
What is the average life span in years for a rat?
3-4
What is the average life span in years for a gerbil?
2-2.5
What is the average life span in years for a syrian hamster?
2-2.5
What is the average life span in years for a Russian hamster?
1.5-2
What is the average life span in years for a guinea pig?
3-8
What is the average life span in years for a chinchilla?
6-10
What is the average life span in years for a chipmunk?
8-12
What is the average life span in years for a ferret?
5-10
What should a respiratory assessment of a small mammal include and what are the important considerations?
Severe respiratory distress is one of the main concerns- this may be obvious with clinical signs such as stertor, wheezes and whistles. Subtle changes such as increased respiratory rate, nasal discharge/blocked nares may all indicate relatively serious respiratory pathology. Most small mammals have a relatively small lung size in relation to their overall body. They, therefore, have less respiratory reserve than larger mammals such as cats or dogs; thus, respiratory disease/pathology can have a more serious effect. In addition, respiratory infections, particularly those associated with Pasteurella spp. and Mycoplasma spp., are very common in small mammals.
Unlike cats and dogs, lung tissue is not segmental so infection tends not to be localised.
The patient should be examined in the cage first, using dimmed, blue or red lighting.
If in doubt, the patient should be admitted and placed in an oxygen enriched atmosphere +/- nebulisation with saline, before any examination is attempted.
Following this an assessment of the patient should be made to see if physical restraint is appropriate. It may be that the preferred/ safest option is for the patient to be sedated/anaesthetised e.g. midazolam / isoflurane in 100% oxygen.
What should the initial assessment of a small mammal that presents in an emergency include?
Initial assessment should consider the following:
1) Is the animal active (should be if in the consulting room-exceptions could include the more nocturnal hamsters)?
2) What is the body condition- is the animal overweight/ underweight? Is the coat in poor condition or is there evidence/ suggestion of parasitism?
3) Are the eyes open or closed?
4) Are the corneas bright?
5) Is there respiratory noise? If so what?
6) Is there abdominal breathing suggestive of dyspnoea?
7) Are the nares clear or is there any other discharge e.g. serous, purulent or haemorrhagic?
8) Is there any faecal clumping at the rear end/evidence of diarrhoea?
9) What is the recent faecal output like?
10) Is there increased urine output/sodden bedding?
11) Is there any blood or melaena in the faeces/urine (may indicate haematuria such as is seen in heavy metal poisoning/cystitis/GI ulceration)?
12) Is there undigested food in the faeces? (may indicate intestinal malabsorption/maldigestion)
13) What is the stance of the animal like? Is it able to support its weight off the ground or is it collapsed?
14) Is there evidence of vomitus/regurgita in the cage/on the fur?
These assessments can be made relatively quickly and often without handling the patient.
What should a detailed examination in a small mammal include and who should carry this out?
Detailed examination- (Depending on the clinical presentation and species some of this examination may have to be performed by a veterinary surgeon- however the veterinary nurse should be aware of the likely procedures to be performed, the required equipment and restraint).
As with all species it is important to be familiar with the findings that would be expected in the healthy patient so that abnormalities can be readily identified.
A detailed examination should include the following:
1) What is the animal’s weight AND what is the muscle condition over the pelvic area like? (body scores of 0-5/ or 0-9 may be given as with other animals)
2) An intra-oral examination using an oroscope. This should allow a close examination of the tongue, the cheek teeth (molars and premolars) and the
palate- it can be difficult to see all of this in the conscious patient. The epiglottis may also be visualised. Abnormalities include overgrown cheek teeth (typically spurs forming on the tongue side of the mandibular cheek teeth and the cheek side of the maxillary cheek teeth); ulcers on the tongue and cheeks; blood
tinged saliva, abscesses; impacted food and broken teeth.
3) A detailed examination of the nares and the eyes. Clinical signs to be aware of include crusting of secretions around the nares or obvious mucopurulent
discharge (common in rabbits); wet fur on the fore paws often associated with nasal discharge; chromodacryorrhoea (red staining around the eyes in ratsthis is due to a viral condition which causes increased tear production and is often associated with stress/debilitation); increased size of the globe
(glaucoma) or protrusion of the eye; cataracts; anterior uveitis (which may suggest Encephalitozoon cuniculi infection in rabbits).
4) A detailed examination of the fur and skin. This may give an indication of the level of husbandry (e.g. matting of fur in long-haired breeds), parasite infestation (dandruff with Cheyletiella infestation in rabbits; alopecia and pruritus with Trixicara caviae infestation in guinea pigs; increased black earwax in Otodectes cynotis infestation in ferrets, guinea pigs or rabbits).
5) A detailed auscultation of the lungs and heart. The lungs may be harsh sounding with infections; however, lung consolidation is not uncommon, particularly in rodents and rabbits, so no lung sounds may be heard over these areas. Heart rates are often very fast in small mammals and so murmurs are rarely heard, other than in rabbits and ferrets. Arrhythmias, however, may be
heard even in smaller rodents.
6) A detailed examination of the limbs may be made for evidence of fractures; or evidence of metabolic bone disease, which may result in abnormally shaped or
thickened bones and decreased range of joint movement (N.B. this would be significant when handling as there may be an increased risk of associated
pathological fractures). Special care in examination is vital in these (typically) prey species, as they are unlikely to show pain response to palpation of a fracture. In addition, in guinea pigs, pain response or swelling of elbows, carpal and tarsal joints is commonly associated with hypovitaminosis C (scurvy).
7) A detailed examination of the perineum and abdomen should also be made. The abdomen is easily palpated in ferrets and rabbits and may result in
detection of masses or foreign bodies. The liver should not normally project beyond the caudal ribcage in any of the species mentioned here. The perineum should be examined for evidence of urine and faecal soiling and matting. This is the prime area for myiasis (blow fly strike) particularly in incontinent/diarrhoeic rabbits
What action should be taken in a rabbit in respiratory arrest?
Rabbits
Should breathing stop, or hypoxia be detected, then emergency ventilation will be required. This is best achieved by immediate endotracheal intubation.
Intubation may be achieved blindly with the rabbit in sternal recumbency. The rabbit’s head is lifted and the endotracheal tube (ET) advanced slowly until breathing sounds are heard through the tube. The tube may then be quickly advanced on inspiration. If a transparent tube is used then seeing condensation from the rabbit’s breath, when the tube is over the glottis, can aid intubation. If the rabbit has stopped breathing then
this technique becomes extremely difficult; therefore, a laryngoscope with a Wisconsin 0 paediatric blade, Flecknell small animal laryngoscope or long auroscope nozzle can be used instead to visualise the glottis (Heard, 2004). This is best achieved with the
rabbit in sternal recumbency, with the chin elevated and the tongue pulled forward. A guide wire/urinary catheter may be inserted through the glottis first and the ET tube threaded over the top. Alternatively, a fine endoscope or needle scope may be used
as a guide wire instead, threading the ET over the scope prior to intubation. Once through the glottis, the ET tube maybe advanced and the scope retracted easily.
Direct intubation may, however, be difficult in the rabbit owing to the narrow oral cavity and relatively large size of the tongue caudally; or the presence of an obstruction, such as a pharyngeal abscess or foreign body. In an emergency, therefore, it may be
necessary to pass a long through-the-needle catheter into the tracheal lumen between two tracheal rings ventrally. A luer adaptor may be attached to allow connection to an anaesthetic circuit for oxygen insufflation.
A tracheostomy may also be indicated with the technique being like that for a cat or a dog. This is an act of veterinary surgery as outlined in The Veterinary Surgeon’s Act 1966. Veterinary nurses should be aware of the procedure so that they can provide appropriate assistance to the veterinary surgeon performing the procedure.
The main difference is that some breeds of rabbit, particularly in the doe, have large dew flaps with plentiful subcutaneous fat depots which may make tracheotomy surgery challenging.
What is the technique for a tracheostomy in a rabbit?
Technique: It is important for the veterinary nurse to be aware of the equipment needed and the surgical procedure to provide appropriate assistance to the veterinary surgeon. A longitudinal incision is made over the trachea caudal to the larynx, followed by blunt dissection onto the trachea itself. The ventral 25% of one tracheal ring is removed and a 2mm tracheostomy tube, or in an emergency, an ET tube is inserted.
This may then be attached to a breathing system for ventilation. Intermittent positive pressure ventilation (IPPV) may then be performed at a rate of 10-20 breaths/minute at a pressure of 10cm H20 (approximately enough to allow the chest to rise and fall by one third of the thorax diameter). It is important to provide appropriate pressure excessive pressure can cause damage to the lungs.
If intubation is a rabbit is challenging what are the options?
If intubation is challenging, insertion of an appropriate sized rabbit Vgel®, is a possibility, although they should not be used without simultaneous capnography. The
size of Vgel® is very important, because if there is a poor fit it is possible for the stomach to inflate. If intubation is not possible then, either, a tight-fitting face mask may be applied, connected to an anaesthetic breathing system with a high flow rate of oxygen (4-5 litres); or an Ambu bag could be used to force ventilate the rabbit.
Alternatively, moving the rabbit in a see-saw manner may aid ventilation by moving the abdominal viscera backwards and forwards onto the diaphragm and acting as apump mechanism (Briscoe and Syring, 2004). This works on the basis that most of the
impetus for inspiration comes from the flattening of the diaphragm rather than the outward movement of the ribcage.
IF a ferret goes in to respiratory arrest, describe how you would intubate and apply IPPV?
Intubation of ferrets is relatively easy to perform. If the ferret is unconscious and has stopped breathing, the mouth may be opened and the epiglottis visualised at the base of the tongue. A small spray of lidocaine onto the epiglottis may aid passage of the tube. Once intubated, the ET tube cuff may be inflated, with care, and intermittent positive pressure ventilation (IPPV) may be performed, if required, at a rate of 10-15
breaths per minute and 10cm H20 pressure as for rabbits
How do you apply oxygen in a ferret that is oxygen-dependent?
If voluntary breathing is still occurring, then a face mask with 100% oxygen, or an intranasal 2-3g French catheter can be placed. The latter can be pre-measured from the tip of the nares to the lateral canthus of the eye and marked. The nostril is deeply
sprayed with lidocaine, using a catheter attached to a syringe. The catheter is then inserted as for a cat. Sedation is usually required to perform this procedure unless the ferret is much debilitated.
How do you apply oxygen in a rodent that is oxygen-dependent?
Rodents
Providing a direct airway may be very difficult in rodents. Access to the epiglottis is difficult, owing to the rodent anatomy whereby the soft palate is locked around the epiglottis. This makes access via the mouth very challenging. If the rodent is still breathing, placing it in a small container and piping in oxygen may be all that is necessary.
If a rodent if not breathing, what three techniques may be attempted?
If the rodent if not breathing, three techniques may be attempted
1. The head of the rodent is placed into a face mask with 100% oxygen. With the rodent positioned in sternal recumbency, two fingers either side of the thorax can be used to perform massage.
2. With the rodent in dorsal recumbency, the trachea is grasped between the fingers of one hand. A 23-25 gauge over-the-needle catheter is passed between two cartilage rings, 1cm caudal to the larynx. The stylet is removed and the catheter is left in place. This may then be used to administer IPPV and 100% oxygen.
3. Tracheostomy- The rodent is placed in dorsal recumbency. A skin incision is made over the trachea, caudal to the larynx, and bluntly dissected down onto
the trachea as with the rabbit. The trachea may then be incised between cartilage rings to insert a tube..
For IPPV, a rate of 20-30 breaths per minute with a pressure of 8-10 cm H20 may be applied, like rabbits
What rate of compressions are delivered to a small mammal that has had a cardiac arrest?
If the patient has had a cardiac arrest, then the focus should be, as with other species, on maintaining a circulation until return of spontaneous circulation (ROSC). The rate of compressions to be delivered is the same as for cats and dogs (100-120/min) - how
these are delivered depends on the size of the patient.
How do you perform CPR on a rabbit?
Rabbits
In mammals <10kg, direct cardiac massage, by compressing the chest directly over the heart, is most effective at increasing thoracic pressure and forcing blood through the arterial vasculature. Heart
compression rates of 100-120 per minute need to be achieved in rabbits - the recommended technique to maximise cardiovascular output is circumferential chest compressions, as is used in human infants, where the chest is compressed over the heart from both sides at once. If there is not already IV access, then
intra-osseous access may be preferred in a patient in circulatory collapse, due to the small size of their veins
ECG leads should be attached to detect a heartbeat or any dysrhythmias. The type of dysrhythmia reported in rabbits during resuscitation techniques has thus far been different from that reported in cats and dogs. The latter may demonstrate electromechanical dissociation (pulseless electrical activity); whereas in rabbits, profound
bradycardia, ventricular asystole and ventricular fibrillation have been reported. Lichtenberger & Lennox (2012) advocate the use of vasopressin for asystole, ventricular fibrillation and pulseless electrical activity. Other authors suggest epinephrine/adrenaline/adrenaline may be used, via the intra-tracheal route (if the rabbit is intubated); or intravenously if no heart beat can be detected clinically
and on ECG (i.e. asystole). In the case of fine ventricular fibrillation, the use of epinephrine/adrenaline has been advocated to convert the electrical activity to coarse
ventricular fibrillation, which is easier to convert
Conversion of coarse fibrillation is based on the use of cardiac massage as described above; or if the clinic has access to defibrillation devices then the use of these
externally at 2-10 J/kg (starting at low energies and increasing if no response is achieved) may be performed (Costello 2004). Greater success is achieved with
defibrillation devices if three initial counter-shocks are applied at low energies.
The effectiveness of CPR can be assessed by detecting a pulse and return of spontaneous breathing. Again, as with cats and dogs, measurement of ETCO2, blood
gas analysis and Doppler can all be used to assess the effectiveness of CPR attempts
How do you perform CPR on a ferret?
Ferrets
As for rabbits, circumferential chest compressions should be started immediately: chest compression rates of 100-120 beats per minute should be attempted, where
possible. Mask ventilation is as effective as attempts to intubate in this situation
ECG leads may be applied to identify any cardiac activity. The commonest arrhythmias in ferrets are sinus tachycardia, atrial and ventricular premature contractions and atrial fibrillation. However, sinus bradycardia may be seen with ferrets that have an
insulinoma with associated hypoglycaemia. It should be noted that second degree AV block can occur as a normal finding in healthy ferrets. However, third degree or high second degree blocks are abnormal. Epinephrine/adrenaline/ adrenaline may be used, via the intratracheal route if intubated; or intravenously if no heart beat can be detected clinically and on ECG (i.e. asystole).
How do you perform CPR on a rodent?
Rodents
As for rabbits, circumferential chest compressions should be started promptly: chest compression rates of > 150 beats per minute should be attempted, where possible.
What drugs should be used in rabbits with severe bradycardia in an emergency situation and why?
If severe bradycardia is detected, glycopyrrolate should be used (Appendix Table 2), in preference to atropine, as 60% of domestic rabbits possess serum atropinesterases, making atropine less effective
What drugs should be used in ferrets with severe bradycardia in an emergency situation and why?
Atropine, however, may be used in ferrets as they do not have serum atropinesterases.
When might lidocaine be used in small mammals in an emergency situation?
Lidocaine may be administered intratracheally or intravenously, if ventricular arrhythmias, such as VPCs leading to ventricular tachycardia, occur (Appendix Table
2). However, lidocaine should not be used in cases of AV block or severe bradycardia, which are more commonly seen in rabbits or in hypoglycaemic
ferrets; or in asymptomatic ferrets, where 2nd degree AV blocks may be found.
If severe bradycardia is detected, as a result of an insulinoma in a small mammal what treatment may be required?
If severe bradycardia is detected, as may occur with an insulinoma, atropine or glycopyrrolate should be used (Table 3). In addition, intravenous dextrose 50%, at
0.25-2ml by slow intravenous bolus, should be administered. This can be followed with an oral glucose solution (such as the human preparation GlucoGel® (previously Hypostop®) used in insulin overdose). If this does not work, then an IV drip of 5% dextrose with dexamethasone should be started. Ongoing management of insulin involves prednisolone therapy and diazoxide (a diuretic that inhibits insulin release;
and promotes glycolysis and gluconeogenesis by the liver).
What species of small mammals can develop cardiomyopathy and valvular insufficiency and what treatment is usually required?
Cardiomyopathy and valvular insufficiency, with resultant congestive heart failure, are also seen in rabbits; as is atherosclerosis. Ferrets may also develop congestive heart failure secondary to cardiomyopathy and valvular insufficiency. The veterinary
surgeon’s initial treatment of congestive heart failure is likely to involve diuretic administration e.g. furosemide (IV) every 4-6 hours as required. ACE inhibitors have
been used, but rabbits are more susceptible than cats and dogs to their hypotensive side-effects. Therefore, reduced dosages and regular monitoring of the systolic blood pressure, using non-invasive techniques devised for cats, is advisable. ACE inhibitors (e.g. benazepril) have been used in ferrets to decrease the preload to the heart. Digoxin may be added where supraventricular arrhythmias are present.
Similar drugs may be used for rodents as are used for ferrets and rabbits. Cardiac disease, specifically dilated cardiomyopathies and associated congestive heart failure, may be seen commonly in guinea pigs and hamsters. Hamsters are also prone to bacterial endocarditis. Atropine may be used where bradycardia is present as, unlike rabbits, rodents do not possess serum atropinesterases.
Gerbils are prone to epilepsy, which is hereditary. Midazolam/diazepam may be used in the management.
What are the limitations of performing an ECG on rodents?
Rodents
Most rodents have such a fast heart rate, and such a small cardiac mass that the ECG trace is difficult to analyse. However, it is still often worth applying ECG leads should an arrhythmia be detected on auscultation; or where cardiac disease is suspected. A
trace speed of 50mm per second should be used, preferably, with a 2cm to 1mV deflection, if possible.
What are the maintenance fluid rates for small mammals?
Maintenance values
Maintenance fluid rates for all small mammals are estimated at 80-100ml/kg/day i.e. ~ twice that estimated for cats and dogs. Because of their smaller size and higher metabolic rates, small mammals have higher glomerular filtration rates and greater
insensible losses.
How do you calculate fluid deficits for small mammals?
These may be calculated as for cats and dogs- however, it is worth noting that a lot of fluid intake, normally, is consumed as ‘food’ i.e. in the form of fresh vegetation. This is difficult to quantify but it is safer to assume that the debilitated small mammal will
not be eating sufficient to affect the fluid calculation.
As with cats and dogs, assume that 1% dehydration equates to a requirement of 10 mls per kilogram body weight fluid replacement, in addition to the maintenance requirements.
Additionally, if a blood sample can be taken, we can use the PCV and total protein levels to assess dehydration level. A table of average PCV’s and total proteins is given below, with a 1% increase in PCV suggesting 10 ml/kg fluid replacement is needed
What signs would be seen in small mammals with 3-5 % dehydration?
3-5% dehydrated: increased thirst, slight lethargy and tacky mucous membranes.
What signs would be seen in small mammals with 7-10% dehydration?
7-10% dehydrated: increased thirst leading to anorexia, dullness, tenting of the skin and slow return to normal, dry mucous membranes and ‘dull corneas’.
What signs would be seen in small mammals with 10-15 % dehydration?
10-15% dehydrated: dull/comatose, skin remains tented after pinching, desiccated mucous membranes.
What is the normal PCV and TP in a ferret?
Ferrets PCV L/L 0.44-0.60
TP - 51-74
What is the normal PCV and TP in a rabbit?
Rabbits PCV - 0.36-0.48
TP - 54-75
What is the normal PCV and TP in a Guinea pig?
Guinea-pigs PCV 0.37-0.48
TP -46-62
What is the normal PCV and TP in a chinchilla?
Chinchillas
PCV - 0.32-0.46
TP - 50-60
What is the normal PCV and TP in a rat?
Rats PCV - 0.36-0.48
TP - 56-76
What is the normal PCV and TP in mice?
Mice PCV - 0.39-0.49
TP - 35-72
What is the normal PCV and TP in a gerbil?
Gerbils PCV - 0.43-0.49
TP - 43-85
What is the normal PCV and TP in a hamster?
Hamsters Av PCV - 0.36-0.55
TP - Av 45-75
Over what time period should you administer rehydration fluids in small animals and why?
The calculated volume of fluids required to rehydrate the patient, is often too great to administer safely in one 24-hour period, due to their small size. The following compromise is advised with the deficit being replaced over 3 days. (This assumes that
there are no ongoing fluid losses requiring to be replaced).
Day one: Maintenance fluid levels + 50% of calculated dehydration factor.
Day two: Maintenance fluid levels + 25% of calculated dehydration factor.
Day three: Maintenance fluid levels + 25% of calculated dehydration factor.
If the dehydration levels are so severe that large volumes are indicated it may be necessary to take 72 hours to replace the calculated deficit rather than 48 hours
Fluids should be warmed to body temperature before use and fluid warming devices used where possible. Blood pressure monitoring can be used to assess the response to fluid resuscitation in hypovolaemic patients
What ongoing loss calculations should be considered in small animals that can vomit when considering a rehydration plan?
In addition, in those species which can vomit, such as ferrets, the volume of expelled vomitus should be considered. As with cats and dogs, assuming 2-4 ml/kg body weight per vomit is appropriate. In other species, where diarrhoea may occur but not vomiting e.g. small herbivores, it is much more difficult to make an estimate of fluid loss. It may approach 100-150 ml/kg body weight per day.
What choice of crystalloids are considered in small mammals?
Crystalloids
Lactated ringer’s solution (LRS), 0.9% saline and hypertonic saline (HTS) may all be used as with cats and dogs.
The fluid of choice for dehydration/ hypovolaemia is likely to be lactated ringer’s solution. HTS can be used in severe hypovolaemia to draw fluid into the vascular
compartment, if the patient is not dehydrated.
How can Protein amino acid/B vitamin supplements assist in small mammals as a fluid additive?
Protein amino acid/B vitamin supplements
These are useful for nutritional support e.g. Duphalyte® (Zoetis) at the rate of 1 ml/kg body weight/day (or added to fluids at a 20% level). They are particularly good to help replace some of the compounds needed for replenishment in situations where the patient is malnourished; or has a protein losing enteropathy, such as heavy parasitism; or a protein losing nephropathy. It is also a useful supplement for patients with hepatic disease or severe exudative skin diseases, such as heater burns.
How can bicarbonate assist in small mammals as a fluid additive?
Bicarbonate
This may be administered if metabolic acidosis exists, such as can occur in rabbits with hepatic lipidosis and secondary ketosis; and guinea pigs with pregnancy ketosis.
Calculations of replacements are as for cats and dogs.
How can electrolytes assist in small mammals as a fluid additive?
Electrolytes
Sodium and potassium levels should be monitored and corrected for deficits, where found. In practice, potassium levels will often be depressed in cases of long-standing diarrhoea and elevated in cases of renal disease. The use of calcium gluconate, when
a patient has a high potassium level, can help minimise hyperkalaemia associated arrhythmias.
How do you administer subcutaneous fluids in rabbits and how much can be administered?
Subcutaneous
Rabbits
The scruff area or lateral thorax make ideal sites. This is a good technique for routine post-operative administration of fluids for patients undergoing surgical procedures such as spaying or castration. It is possible to give a maximum of 30-60 ml/kg split into two or more sites, depending on the size of rabbit, at one time.
How do you administer subcutaneous fluids in ferrets?
Ferrets
This is an easily used route for post-operative fluids and management of mild dehydration in these species. The scruff area or lateral thorax are preferred sites.
How do you administer subcutaneous fluids in rodents and how much can be administered?
Rodents
The scruff area is easily utilised for volumes of 3-4 mls of fluids, for smaller rodents; and up to 10 mls at any one time for rats. The use of a 25-gauge needle is
recommended. However, the subcutaneous route may be painful for Guinea-pigs: but doses of 25-30 mls may be given at one time, preferably at two or more sites. It is important to be aware of fur slip when handling chinchillas
When would oral fluids be sufficient in rabbits that are hospitalised and how would they be administered?
Rabbits
Not such a good route for seriously debilitated animals but useful for those with nasooesophageal feeding tubes in place. It may be useful for mild cases of dehydration where owners wish to home treat their pet. This route, though, is restricted to small
volumes with a maximum administration of 10 ml/kg at any one time. In practice, it may be possible to administer much less than this.
When would oral fluids be sufficient in ferrets that are hospitalised and how would they be administered?
Ferrets
This route can be used as for rabbits. Naso-oesophageal/gastric tubes may be placed
as for cats; and doses of 10 ml/kg may be administered at any one time.
When would oral fluids be sufficient in rodents that are hospitalised and how would they be administered?
Rodents
Gavage tubes or avian straight crop tubes can be used to place fluids directly into the oesophagus. The rodent needs to be firmly scruffed to adequately restrain it and to keep the head and oesophagus in a straight line. This method is often stressful for the rodent: but the alternative is to syringe fluids into the mouth, which often does not work, as rodents can close off the back of the mouth with the cheek folds. This is a normal anatomical function, allowing rodents to gnaw structures without ingesting fragments of the material they are gnawing. Maximum volumes which can be given
via the oral route, in rodents, vary from 5-10 mls per kg body weight. Nasooesophageal/gastric tubes are not a viable option in rodents due to their small size.
How do you administer intraperitoneal fluids in rabbits and how much can be administered?
Rabbits
The rabbit needs to be positioned in dorsal recumbency with its head downwards, to allow the gut contents to fall out of the injection zone. The needle is inserted in the caudal right quadrant of the ventral abdomen. Once it is inserted just through the abdominal wall, drawing back on the plunger of the syringe checks for evidence of puncture of the bladder/gut. With correct positioning correctly, there should be no resistance to injection: a maximum volume of 20ml/kg may be given in one
administration. Concerns about concurrent respiratory/cardiovascular disease should
be considered when positioning and injecting.
How do you administer intraperitoneal fluids in ferrets and how much can be administered?
Ferrets
The same principles are applied as for rabbits and rodents. The patient should be placed in dorsal recumbency with its head downwards. The injection is administered in the right caudal-ventral quadrant with the bevel of the needle inserted just through the abdominal wall. The plunger should be pulled back to ensure that no puncture of the bladder/gut has occurred. Doses of 15-20 mls, at any one time, are generally well tolerated.
How do you administer intraperitoneal fluids in rodents and how much can be administered?
Rodents
Again, as for rabbits, the rodent should be placed in dorsal recumbency, with its head downwards, to encourage the gut contents to fall cranially and away from the injection site. The needle, preferably 25 gauge or smaller, is advanced slowly to just pop through the abdominal wall in the caudal right ventral quadrant. A maximum volume of 1-4 mls in smaller rodents and up to 10 mls in large rats may be given. This is a good route for more serious cases as intravenous fluids are not so well tolerated, particularly in chinchillas and Guinea-pigs.
What is the ideal intravenous route in rabbits and how can this be secured?
Rabbits
It is advisable for all sick rabbits or those undergoing anaesthesia to have an IV catheter placed. The marginal ear vein, cephalic vein and lateral saphenous vein can all be used for IV catheter placement. Complications of using the marginal ear vein
include necrosis sloughing of the ear tip or cartilaginous fracture in non-lop breeds especially after long-term use of the marginal ear vein (>2-3 days): this is relatively rare with careful venepuncture technique although some authors advise avoiding this vein. Sedation, with midazolam prior to placement, is advisable and application of a topical local anaesthetic cream is advantageous prior to placement of a 23-25 gauge Jelco ® catheters, or equivalent. If the catheter is placed in the ear vein, a rolled-up gauze swab or section of foam (e.g. cold water insulation foam) can be placed on the pinna with the catheter being taped in place around this to secure.
How do you administer intravenous fluids in ferrets?
Ferrets
Catheters are ideally placed in a sedated (midazolam) or anaesthetised patient. Ferrets frequently chew dressings off, therefore plenty of additional dressing material is required to allow sufficient time for the fluids to be administered. Care must be taken to avoid over administration (Orcutt, 2005) so fluids can be administered as boluses: if the patient is very debilitated a syringe driver could be used.
How do you administer intravenous fluids in rodents and how much can be administered?
Rodents
This route can be difficult especially in conscious animals. It is very difficult to place intravenous catheters in hamsters and gerbils, as they have few peripheral veins.
Using the tail veins in gerbils is dangerous due to the risk of causing tail separation, in this species. In mice and rats, an attempt could be made to use the lateral tail veins.
An intravenous bolus of fluids can be given by inserting a butterfly needle or using a 25 or 27-gauge insulin needle. Sedation, applying local anaesthetic cream and warming the tail to help dilate the vessels can make veni-puncture easier. Another option that may be considered is ‘cut-down’ jugular catheterisation, under GA. Volumes of 0.2 mls in mice and up to 0.5 mls in rats may be administered IV as a
bolus.
The cephalic and saphenous veins may be utilised in guinea pigs and chinchillas but as well as being very small and difficult to catheterise, this method is often poorly tolerated. Cut down over the vein, in an anaesthetised patient, is most likely to be effective. Where a catheter can be implanted, 25 or 27-gauge butterfly catheters or winged, over-the-needle catheters are required. In many cases the veins are so small, and the access so difficult that, other than the lateral tail veins, administration of a bolus of fluids, using a needle and syringe, is often all that can be achieved.
How do you administer intraosseous fluids in rabbits and how much can be administered?
Rabbits
Intraosseous needles may be placed into the proximal tibia or the proximal femur, in the trochanteric fossa, parallel to the long axis of the femur. An 18-23 gauge, 1-1.5 inch (2.5 – 3.7cm) spinal or hypodermic needle should be used. Strict aseptic preparation and analgesia is required when an intraosseous needle is placed. The veterinary surgeon may consider that prophylactic antibiosis e.g. Enrofloxacin (Baytril
® 2.5% Bayer) may be justified.
Intraosseous and intravenous fluid administration should be accurately delivered using an infusion pump or titrated using syringe drivers: if these are not available small volume boluses are preferred rather than relying on giving set gravity administration
(Fisher, 2010). Even a small error in fluid administration could be very significant considering the small size of many rabbits.
How do you administer intraosseous fluids in ferrets?
Ferrets
The proximal femur is the most easily accessed site. Access is via the natural fossa created by the hip joint and the greater trochanter as described below. Infusion devices, such as syringe drivers, are advised for this route of administration.
How do you administer intraosseous fluids in rodents?
Rodents
In larger rats, the proximal femur may be tolerated, as for rabbits but smaller species often have too small a medullary cavity for needles to be safely inserted. This is also the preferred route for severely dehydrated chinchillas and Guinea-pigs, with the proximal femur being the easiest site to access. Access is also via the natural fossa created by the hip joint and the greater trochanter (described below). Infusion devices
such as syringe drivers are advised for this route of administration
Describe the step by step process for the Placement of intraosseous catheters in small mammals?
Placement of intraosseous catheters in small mammals
1. Sedation or anaesthesia is required for all but unconscious or debilitated animals. Appropriate analgesia should also be provided.
2. The area overlying the lateral aspect of the hip joint (or proximal tibia) is clipped and surgically prepared. It is important that the whole technique is performed
aseptically.
3. A 20-22 gauge (for ferrets, rabbits and guinea pigs) or 24-26 gauge (for other rodents) spinal or hypodermic needle of appropriate length is insert in the fossa
between the hip joint and the greater trochanter parallel to the long axis of the femur.
4. The needle is flushed with heparinised saline (the advantage of a spinal needle is that it the central stylet prevents it from becoming plugged).
5. The needle is taped securely in place and antibiotic cream applied around the site. Ideally the area is X-rayed to ensure correct intramedullary placement of
the needle.
6. Once this has been confirmed, intravenous tubing is attached to the needle and kept securely in place by wrapping bandage material around it and the patients’
abdomen.
7. An Elizabethan collar or tubing guard is applied.
8. A syringe driver is used to ensure the correct volume of fluids are provided.
What are the clinical signs of early compensatory shock in small mammals?
Early (Compensatory)
Normal or increased blood pressure
Increased heart rate
Slightly depressed or normal mentation
What are the clinical signs of early decompensatory shock in small mammals?
Early decompensatory Increased heart rate Increased CRT Normal or decreased heart rate Normal or decreased blood pressure Depressed
What are the clinical signs of decompensatory shock in small mammals?
Decompensatory Decreased heart rate Decreased blood pressure Decreased / absent CRT Hypothermia Depression /comatose (the floppy bunny!)
What fluids may need to be considered in a small mammal that presents in shock?
If systemic blood pressure is lower than 90mmHg then the circulation can be supported using hypertonic saline and colloids. Initially a bolus of 3-5ml/kg of
hypertonic saline may be given slowly IV followed by 3-5ml/kg of colloid as a slow IV bolus. As with other species, larger volumes of isotonic, crystalloid fluids may be used to perform fluid resuscitation. If there is a poor response, further boluses may be carefully administered. Once normovolaemia has been achieved any other fluid deficits will need to be replaced.
The response is to fluid therapy is assessed by monitoring patient demeanour, behaviour, blood pressure and body temperature. Further investigation, including blood glucose, lactate, BUN, acid base status, electrolytes, PCV, TP and ECG, may be carried out.
What are the important considerations when performing a blood transfusion on a small mammal?
Blood Transfusions
Blood transfusions are sometimes performed as in other species e.g. if the PCV falls acutely below 20%. Blood transfusion has associated risks- the principles are similar to those for dog and cat transfusions. They
are best performed by direct, same species transfers (i.e. rat to rat, or Guinea-pig to Guinea-pig).
A healthy donor should be able to donate 1% of body weight (g) as blood, without any deleterious effects- this equates to ~ 10 ml/kg (Vennen and Mitchell, 2009)
E.g. 5 kg rabbit – 5000 g.
1% of 5000 = 50 ml.
Or 10 ml/kg= 10 x 5= 50 ml.
This may be increased to 4%, if required, although very careful monitoring of the donor is essential. There is very little information available currently about cross-matching blood groups of small mammals. Ferrets are not currently recognised to have appreciably detectable groups. They are severely anaemic if the PCV is < 30 % and candidates for blood transfusion if PCV < 20% (Mayer, 2006).
Rabbit blood volume is 55-65 ml/kg blood samples should ideally be cross-matched prior to transfusion.
If available, 5-6 mls of whole blood should be mixed with 1 ml of citrate acid dextrose anticoagulant; alternatively, the sample may be collected into directly into a preheparinised syringe for immediate transfusion. Intravenous catheters are advised for
blood transfusion as the rate of administration should be slow, especially initially: ~ 1 ml over 5-6 minutes if possible. It may be increased to a maximum rate of 6-12ml/kg/hour and must be complete within 4 hours. The recipient must be very wellrestrained and will ideally be sedated. Intraosseous donations may be made if vascular access is not possible. Oxyglobin®, if available, may be used at 4-10ml/kg replacement for severe deficits; and 1-2ml/kg replacement for minor losses
How do you perform a pulse assessment on a small mammal?
Pulse assessment
Femoral pulses may be detected by palpation in ferrets and rabbits, in the same location as cats and dogs. In small mammals, it is preferable to use a Doppler ultrasound monitor to assess the pulse and blood flow. This may be placed directly over the heart in small species; or over a significant vessel, such as the ventral tail artery, in ferrets and rabbits; the central ear artery, in rabbits; and the lateral tail vein, in mice or rats
How do you perform a cardiac and respiratory auscultation on a small mammal?
Cardiac and respiratory auscultation
Normal heart and respiratory rates are found in Table 1. Where consolidation has occurred, there may be no appreciable lung sounds- this is common in rabbits and
rats. However, most other small species with lung disease will have audible wheezes and crackles, as cats and dogs would. Bilateral auscultation of the lateral thorax, for the lungs, and immediately caudal to the shoulder, for the heart, is preferred. Rodents may have such a rapid heartbeat that counting the rate accurately may be impractical.
How do you perform blood pressure monitoring on a small mammal?
Blood pressure monitoring
This may be performed relatively easily via indirect methods e.g. Doppler or oscillometry. The cuff is applied to the forelimb, proximal to the elbow, with the Doppler probe placed over the palmar carpal area; or the cuff is applied to the hind limb, proximal to the tarsus, with the probe placed over the plantar metatarsal area. Four or five readings should be taken to ascertain an average blood pressure: this will give
systolic blood pressure figures (normal ~ 110-120mmHg).
How do you perform a neurological assessment on a small mammal?
Neurological assessment
In larger species, such as the rabbit, many of the reflex actions used in cats and dogs may be assessed: these include the withdrawal response, placement reflexes, flank needle prick (panniculus) reflex and righting reflexes. However, being a prey species, some rabbits and many of the smaller herbivores may exhibit freeze responses to manipulation, therefore rendering many of the tests invalid. Ferrets are difficult to
assess owing to their hyperactivity.
In most cases a simple assessment looking at the following points will give a good idea of the animal’s neurological function:
1) Can it stand upright?
2) Can it maintain this stance when gently pushed from either side?
3) Can it take food offered to it?
4) Does it have a head tilt?
5) Is there any evidence of nystagmus? (Note rabbits often show a whole head nystagmus, rather than an ocular one, as their eyes are so large they have little mobility)
6) Does the nystagmus worsen/appear/disappear when the head is tilted: suggesting a central versus peripheral nystagmus?
7) If obstacles are placed in front of the animal can it avoid them?
8) Is there a panniculus reflex; and if so is there any cut-off point, suggesting a spinal injury?
How do you perform pulse oximetry on a small mammal?
Pulse oximetry
Pulse oximeter probes may be applied to the same vessels as the Doppler probe.
Probes can be difficult to attach- the tongue may be useful in anaesthetised animals. In rabbits, other options include the pinna using the central ear artery; or the tail artery, ventral to the tail base.
In anaesthetised ferrets, the tongue may be used; or the tail artery, ventral to the tail base. In most rodents, the tail (position the probe laterally in mice and rats); ears (for Guinea pigs and chinchillas) or feet may be used.
Pulse oximetry assesses the oxygen saturation of haemoglobin (SaO2) rather than PaO2.The pulse oximeter readings show a logarithmic association with PaO2 readings meaning there are limitations to the assessment of readings i.e. a pulse oximeter
reading of 98% may indicate a PaO2 anywhere between 100mmHg and 500 mmHg.
100% on a pulse oximeter reading should equate to 500mmHg but it could be significantly less, which is a serious concern especially if the animal is breathing 100% oxygen. In cyanotic animals, pulse oximeter readings may be <70%, equating to PaO2 of 40-50 mmHg
How can kidney function be assessed in small mammals?
Kidney function may be assessed by measuring urea and creatinine levels in small mammals, but as with cats and dogs they are not a sensitive indicator of renal function.
There is no specific enzyme for liver damage in small mammals. ALT is of limited value in rabbits (
Why should blood glucose levels be monitored in rabbits and what could a high or low blood glucose indicate?
Blood glucose monitoring is important in all exotics, but most work has concentrated on rabbits. As with cats, rabbit blood glucose levels will rise if the patient is stressed, which can be due to pain and/or underlying pathological conditions, including dental
disease, arthritis, gut stasis, or renal /bladder stones. Remember that gut stasis will develop if the rabbit stops eating, either due to a primary GI problem or because they are in pain. Low blood glucose is abnormal and suggests that the rabbit, or other small
mammal, may be becoming shocked – so something needs to be done urgently; equally high blood glucose is abnormal, suggesting stress/pain and so equally
something needs to be done! It has also been suggested that these blood glucose levels could be used to differentiate gut stasis from impaction where very high blood glucose has been found in cases with
a blockage, requiring surgery. As with many conditions, this is not a hard and fast rule and serial values are of more value than a single reading; but suggests that further investigation is required as a blockage is high on the list of differentials.
Recent work by Harcourt-Brown & Harcourt-Brown (2012, 2014) suggests that blood glucose values in rabbits between 4.2 - 10 mmol/l are normal (although some laboratories consider blood glucose of 4.2-7.8 mmol/l as normal); values between 10 and 15 mmol/l suggest continuous monitoring is required; values between 15 and 20 mmol/l indicate a cause for concern, with effective analgesia being a priority; and
values above 20 mmol/l indicate that the rabbit’s condition is very serious- surgery may be required as there may be a gastrointestinal obstruction.
Why are female ferrets prone to anaemia?
Female ferrets are prone to anaemia, associated with persistent oestrus. Being induced ovulators, if kept alone or with another female/ castrated male
they will not ovulate and come out of oestrus. This is compounded by the fact they are very sensitive to the immunosuppressive effects of oestrogens. Bone marrow suppression is a common sequel to prolonged oestrous periods and can cause leucopenia, thrombocytopenia and aplastic anaemia
Why can rabbit urine sometimes appear cloudy?
Urine analysis
Rabbit urine can sometimes appear cloudy- rabbits absorb all the calcium present in their diet and excrete excess in the urine. This means that if they are on a high calcium diet (e.g. ad lib pellets), then the urine can take on a thick, white, pasty appearance- the worst cases can look like toothpaste due to calcium carbonate crystals. This is not normal and increases the risks of renal/bladder calculi.
What ongoing care, medication and monitoring should be carried out on a rabbit that has presented in an acute emergency?
Rabbits
Many rabbits presented as an acute emergency- they often already have gastrointestinal stasis (Girling, 2015) or are at high risk of developing it (Clark and
Saunders, 2012). The investigation and diagnostics are similar to other ECC cases (history, clinical examination, and diagnostics). The main differential that should be
ruled out is obstruction of the GI tract- radiography of the abdomen is valuable.
Affected rabbits should be hospitalised for management- as stress is a major cause of GI stasis it is important that they are housed away from dogs and cats and provided with a box to hide in.
Medical management includesAnalgesia (opioid +/- NSAID)
Fluid therapy
Assisted feeding
Prokinetics (if GI obstruction ruled out)- Ranitidine and cisapride act as prokinetics
along the whole length of the rabbit gut, whereas metoclopramide acts on the foregut.
Depending on the underlying cause antibiotics may be required. Enrofloxacin is licensed in the UK for rabbits and is effective against most Pasteurella spp. infections.
Other antibiotics that can be administered include trimethoprim sulfonamides and other fluoroquinolones. Penicillin and potentiated penicillins, amoxicillin and ampicillin, must never be given by mouth, but can be administered systemically, based on culture
and sensitivity results.
What ongoing care, medication and monitoring should be carried out on a ferret that has presented in an acute emergency?
Ferrets
There are several medications that may be administered depending on the disease process.
These could include:
furosemide, ACE inhibitors and digoxin for congestive heart failure
prednisolone or diazoxide for insulinoma
ranitidine and sucralfate for gastric ulceration. Chronic vomiting associated with gastric ulcers, either due to foreign bodies or Helicobacter mustelae infection, is common in ferrets. Ferrets rarely need prokinetics.
Antibiotics rarely pose a problem for ferrets, and most products suitable for cats can be used for ferrets. Foul tasting antibiotics (or other medications), such as oral
enrofloxacin (Baytril ® 2.5% solution, Bayer) liquid, should not be used in ferrets, as it will often result in severe scratching, with claw associated trauma to their tongue
What ongoing care, medication and monitoring should be carried out on a rodent that has presented in an acute emergency?
Rodents
Herbivorous rodents, such as guinea pigs and chinchillas, will often benefit from the use of prokinetics, as with rabbits. Dosages are similar. The antibiotics that are harmful to rabbits should also be avoided in small herbivores. It is possible to use fluoroquinolones and trimethoprim sulphonamides at rabbit doses safely. Enrofloxacin (Baytril ® 2.5% solution, Bayer) is licensed for use in all exotic sp. in the UK
How do you calculate the energy required for a debilitated rabbit?
Rabbits
The levels of energy required for a debilitated rabbit should approach that calculated for growing to lactating rabbits using the formula MER = k x (wt[kg])0.75, where k=200 for growth and 300 for lactation, (Carpenter and Kolmstetter 2000).
Therefore, for debilitation, the following daily energy requirement may be used.
MER = 250 x (wt [kg]) 0.75
To re-populate the intestinal flora, transfaunation of caecotrophs from a healthy rabbit may aid the return of normal bowel function (Kelleher, 2010). The use of commercial probiotics, designed for rabbits, has also been advocated; and reduces the risk of
transferring potential parasites and other agents to the debilitated patient.
How do you calculate the energy required for a debilitated ferret?
Ferrets
The same formula may be used for ferrets. The value used for k should be changed to 350.
formula MER = k x (wt[kg])0.75, where k=200 for growth and 300 for lactation
Therefore, for debilitation, the following daily energy requirement may be used.
MER = 250 x (wt [kg]) 0.75
How do you calculate the energy required for a debilitated rodent?
Rodents
The same formula may be used for rodents. The value of k will change depending on the species and has been estimated as 300 – 450.
formula MER = k x (wt[kg])0.75, where k=200 for growth and 300 for lactation.
Therefore, for debilitation, the following daily energy requirement may be used.
MER = 250 x (wt [kg]) 0.75
Describe the assisted feeding techniques for rabbits, the food available and how much?
Rabbits and other small herbivores
Assisted syringe feeding of a high fibre, herbivore Critical care diet (e.g. Oxbow Herbivore Critical Care, Burgess Critical Care, Science Recovery® (Supreme
Petfoods), or (Lafeber Emeraid® Herbivore) at a rate of 10-15ml/kg every 2-3 hours. If the rabbit is not willing to accept syringe feeding, a 5Fr nasogastric tube should be placed and a liquid diet e.g. Lafeber Emeraid® Herbivore delivered via tube (PaulMurphy, 2013). This can be a very effective way of providing nutrition to ill rabbits.
Describe the assisted feeding techniques for ferrets, the food available and how much?
Ferrets
A debilitated ferret may be supported with nasogastric or oral syringing of meat based baby food, or commercial critical care diets e.g. Lafeber Emeraid® Carnivore or Oxbow Carnivore™. If these are not available, liquid, meat based formulas, designed
for cats and dogs, such as Hills a/d™ could be used. The amount to be fed is ~10ml/kg three to six times daily. Care during administration is important to avoid aspiration pneumonia (Bament, 2013). It is vitally important that a debilitated ferret goes no longer than 4 hours without nutritional support, as they become hypoglycaemic rapidly
How would you place a naso-gastric feeding tube in a rabbit?
Placement of feeding tubes
Naso-gastric tube in rabbits
A 3-5 French gauge naso-gastric tube is pre-measured, with the head and neck extended, from the external nares to the last rib, and then lubricated. After spraying the external nares with lidocaine spray, it is introduced into the nasal cavity. After placement, a syringe is attached to the tube and aspirated- stomach contents should be seen, confirming correct placement. The tube may then be glued, taped or sutured to the dorsal aspect of the head, with the free end taped around an ear (using gauze or foam in the pinna, to prevent soft tissue compression). It is not normally necessary
to place an Elizabethan collar on the rabbit to prevent removal- this could cause significant stress
How would you place a naso-gastric feeding tube in a ferret?
Naso-gastric tube in ferrets
The technique is the same as for rabbits. An Elizabethan collar may be applied to prevent removal.
A 3-5 French gauge naso-gastric tube is pre-measured, with the head and neck extended, from the external nares to the last rib, and then lubricated. After spraying the external nares with lidocaine spray, it is introduced into the nasal cavity. After placement, a syringe is attached to the tube and aspirated- stomach contents should be seen, confirming correct placement. The tube may then be glued, taped or sutured to the dorsal aspect of the head, with the free end taped around an ear (using gauze or foam in the pinna, to prevent soft tissue compression). It is not normally necessary
to place an Elizabethan collar on the rabbit to prevent removal- this could cause significant stress
How would you place a pharyngostomy feeding tube in a ferret and why would you consider placing it?
Pharyngostomy tube in ferrets
This could be used in ferrets where oral trauma prevents normal prehension and mastication of food. The same technique can be used as for cats using an 8-10 French tube. Butterfly tape is applied around the tube, at the neck, to suture the tube to the skin. This can then be bandaged in place.
What type of bacteria do small herbivores often have in wounds?
small herbivores wounds often have more Pasteurella and Streptococcal Spp. Rabbits also tend to develop walled-off abscesses in response to infection, meaning that flushing and debriding wounds in this species is often unrewarding. Instead, surgical excision of infected abscessed areas is often required.
What type of anaesthesia is preferable for most rodents?
For most rodents, gaseous anaesthesia with sevoflurane or isoflurane is preferable
What type of anaesthesia is most commonly used in healthy ferrets?
In healthy, non-cardiovascularly compromised ferrets, triple combination (Ketamine + Medetomidine +Butorphanol) may be administered IM. The medetomidine may be reversed with atipamezole. A veterinary formulary should be consulted for the
appropriate dose.
What anaesthesia protocol is often used in healthy rabbits?
For rabbits, medetomidine (or midazolam) plus buprenorphine can be administered IM. Once sedated, induction is achieved with IV Ketamine to effect (diluted with sterile water to facilitate accurate incremental dosing). A veterinary formulary should be
consulted for the appropriate dose which should be selected by a veterinary surgeon.
The rabbit is then intubated and anaesthesia is maintained with isoflurane and oxygen
What anaesthesia protocol is often used in debilitated small mammals?
Debilitated small mammals
In most cases where the animal is severely debilitated, sedation with midazolam maybe preferable to anaesthesia. If anaesthesia is essential, or the patient is only mildly debilitated, then midazolam sedation followed by gaseous anaesthesia with sevoflurane or isoflurane in 100% oxygen should ideally be used. Sevoflurane is preferred in small mammals as they are adversely affected by the taste of isoflurane.
Gaseous induction of rodents is easy- they may be placed in an induction chamber (preferably) or their head placed in a face-mask. Pre-oxygenation for 3-4 minutes, followed by a gradual increase in concentration to 3-4% sevoflurane (2-3% isoflurane)
is used for induction. 2-3% sevoflurane (1-2% isoflurane) is used to maintain anaesthesia via a mask or ET tube
Ferrets are easily induced in a chamber but care must be taken with mask induction, as they will object, wriggle and bite if possible.
There are various anaesthetic techniques for rabbits and guinea-pigs. If gaseous induction is performed they should be sedated (e.g. midazolam) first. Chamber
induction is far preferable to manual restraint. The rabbit is given 100% oxygen for 2 minutes. If still breathing, 1% sevoflorane (0.5% isoflurane) is added and the breathing monitored for another 2 minutes. If still breathing, then 2% Sevoflurane (1% isoflurane)
is administered for a further 2 minutes and so-on increasing in 0.5% increments until anaesthesia is induced at around 2-3% Sevoflurane (1.5-2% isoflurane). If the rabbit stops breathing at any point the volatile agent should be reduced by 0.5% and
breathing usually restarts. Alternatively, the rabbit can be intubated and the volatile agent to given to effect.
What type of pain scoring charts have been developed for small mammals?
. Grimace scales have been developed for rabbits, rats and mice as ‘facial expression provide a reliable and rapid means of assessing pain in mice, rabbits and
rats’
When should meloxicam be avoided in rabbits?
The standard dose of meloxicam in rabbits is higher than that used in cats and dogs but the same care should be taken when using NSAIDs in rabbits i.e. they should be avoided where renal disease or gastro/intestinal ulceration/perforation is suspected.
How does the body temperature of birds differ to small mammals?
the body temperature of all birds is higher than mammals- all are similar at around 41-42ºC.
Do birds get pyrexia?
Birds do not seem to show a physiological pyrexia in the presence of infection.
How do you perform an initial assessmenton a bird?
Do not be too quick to physically restrain the sick avian patient, particularly if it is a small species and/or in respiratory distress. Severe respiratory distress is obvious, with stertor, wheezes and whistles: but subtle changes, such as increased respiratory rate, tail bobbing and nasal discharge/blocked nares may all indicate relatively serious respiratory pathology. Birds have no diaphragm and therefore rely on the outward
movement of the ribcage and downward movement of the keel to allow inspiration.
Any restriction of this (coupled with the increased oxygen demand of a stress response) will lead to hypoxia, possible cardiac ischaemia and arrest.
Examine the bird in the cage first, using dimmed blue or red lighting- much can be learned by observing the bird before handling it. If there is respiratory distress or
suspicion of respiratory disease, the bird should be placed in a warm, dark, oxygen enriched atmosphere (+/- nebulisation with saline) for 15 minutes before any
examination. Following this, it may be considered that sedating (e.g. midazolam IM) or anaesthetising the patient (100% oxygen with isoflurane) is the preferred method for a physical examination. N.B. Owners should be warned of the risks of handling a sick bird- its condition may worsen or it could die.
The initial assessment should consider the following points. NOTE the observations and questions that should be asked are like those for small mammals.
1. What is the feather condition – chewed feathers, fret lines, discolouration,
feather loss etc?
2. Are the feathers fluffed up?
3. Are the eyes lemon shaped or closed?
4. Are the corneas bright?
5. Is there increased respiratory noise?
6. Is there tail bobbing when breathing indicating dyspnoea/hyperpnoea?
7. Are the nares clear or is there a serous or purulent discharge?
8. Is there any faecal clumping to the feathers around the vent?
9. What is the recent faecal output like?
10.Are the urates green/mustard yellow? (may indicate liver disease)
11.Is there any blood in the faeces (may indicate urolithiasis, cloacitis, intussusception, urogenital (UGT) disease or haematuria which can be associated with heavy metal poisoning in Amazons)?
12.Is there undigested seed in the faeces? (May indicate psittacine pro-ventricular dilatation disease or pancreatitis).
13.What is the bird’s weight AND what is the muscle condition of the pectoral
region like (compare with normal for species)? (Body condition scores of 0-5 may be given as with other animals). All birds should be weighed.
14.What is the bird’s stance? (Upright on the perch, on the floor, leg weakness, respiratory distress etc.)
15.Is there evidence of vomitus/regurgitation on the feathers around the head?
What should you observe when doing an initial assessment on a bird?
The initial assessment should consider the following points. NOTE the observations and questions that should be asked are like those for small mammals.
1. What is the feather condition – chewed feathers, fret lines, discolouration, feather loss etc?
2. Are the feathers fluffed up?
3. Are the eyes lemon shaped or closed?
4. Are the corneas bright?
5. Is there increased respiratory noise?
6. Is there tail bobbing when breathing indicating dyspnoea/hyperpnoea?
7. Are the nares clear or is there a serous or purulent discharge?
8. Is there any faecal clumping to the feathers around the vent?
9. What is the recent faecal output like?
10.Are the urates green/mustard yellow? (may indicate liver disease)
11.Is there any blood in the faeces (may indicate urolithiasis, cloacitis, intussusception, urogenital (UGT) disease or haematuria which can be associated with heavy metal poisoning in Amazons)?
12.Is there undigested seed in the faeces? (May indicate psittacine pro-ventricular dilatation disease or pancreatitis).
13.What is the bird’s weight AND what is the muscle condition of the pectoral
region like (compare with normal for species)? (Body condition scores of 0-5 may be given as with other animals). All birds should be weighed.
14.What is the bird’s stance? (Upright on the perch, on the floor, leg weakness, respiratory distress etc.)
15.Is there evidence of vomitus/regurgitation on the feathers around the head?
What would a detailed examination of a bird involve?
Detailed Physical Examination
A physical examination is often carried out under sedation or anaesthesia and will include the following:
1. An intra-oral examination using a mouth gag or a pen/pencil to encourage the bird to open its beak. This should allow a close examination of the tongue, the
choana (slit in the dorsal mouth - the communication between the nasal passages and the mouth) and the palate. The glottis may also be visualised.
Abnormalities such as a foreign body or discharge from the choanal slit, wide choana or loss of papillae on the choana (often indicative of vitamin A deficiency), an abnormal or foul odour and evidence of white or yellow plaques on the mucosa should all be noted. If possible a sample should be obtained using a swab, dampened with sterile saline, for microscopic examination for
Trichomonas (motile protozoa) spp.; and cytological examination of a Diff Quick stained examination for evidence of fungal or bacterial infection.
2. A detailed examination of the nares and the eyes will allow an assessment for upper respiratory tract disease. Clinical signs include abnormal shaped nare(s); rhinoliths (hard concretion of exudate in the nares); loss of the rostral concha (e) (operculum); sinking of the globe of the eye; swelling above or below the globe (the region of the infraorbital sinus); discharge from the eye; swelling of the conjunctiva and corneal blemishes. The ear canal should also be examined (caudal and ventral to the lateral canthus).
3. A detailed examination of the feathers, particularly the blood or newly emerged ‘pin’ feathers. These may be plucked from the body area and the sheath
carefully slit to reveal the pulp, which may then be smeared and stained for microscopic examination. This can give evidence of skin or feather pulp
infections. Examination of the shaft and vane of the feather may also reveal evidence of lice and mites. Fret bars or lines (lines of weakness, perpendicular
to the direction of the feather vein) indicate illness, stress (or steroid administration) at the time that that feather was developing.
4. A detailed auscultation of the lungs and air sacs. The lungs are best auscultated from the dorsum, between the wings, as they are closely attached to the ventral
aspect of the thoracic area. The air sacs are dispersed throughout the body.
The most easily auscultated and those most likely to have pathological changes are the abdominal and caudal thoracic air sacs- these may be auscultated on
the lateral aspect of the body caudally, just behind the keel bone. The heart may be auscultated from the lateral body wall just underneath the wings.
5. A detailed examination of the wings may be made. When anaesthetised, an idea of wing integrity and the elasticity of the propatagium (wing web) may be
made by placing the bird on its sternum and extending both wings laterally, to an equal distance, and comparing the tension, which should be similar. Any
asymmetry should be noted as it may indicate joint or propatagial injury.
6. A detailed examination of the preen gland (dorsal body just cranial to insertion of tail feathers – absent in Amazon parrots and pigeons), vent and caudal
abdomen should also be made. The caudal abdomen, unprotected by the keel bone, should be naturally concave. Any convexity may indicate breeding
activity, a space occupying mass in the coelomic cavity, organomegaly (such as hepatomegaly), constipation or ascites. In smaller birds (<300gm) slight wetting the feathers over the ventrum, in this region, will allow the clinician to see through the thin skin of the body wall. If hepatomegaly is present, the dark black-brown shadow of the liver will be seen: normally the liver does not extend caudal to the keel bone
How do you check a birds airway and breathing in the ABC emergency protocol?
Airway and breathing
Pollock (2007) discussed the approach to the dyspnoeic avian patient (including video footage). Air-sac tube placement
In some cases of severe respiratory distress, such as syringeal aspergillosis, foreign body aspiration or when performing tracheal washes in a dyspnoeic bird, the placement of an air-sac tube may be vital to maintain a patent airway. Brown and Pilny (2007) describe the placement technique.
1. The anaesthetised bird should be placed in right lateral recumbency.
2. The uppermost wing (left) is abducted dorsally and attached to a wedge or similar. The left leg is abducted caudally and fixed in position with tape.
3. A small skin incision is made between the last rib and penultimate rib at a level 2/5th of the depth from top to bottom.
4. Either a specifically designed air-sac tube or a short length of 2-3.5 gauge endotracheal tubing, held by haemostats, is passed through the body wall. The cuff on the endotracheal tube or air sac tube may
be inflated and the tube sutured to the body wall.
5. The outlet of the anaesthetic circuit may then be attached to the free end of the tube to administer the anaesthetic. The oxygen flow rate should be reduced to 300ml/kg/minute - even so breathing may cease
due to the effect of CO2 wash out. No further action should be required, as oxygen is passing over the absorptive surface- respiration should restart when the anaesthetic is stopped or oxygen is removed.
Alternatively, in the conscious bird, the air sac tube can be cut short and left in place to support breathing for 3-4 days whilst its condition is treated.
The air-sac tube works so well because of the unique physiology and anatomy of the avian airways – the tube is placed in the caudal thoracic air sac, at a point before inspired air passes over the absorptive surface
How do you check a birds cardiovascular system, in the ABC emergency protocol?
The prognosis for respiratory arrest caused by anaesthetic overdose is good. In this situation, respiratory arrest usually occurs 2-3 minutes prior to cardiac arrest- so, if the bird is carefully monitored, the volatile agent is reduced and IPPV is administered (one breath every 5 seconds) a positive outcome is expected.
However, cardiac arrest in birds carries a worse prognosis than for mammals. This is, partly, due to the difficulty in performing cardiac compressions due to the presence of the keel/sternum of the bird. Most birds become bradycardic immediately before arrest- if this is recognised, rapid administration of epinephrine/adrenaline/adrenaline and atropine can be effective. Atropine premedication should always be used when surgery of the head or neck is involved. Epinephrine/adrenaline/adrenaline may be given by the intravenous, intraosseous or, often more effectively, the intra-tracheal route.
How do you perform an ECG on a bird?
ECG
ECG traces may be taken from birds, although it is preferable to anaesthetise the bird to perform them. It is also helpful to blunt the alligator teeth on the clamps, as these can traumatise the skin very easily. The bird is placed in dorsal recumbency and the clips are attached to the skin of the thighs and the skin of the propatagium (the skin between the shoulder and carpus on the wing’s leading edge). Alternatively, hypodermic needles can be placed through the skin of the wings and thighs and clamps attached to these.
In most avian species, the lead II trace is like a mammalian one, except the QRS complex appears inverted. The complex is not actually inverted- it occurs because the S wave is the dominant deflection with the Q wave hardly being recorded at all. For this reason, bird QRS waves are often referred to as ‘RS’ waves. P and T waves are as for mammals. Occasionally in some species (pigeons and some parrots) there is a small depression wave known as a Ta wave immediately after the P wave- this is normal (it represents atrial repolarisation). In addition, the P on T phenomena (where the P wave is superimposed onto the following T wave) is a normal finding in some African grey and Amazon parrots.
Normal lead II annotated trace for pigeons and some psittacine birds showing the Ta wave associated with atrial repolarisation
Normal values for bird ECGs have been published but they vary between species and exceed the scope of this text. Many birds have heart rates which are too fast to record even on 50mm/second traces. It is more important to look for arrhythmias, such as severe bradycardia instead- this is a common abnormality before cardiac arrest in anaesthetised birds.
what are the fluid maintainance value for a bird?
Maintenance values
If a bird is assessed as being dehydrated, the volume of fluids required to rehydrate can be calculated. Maintenance for birds is assumed to be 50 ml/kg/day. The fluid deficits is calculated as for other species by assessing the percentage dehydration.
What signs may you see in a bird that is 3-5% dehydrated?
3-5% dehydrated: increased thirst, slight lethargy, tacky mucous membranes, increased heart rate
What signs may you see in a bird that is 7-10% dehydrated?
7-10% dehydrated: increased thirst leading to anorexia, dullness, tenting of the skin and slower return to normal over eyelid or foot, dry mucous membranes, dull corneas, red or wrinkled skin in chicks. Basilic vein (vein which runs over the ventral aspect of the elbow joint of the wing-often used for blood sampling) refill time exceeds 2 seconds
What signs may you see in a bird that is 12-18% dehydrated?
12-18% dehydrated: dull-comatose, skin remains tented after pinching, desiccated mucous membranes, sunken eyes.
How much replacement fluids are needed for a 1% increase in PCV suggest?
Performing a PCV and total protein assessment will give more detailed information on hydration status. 1% increase in PCV suggests 10 ml/kg of replacement fluids are needed
What time period are replacement fluids given over in birds?
As with small mammals, the volume that is required to replace the deficit may be too large to administer over one day – hence the deficit may be calculated and administered over the first 3 days.
Day one Maintenance fluid levels + 50% of calculated dehydration factor.
Day two Maintenance fluid levels + 25% of calculated dehydration factor.
Day three Maintenance fluid levels + 25% of calculated dehydration factor.
What choice of crystalloids can be used in birds?
Crystalloids
Lactated Ringer’s is the fluid of choice in most cases although normal saline may also be indicated. Once the fluid deficit has been corrected, ongoing maintenance fluid therapy, hypotonic glucose saline combinations may be used.
When might colloids be used in birds and what is the dose?
Colloids
These may be used, as with mammals, although their use is currently controversial. Bolus administration of gelatins, such as Gelofusine® and Haemaccel®, may be given to aid in the treatment of low colloid osmotic pressure associated with hypoproteinaemia. The infusion rate is 10-15 mls/kg intravenously 4 times over a 24-hour period. Hetastarch® may be used at a similar rate.
When might potassium be added to fluids in birds?
Potassium
May be added to fluids where hypokalaemia is present. This is useful in cases of chronic diarrhoea or thermal burns.
When might bicarbonate be added to fluids in birds?
Bicarbonate
May be given at rates of 1 mEq/kg up to a maximum of 4mEq/kg without measuring bicarbonate deficits where metabolic acidosis is suspected.
What type of diet do
parrots, cockatiels and budgies
pigeons and poultry
raptors, carnivores and piscivores
require when considered crop feeding?
Oral
Crop tubing
Once rehydrated, critical care diets appropriate to the species are very useful:
herbivore for parrots, cockatiels and budgies
omnivore for pigeons and poultry
carnivore for raptors, carnivores and piscivores (fish eating birds)
What is the maximum volumes of fluid that may be crop-tubed in a budgerigar?
0.5-1 ml
What is the maximum volumes of fluid that may be crop-tubed in a cockatiel?
2.5-5 mls
What is the maximum volumes of fluid that may be crop-tubed in a conure?
5-7 mls
What is the maximum volumes of fluid that may be crop-tubed in a cockatoo?
10 mls
What is the maximum volumes of fluid that may be crop-tubed in an african grey?
8-10 mls
What is the maximum volumes of fluid that may be crop-tubed in a macaw?
15 - 20 mls
What is The optimal subcutaneous site in birds?
Subcutaneous
For mildly dehydrated birds, and for follow-up maintenance fluids, the subcutaneous route may be used. The optimal subcutaneous site is between the scapulae, i.e. dorsally between the wings.
why is intracoelomic/intraperitoneal fluids contraindicated in birds?
Intracoelomic
Intracoelomic/intraperitoneal fluids are CONTRAINDICATED IN BIRDS due to the absence of a diaphragm and the presence of air sacs.
What are the preferred veins to use in birds?
It is advised that all sick birds and those undergoing anaesthesia, lasting more than 10 minutes, should have an indwelling I/V catheter placed. The preferred veins are the superficial ulna vein, ventral to elbow, the medial tarsal vein or the right jugular vein. In waterfowl or long legged birds (not parrots) the medial metatarsal vein can be used. Catheters are typically placed under GA.
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in finches?
0.5ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in budgies?
1ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in cockatiels?
2ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in conures?
6ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in amazon parrots?
8ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in barn/tawny owls?
10ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in cockatoos?
14ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in buzzards?
12-14ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in macaws?
14ml
What are the maximum volume of fluids that may be administered safely by slow intravenous bolus injection in swans?
25-30ml
What bones are not suitabel for intraosseous catheters in birds?
The following bones should NOT be used for intraosseous fluid therapy as in many species they are pneumonised (connected to the air sacs and lungs): humerus; femur; vertebrae; sternum and pelvis. Administration of fluids via this route would result in drowning.
What do you use as an intraosseous catheter in birds and how should it be placed?
If vascular access is not possible, a hypodermic needle or a spinal needle, in larger species (>200g), may be placed in the distal ulna or the proximal tibio-tarsus and intraosseous fluids may be administered slowly. Again, if the bird is conscious and struggling, it is preferable to do this procedure under gaseous anaesthesia. In collapsed and comatose patients anaesthesia may not be required.
How are blood transfusions performed in birds?
Blood transfusions
These are indicated when the PCV has dropped below 20%- they may be given via intravenous or intraosseous routes. Prior blood-typing and cross-matching of samples, does not appear to be necessary for one off transfusions. The transfusion should, however, involve the same genus e.g. pigeon to pigeon; African Grey Parrot to African Grey Parrot etc. Donor blood is collected into a pre-heparinised syringe. In healthy birds the volume of blood collected can be up to 2% of the donor’s body weight. Administration of the blood, into the recipient, should be performed over a 60-120 minute period.
If a donor is not available, then 3-10ml/kg of Oxyglobin® may be used (Oxyglobin® is currently unavailable).
How and where can pulse assessments be carried out on birds?
Pulse assessment
Assessing a pulse can be very difficult in birds owing to their rapid heart rates and thin walled vessels, which can be easily occluded. In most cases, pulse assessment is made using Doppler ultrasound probes attached to one of the following blood vessels:
1) Superficial ulna vein (aka basilic vein)- runs over the ventral aspect of the elbow joint of the wing
2) Medial tarsal vein- runs over the medial aspect of the lower un-feathered leg and may be clearly seen in waterfowl
What should the capillary refill time be on birds and where can this be assessed?
The capillary refill time should be less than 2 seconds as with mammals. This may be assessed on mucous membranes around the mouth, face or vent.
Where can the lungs be auscultated in birds and why is thi sless informative in birds than in mammals?
Auscultation is less informative in birds than in mammals. To auscultate the lungs, the stethoscope diaphragm must be placed over the dorsum of the bird, as the lungs are closely adherent to the underside of the dorsal thoracic wall. Air sounds may be heard over the flanks and ventrum but these are due to referred sounds and the sounds of air moving through the air sacs (balloon like structures inside the bird). With rigid lungs and having no diaphragm, the bird inspires by moving its keel bone ventrally and ribcage laterally. This creates negative pressure inside the single body cavity (coelom) which allows air to be drawn in through the trachea down to the lungs. Gaseous exchange occurs in the lungs, which, being semi-rigid in nature, do not inflate or deflate significantly. Due to the lack of inflation, air already in the lungs moves caudally into the air sacs. On expiration, the process is reversed, meaning that air in the caudal air-sacs is pushed back through the lungs, allowing extraction of oxygen on both inspiration and expiration. The air then goes into the cranial air sacs. From entering the trachea to its final expulsion, an air molecule is involved in two cycles of inspiration and expiration.
Where can the heart be auscultated in birds?
The heart may be auscultated over the back or pectoral area. The heart rate is usually very fast, even in the larger parrots so, detecting murmurs is difficult. Arrhythmias however may be detected.
What can fitting and collapse in birds be caused by?
Fitting or collapse in birds could be caused by the following:
heavy metal poisoning (particularly lead but also zinc)
hypocalcaemia (particularly in a hen bird that is egg laying); any psittacine species, fed on a seed based diet, may also be affected (especially common in the African Grey Parrot which has a recognised hypocalcaemia syndrome)
hypoglycaemia (particularly common in birds of prey that are underweight and being flown regularly)
hyperglycaemia in raptors of excitable species (e.g. goshawk) which are being stressed
Chlamydiosis or Psittacine Proventricular Dilation Syndrome in a percentage of affected psittacine birds
cranial trauma
CNS infection of parasitic (usually protozoal), fungal or bacterial origin
How do you assess a birds ability to perch?
The bird should be assessed for ability to perch using both legs. When held in the hand, the bird should be checked to see if it can grasp a finger/towel with both feet. Unilateral leg paresis through to full paralysis is common, particularly in small psittacine birds such as the budgerigar, and is often associated with renal or gonadal neoplasia. The gonads are located internally in both males and female birds. Neoplasia can put pressure, bilaterally on the sciatic nerves, as they pass under the middle lobe of the kidney. Similar clinical signs may be seen in toxicities such as heavy metal (lead or zinc) poisoning; or organic toxins such as those associated with aspergillosis infection of the airways.
What is paresis affecting one or both legs in young poultry typically due to?
In young poultry (usually ~ 3-4 months of age), paresis affecting one or both legs is typically due to Marek’s disease, a viral disease for which there is no treatment. The patient should be euthanased. Vaccination, which must be administered before the bird is three days of age, may prevent this condition.
What are the signs of lead poisoning in birds and how might this come about?
Birds of prey may develop lead poisoning having consumed lead shot in prey that they have been fed. They will often sit on the floor, rocked back onto their ‘hocks’, with drooped wings and a slightly drunken appearance. In birds <1kg, lead will be evident on radiographs: whilst in birds >1kg, the lead has often been regurgitated in their casting and, hence, will not be apparent on radiographs. Blood collected into an EDTA tube, can be used to detect elevated plasma lead levels.
What is the dropping of both wings in a bird most likely to be due to?
Both wings drooping is more likely to be lead, permethrin or organophosphate poisoning
How do you perform a neurological exam on a bird?
The bird should be assessed to see if it holding both wings normally. One wing drooping is most likely to be associated with a muscular/skeletal injury. Both wings drooping is more likely to be lead, permethrin or organophosphate poisoning. In normal birds, physiological nystagmus can be induced by moving the head horizontally from side to side. If this is absent, Cranial Nerve VIII may be affected. Birds will not normally demonstrate pathological nystagmus as the eyes are held in place by the bony ossicles in the sclera- instead they move the whole head. Head tilts are common with peripheral and central vestibular disease. Pupillary reflex responses are not consistent in birds, as they have skeletal muscle in their irises, over which they have a degree of conscious control. Also, the eyes are so big and the midline bone, separating them, is often so thin that light shone into one eye frequently passes through the bony septum into the opposite eye as well.
The bird should be assessed for ability to perch using both legs. When held in the hand, the bird should be checked to see if it can grasp a finger/towel with both feet. Unilateral leg paresis through to full paralysis is common, particularly in small psittacine birds such as the budgerigar, and is often associated with renal or gonadal neoplasia. The gonads are located internally in both males and female birds. Neoplasia can put pressure, bilaterally on the sciatic nerves, as they pass under the middle lobe of the kidney. Similar clinical signs may be seen in toxicities such as heavy metal (lead or zinc) poisoning; or organic toxins such as those associated with aspergillosis infection of the airways.
How do you perform pulse oximetry on a bird?
Pulse oximeter probes may be attached to the distal legs, over the medial metatarsal area or over the basilic (brachial) vein, on the ventral aspect of the elbow. It should be noted, however, that avian haemoglobin is different from mammalian and therefore the accuracy of the oximeter readings is questionable. However, changes in the SpO2 are relevant and should be acted upon, even though the actual values (providing the readings stay above 90%) are not reliable
What is the only useful indicator of renal function in birds?
It should be noted that, as with reptiles, uric acid is the only useful indicator of renal function, although usually more than three quarters of kidney function needs to be lost before uric acid levels become elevated. Urea and creatinine levels do not provide information on renal function in birds and reptiles.
What three main conditions in parrots will elevate the white cell count?
In parrots, three main conditions will elevate the white cell count over 20x109/L.:
1) Psittacosis (infection with Chlamydophila psittaci)
2) Aspergillosis
3) Egg yolk coelomitis (equivalent to peritonitis where the yolk ruptures internally instead of being shed into the oviduct)
4) Mycobacteriosis (rare)
in general what is the range of white blood cells in birds?
In general, white cell counts are in the range of 5-9x109/L.
What is the avian equivalent of the neutraphil? What does evidence of degranulation and rupture of the cells indicate?
The avian equivalent of the neutrophil is the heterophil- so named because of its part basophilic/ part eosinophilic staining with Romanowsky stains. It has a polymorphic, bi/trilobed nucleus and brick red, cigar-shaped granules in its cytoplasm. On a blood smear, evidence of degranulation and rupture of the cells (toxicity) is a strong indicator of active and severe infection.
Why can’t erythrocytes and platelets in birds be counted in an automated cell counter?
Erythrocytes and platelets in birds are nucleated and hence cannot be counted in an automated cell counter. Erythrocytes are rugby ball shaped with a similar shaped nucleus; platelets are oval and smaller and other cells are like mammals.
How does normal excrement appear in birds?
Normal excrement of birds comprises, faeces (green, brown or black), urates (white) and an aqueous fraction.
How do you assess a birds urine?
This is less useful than for mammals owing to the faecal contamination of avian urine that occurs prior to production. It is, however, important to look at the urates (white portion of the dropping) to see if there is any blood; or if they are discoloured (mustard yellow or lime green)
What may mustard yellow or lime green urate in birds indicate?
If the latter has occurred, this is evidence of biliverdinuria which, in birds, is an indicator of liver damage / inflammation/ (commonly associated with psittacosis in parrots; and lead poisoning in birds of prey). Biliverdin is the main excretory product of the liver, as opposed to bilirubin in mammals.
What is blood in urates in amazon parrots often associated with?
In Amazon parrots, only, blood in the urates is often associated with heavy metal poisoning, particularly lead.
If the faeces are well formed, but the excrement is excessively wet in birds what does this indicate?
If the faeces are well formed, but the excrement is excessively wet, this indicates PUPD, rather than diarrhoea, and is an important differentiation to make
What type of birds are ongoing nutritional supplementation with calcium/vitamin D3 replacers useful in ?
Ongoing nutritional supplementation with calcium/vitamin D3 replacers are useful in cases of African Grey Parrot hypocalcaemia: examples include Nutrobal® and Zolcal-D® produced by Vetark Professional Ltd. Being a liquid preparation Zolcal-D® is preferred as it allows for accurate dosing. Other mineral and vitamin supplements are often required if the diet is poor or restricted to seeds, as may be the case with parrots. In addition to introducing fruit and vegetables, vitamin A, particularly, may need to be supplemented in these parrots.
How do you calculate energy requirments in birds?
Critical care nutrition- including calculation of energy requirements
The estimated daily food requirements may be calculated using the Emeraid® calculator on www.lafebervet.com. The calculation is based on species, patient weight and body condition. As a starting point, 2% of the bird’s bodyweight is administered by gavage, 3 times per day (birds over 500g) or 4 times per day for smaller birds. The patient should always be weighed each morning, prior to food or fluid administration: if it is failing to maintain its weight, the volume or frequency of feeding must be increased accordingly.
How do you provide assisted feeding to a bird?
Assisted feeding techniques and foods
Most cage birds are granivorous, herbivorous or frugivorous. Members of the parrot family should not be fed meat or dairy products as this could result in renal, hepatic and cardiovascular disease. The nurse/clinician must never wait for a sick (especially small) bird to feed themselves. Once hypovolaemia and hypothermia is addressed, supplementary feeding should be commenced. A scientifically and commercially produced, appropriate, exotic animal critical care diet, should be given e.g. Lafeber Emeraid® (Herbivore, Omnivore, Carnivore or Piscivore). Assisted feeding involves using a purpose designed, steel, blunt ended crop tube (for psittacine birds). A plastic or rubber tube, which is attached to a syringe containing the food formula, may be used in poultry, waterfowl, pigeons or raptors. In psittacine birds, the beak is opened and the metal tube passed down. In other species, the index finger of the left hand (restraining the head) is used as a gag, by placement in the cheek tissue on the left side. See section headed ‘Crop Tubing’ for maximum values that may be crop-tubed at one time. In the absence of an appropriate food, a suitable liquidised baby food, (avoid products containing lactose) such as Cow and Gate or Milupa or a suitable bird hand rearing food may be administered by crop tube.
What are the important considerations when performing asisted feeding in raptors and what should be avoided?
Raptors are by nature carnivorous and when sick should be fed a meat based diet. Whilst sick, fur or feather (casting material) should be withheld so that feeding can be repeated several times a day. This avoids having to wait for the bird to ‘cast’ (bring up their casting) which occurs, typically, 10-18 hours after the normal daily meal. Products such as Hills a/d®, Oxbow Carnivore Care™ or Lafeber Emeraid® Carnivore are suitable.
What care do you provide to a bird with a superficial wound?
Avian wounds generally gape due to their skin being very thin and inelastic. Skin wounds must be covered with hydrocolloid gels, dressings or sutured closed to prevent desiccation. An Elizabethan collar may need to be applied, especially in parrots, to prevent self-trauma. A foam neck extension collar, of appropriate length and internal diameter such that it doesn’t cause distress, (fashioned from cold water insulation foam) is fitted. If necessary a horizontal plastic Elizabethan collar may be applied- to avoid chewing, this must extend beyond the lateral reach of the beak. Birds are typically very distressed after initial collar application. Hospitalisation is recommended in these cases allowing for midazolam sedation whilst the bird becomes accustomed to the collar.
What care do you provide to a bird with a deep wound?
For deeper wounds, associated with e.g. bites, debridement of the wound under anaesthetic is essential, particularly if they occur over the distal legs. This helps to prevent infection of the avascular tendons or, worse, osteomyelitis. Once cleaned the wounds may be dressed with Granuflex®, to aid granulation; or Intrasite Gel® (or similar) and covered with Allevyn® or Allevyn Thin® (or similar). Secondary and tertiary bandage material may be applied over the top to give the bird something to chew before it reaches the wound. Collars are preferable, as a chewing parrot will readily chew through its own limb or digits.
How do you stabilise a wing fracture?
For stabilising wing fractures, a figure of eight bandage may be used for a short period (<48hrs). This contracts the wing, binding it to the lateral body wall but should never be tight over the propatagium. Any longer immobilisation leads to fibrosis of the joints and will result in severe impairment of wing function.
How do you stabilise a leg fracture in a bird?
Leg fractures, in small cage birds such as canaries, may be stabilised using the so-called ‘Altman’ splint. This is simply two pieces of heavy zinc tape applied laterally and medially to the fractured leg, sandwiching the leg between them. These can be left on, or replaced as the bird chews them off, for the duration of the healing process. Finger splints or tongue depressors may be used in larger birds, for temporary stabilisation, but these will generally require surgical fixation.
When is ingluviotomy tube placment in birds useful?
Ingluviotomy tubes (i.e. placed via the crop into the proventriculus) are invaluable in birds where gavage is impossible; or in highly trained birds, who will permit tube feeding without restraint, but may be highly stressed by physical restraint.
When are air sac tubes indicated in birds? where and how are they placed?
Air sac tubes (equivalent to a tracheostomy tube in a mammal) facilitate breathing where there is compromised tracheal breathing e.g. tracheal blockage due to a foreign body or a fungal granuloma. The granuloma is usually Aspergillus spp. which like to grow at the bottom of the trachea where it divides into the primary bronchi. An aperture is created between the last and penultimate rib on the left side (2/5th way from top to bottom) and a small tube placed via this hole into the caudal thoracic air sac, to allow the bird to breathe.
What colour lighting aids bird restraint and why?
Apart from owls, birds are always best caught and restrained using dim, red or blue lighting- the bird can be seen but it cannot see the handler, thus decreasing the stress.
How do you manually restrain parrots?
For parrots the head is firmly caught and held between thumb, first and second finger, thereby controlling the beak (the most dangerous part of the parrot): the rest of the bird is then wrapped in a towel.
How do you manually restrain a small bird?
Small cage birds may be carefully restrained using a small hand-towel or piece of kitchen roll/paper roll in a latex gloved hand. The paper provides a larger surface area and means the bird is less likely to fly around the cage to avoid it. Once caught, the paper may be dropped and the bird held in one hand grip, where the head is gently caught between thumb, first and second finger and the rest of the body is cupped in the palm, to prevent wing flapping and damage
How do you manually restrain raptors?
For raptors, the bird is grasped with a towel over its back, in two hands. The legs are controlled, on each side, between the third and fourth finger. The bird is held between elbow and chest, with the legs restrained by the other hand. Once restrained, the bird’s legs are grasped between the fingers of one hand, always with one finger between the two legs, to prevent self-trauma if it struggles. A towel may be draped over the bird’s head to prevent biting. A second person may then examine each part of the body. In all cases, care should be taken not to tighten the grip and put too much pressure on to the chest/body. Birds have no diaphragm and rely solely on the outward movement of the body wall to allow inspiration to occur.
Why is chemical restraint in birds used?
Chemical restraint is not used to immobilise birds, but instead to reduce the stress to the bird for physical examination, collection of clinical pathology samples, radiography etc.
What are the options of chemical restraint in birds?
Isoflurane
sedation (midazolam IM)
parenteral anaesthesia e.g. ketamine mixed with medetomidine and administered IM, or preferably IV.
What isoflurane levels are used to chemically restrain a bird?
Isoflurane- induction for manual restraint 4-5%, reduced to 2.5-3% for maintenance (an experienced avian anaesthetist should be present to monitor and adjust depth of anaesthesia)
How do you provide parenteral anaesthesia in a bird for chemical restraint?
parenteral anaesthesia e.g. ketamine mixed with medetomidine and administered IM, or preferably IV. Weighing the bird is essential to ensure accurate dosing. This combination should give ~ 25 minutes of surgical anaesthesia. The bird should be intubated and maintained on oxygen. The medetomidine dose can be repeated once if necessary and it should be reversed with the same volume of atipamazole (IM). Using this regime, the bird should be standing and normal ten minutes later.
Why is intubation always advised in birds over a standard face mask?
Standard face masks may be used for parrots and raptors; or masks adapted from plastic bottles for long-beaked birds. Once anaesthetised, intubation is always advised as this allows intermittent positive pressure ventilation (IPPV) should the patient stop breathing: it also protects the airway and permits scavenging of exhaled volatile gases. The glottis (there is no epiglottis) is clearly visible at the base of the tongue in all birds: although in parrots, the tongue needs to be pulled forward, with atraumatic forceps, to visualise it.
Why is it not advisable to inflate a cuff on an ET tube on an intubated bird?
It is NOT advisable to inflate the cuff on the ET tube as birds have complete tracheal cartilage rings: over-inflation of the cuff can lead to damage to the cartilage rings and subsequent tracheal stricture formation. In addition, it is best to position the patient in ventral or lateral recumbency where possible: as dorsal recumbency will result in a 10-60% reduction in tidal volume. If dorsal recumbency is required, then manual or automatic intermittent positive pressure ventilation (IPPV) may be required.
Why should fluids be commenced in any reptile that is sick?
Renal failure is a common sequel of many disease processes in reptiles: it is vitally important, whenever treating a sick reptile, to maintain hydration status to encourage tubular urine flow. In any sick reptile, unless there is evidence to the contrary, it is assumed that the patient is 10% dehydrated (i.e. 100ml/kg deficiency). Fluids should be commenced immediately at 10-15ml/kg BID. The aim is to provide maintenance fluids (15-30ml/kg) and replace the deficit at the following rate: 50% day one, 25% day two, 25% day three). There are several factors, relating to medications that need consideration
What are the maintenance fluid rates in reptiles and in what time frame should replacement fluids be given over?
the aim is to provide maintenance fluids (15-30ml/kg) and replace the deficit at the following rate: 50% day one, 25% day two, 25% day three)
What drugs have nephrotoxic effects in reptiles?
Several drugs used in reptile medicine, such as aminoglycoside, potentiated sulphonamide and tetracycline antibiotics, have potentially serious nephrotoxic effects. These drugs should be avoided altogether when severe dehydration is present. Even when well hydrated, a reptile receiving potentially nephrotoxic medications should always have supplemental fluid therapy, to reduce the risk of drug associated renal damage.
what is the renal portal system in birds and reptiles and why is this thought to occur?
Reptiles and birds possess a unique venous blood return from the caudal half of the body, including the bone marrow, subcutaneous tissues, musculature of the hind-limbs, lumbar region and tail. This venous blood can bypass glomerular filtration to perfuse the renal tubular area- this is called the renal portal system (WikiVet, 2013). It is thought to be an adaptation for water retention, to prevent ischaemic tubular damage when glomerular blood-flow is reduced. It is thought that depending on hydration status, blood may pass through this system; or bypass it and pass directly into the systemic circulation.
Where should drugs ideally be administered in reptiles in accordance to thier renal portal system?
Fluid therapy and many drugs that are not nephrotoxic can be safely injected into the caudal half of the body and the renal portal system does not need to be considered. However, drugs that are potentially damaging to the renal tubules, and drugs which are removed by renal tubular excretion should ideally be administered into the cranial part of the body to avoid any ‘first pass’ consequences of this portal system. Recent research in healthy snakes and chelonians, however, has found that choosing cranial or caudal injection site appears to make no difference to the distribution or pharmacologic parameters of the drugs tested. In the presence of marked dehydration, however, there could still possibly be damaging effects.
How do you perform an initial assessment on a reptile that presents in an emergency?
Initial assessment
An initial assessment should be made of the reptile patient before it is removed from its carry cage/box. This should focus on the following points:
1) Is it potentially hazardous/dangerous to the handler? (e.g. male green iguanas, snapping turtles, aggressive snakes and, unlikely but possible, venomous snakes)
2) Is it mouth breathing and, therefore, possibly in respiratory distress?
3) Is it a fragile species? (many geckos will shed their tails very easily, as will many iguanas)
4) Is it possibly suffering from metabolic bone disease making it a risk to handle? (Look for deformed limbs, shell, spine etc. Inability to support its own weight, in a lizard or chelonian, may suggest this condition is present).
5) How large is the animal? (Many larger species of tortoise are surprisingly heavy and strong; snakes longer than 3-4 feet require more than one handler to avoid damaging the patient and putting the handlers at risk).
During the initial examination of a reptile that presents in an emergency, what question can be asked to the owner regarding the husbandry?
Whilst examining the patient from a distance to ascertain if it is safe to handle, it is a good idea to question the owner about the husbandry of the reptile at home e.g.
1) What do they feed it?
2) Do they feed vitamin/mineral supplements?
3) Is there a UV lamp? (may not be necessary for most snakes but is necessary for most lizards and chelonia – required in all species which naturally bask in sun light)
4) What temperature range do they keep the vivarium at (refer to a reliable website for correct data (e.g. www.anapsid.org or www.lafebervet.com)
5) What hides/cage furniture is present in the tank?
6) What humidity is the vivarium kept at?
7) Are there any other reptiles/pets?
8) If a snake, when did it last shed its skin and was it a complete shed?
How do you perform a detailed examination of a reptile?
1) An intra-oral examination can be performed using a mouth gag, tongue depressor/pen/pencil to encourage the reptile to open its mouth. A metal, hard plastic or wooden gag should never be used on a reptile. If they bite on this, it can cause damage to the jaw or teeth, especially if they have infection or metabolic bone disease. Care should be taken with chelonia as they have powerful jaws. It should be possible to perform a close examination of the tongue, the roof of the mouth and the nasal passages. There is no hard palate in reptiles, other than crocodylia. The glottis may also be visualised at the base of the tongue. Abnormalities such as a discharge from the nasal passages; petechiae or haemorrhages in the mouth; an abnormal or foul odour and evidence of white or yellow plaques on the mucosa should all be noted. Samples should be obtained for cytology and culture using a swab dampened with sterile water. It should be noted that many lizards have a two-coloured tongue e.g. the green iguana has a bright red tongue tip and a pale pink body to the fleshy tongue.
2) A detailed examination of the nares, ears and eyes will allow an assessment for upper respiratory tract disease. Clinical signs include: abnormal shaped nares; sinking of the globe of the eye; swelling below the globe of the eye (the region of the infra-orbital sinus); clouding of the cornea (an indication of impending shedding); discharge from the eye; swelling of the conjunctiva and corneal blemishes. Swollen, weeping or painful eyes, are often an indication of a retained shed (spectacle)- a common finding in snakes and lizards. The ear canal is located caudal and ventral to the lateral aspect of the eye. Ascending infection via the eustachian tube commonly results in abscessation. This is seen as distension of the skin overlying the ear canal. If present, this will require to be lanced, drained, cleaned and samples taken for culture and sensitivity.
3) A detailed examination of the skin/shell. This may show areas of retained slough: snakes should shed (ecdysis) their skin in one piece (including the eye-caps) about once per month; lizards shed in small patches and chelonia in small patches from the limbs and head/neck/tail. Dysecdysis (incomplete or improper shed) has many potential causes including disease, poor nutrition and/ or husbandry (Pollock, 2013). Geckos and small lizards often have retained shed on their toes, so these should be checked carefully- this will also allow any petechiae or ecchymoses to be observed, a possible indicator of septicaemia. These are often seen on the plastron (underneath shell of a tortoise). Abscesses appear as firm, inspissated subcutaneous masses.
4) A detailed auscultation of the lungs and air sacs. The lungs are best auscultated from the dorsum in chelonia and lizards. To improve sound conductivity, a damp towel/cloth may be placed over the reptile and the diaphragm of the stethoscope applied to this. Snakes are difficult to auscultate owing to their long thin lungs.
5) The heart is very difficult to auscultate and it is often preferable to use a Doppler probe to assess blood flow through/out of the heart to determine heart rate. It should be noted that the heart rate and heart sounds of reptiles are significantly different from mammals, owing to the different anatomy and physiology of the three-chambered heart (one ventricle and two atria). In addition, environmental temperatures will significantly alter heart rates.
6) A detailed examination of the limbs should be made. The long bones should be palpated as metabolic bone disease is common. Fibrous dystrophy, where the poorly ossified bone swells due to cartilage deposition, makes the limb look fat and muscular. Palpation reveals, however, that it is bone mass and not muscle. Shells of chelonia may be deformed and soft to touch. The mandibles of lizards may be bowed and malleable with this condition.
7) A detailed examination of the vent and caudal coelom should also be made. Many lizards have kidneys tucked into the pelvic area: so, these should not be palpable cranial to the wings of the ilia in normal animals. Snakes may be palpated, by running a finger along the ventrum, to feel for masses or obstructions. Chelonia are obviously difficult to palpate, although gentle palpation or ballottement of eggs or masses, by placing a finger cranial to a hind limb and rolling the animal onto its side and away again is possible. Snakes may be sexed by probing the cloaca in a caudal direction: a sulcus of 3-4 scales denotes a female, 6-7 scales length a male. Male lizards have hemipenes, bilateral grasping organs, used in copulation. Prolapse and infection is not uncommon. Cloacal prolapses are common in reptiles: a diagnostic work up of the cause of excessive straining is essential. Replacement and purse string suture is, however, not appropriate.
How do you assess the airway and breathing in a reptile that presents in an emergency?
Airway and breathing
The stimulus for respiration in the reptile is not elevated PaCO2 levels, as with mammals, but lowered PaO2. ‘Hypoxia stimulates respiration in reptiles and supplemental oxygen can actually suppress voluntary respiration, reducing effective oxygenation and making a bad situation even worse’ (Pollock, 2014a). This means that if 100% oxygen is administered to a reptile, it will, in effect, not need to breathe of its own accord as the PaO2 will be adequate. On recovery from GA, room air should always be administered via the Ambu bag rather than oxygen.
Why should antibiotic administration be considered in sick reptiles and why?
It should be noted that most sick reptiles are borderline or fully septicaemic. They tend to be attacked by their own gut bacteria, which are generally Gram negative in nature (often the Salmonella and Pseudomonad bacterial family). Strict hygiene must be adhered to when nursing reptiles with appropriate consideration of the zoonotic risks. A bacteriocidal antibiotic, active against Gram negative bacteria, is likely to be required- i.e. the fluoroquinolones and third generation cephalosporins. In accordance with the veterinary prescribing cascade, the only antibiotic licenced for reptiles in the UK, is enrofloxacin as Baytril 2.5%® (Bayer plc)- this should, in theory, be usedinitially. However, it is appropriate for other antibiotics to be used instead, if culture and sensitivity testing indicates this.
If carrying out CPR on a rabbit, at what rate should the compressions be performed?
Select one:
a. 75 - 100 bpm
b. 130- 160 bpm
c. 50 - 70 bpm
d. 100 - 120 bpm
The correct answer is: 100 - 120 bpm
Maintenance fluid requirements for most small mammals are:
Select one:
a. 40-50ml/kg/day
b. 20-30ml/kg/day
c. 80-100ml/kg/day
d. 60-70ml/kg/day
The correct answer is: 80-100ml/kg/day
What volume of fluid is required to replace the deficit, only, if a rabbit is considered to be 7% dehydrated?
Select one:
a. 0.7ml/kg
b. 7ml/kg
c. 700ml/kg
d. 70ml/kg
The correct answer is: 70ml/kg
The maximum volume of fluids which should be given orally to a rabbit is:
Select one:
a. 7ml/kg
b. 1ml/kg
c. 5ml/kg
d. 10ml/kg
The correct answer is: 10ml/kg
The maximum total volume of fluids which could be administered by the intraperitoneal route to a a ferret is:
(N.B. Need to be aware of who can perform this legally in the UK)
Select one:
a. 25-30ml
b. 5-10ml
c. 15-20ml
d. 35-40ml
The correct answer is: 15-20ml
Which of the following has prokinetic gut activity in a rabbit?
Select one:
a. Metoclopramide
b. Ranitidine
c. Cisapride
d. All of the answers
The correct answer is: Right lateral recumbancy
How should a bird be positioned for air sac tube placement?
Select one:
a. Left lateral recumbancy
b. Right lateral recumbancy
c. Dorsal recumbancy
d. Ventral recumbancy
The correct answer is: Right lateral recumbancy
The average maintenance fluid value for a bird is:
Select one:
a. 25ml/kg/day
b. 100ml/kg/day
c. 50ml/kg/day
d. 75ml/kg/day
The correct answer is: 50ml/kg/day
What is the maximum volume of food which should be given to an African Grey parrot via crop tube?
Select one:
a. 4-6ml
b. 2-4ml
c. 6-8ml
d. 8-10ml
The correct answer is: 8-10ml
Choose four clinical signs found in rabbits under each section header, signifying whether they indicate dehydration or shock
The correct answer is:
Choose four clinical signs found in rabbits under each section header, signifying whether they indicate dehydration or shock.
Dehydration:
[Tacky mucous membranes][Increased thirst][Skin tenting][Dull corneas]
Shock:
[Increased heart rate][Decreased blood pressure][Hypothermia][Increased CRT]
The minimum recommended fluid maintenance rate for a domestic rabbit is:
Select one:
a. 25ml/kg/24 hours
b. 40ml/kg/24 hours
c. 80ml/kg/24 hours
d. 100ml/kg/24 hours
The correct answer is 80ml/kg/24 hours. (100ml/kg/hr is the maximum amount)
The correct answer is: 80ml/kg/24 hours
Which small mammal has six incisors?
Select one:
a. The chinchilla
b. The rabbit
c. The guinea pig
d. The rat
The correct answer is: The rabbit
Which of the following could not be used to assess the pulse and blood flow, using Doppler, in a ferret?
Select one:
a. heart
b. auricular artery
c. femoral artery
d. ventral tail artery
The correct answer is: auricular artery
Atropine premedication should be administered to a bird undergoing surgery of the:
Select one:
a. leg
b. wing
c. head
d. cloaca
The correct answer is: head
Intraosseous fluid may be administered to birds via the:
Select one:
a. ulna
b. humerus
c. pelvis
d. femur
The correct answer is: ulna
Respiration is stimulated in reptiles by:
Select one:
a. elevated PaO2
b. lowered PaO2
c. elevated PaCO2
d. lowered PaCO2
The correct answer is: elevated PaCO2
Which one of the following can be used for intraosseous fluid administration in both lizards and tortoises?
Select one:
a. proximal tibia
b. distal femur
c. proximal femur
d. plastron-carapacial junction
The correct answer is: proximal tibia
Refeeding syndrome in a snake results in life-threatening:
Select one:
a. hypocalcaemia
b. hyperkalaemia
c. hyperglycaemia
d. hypophosphataemia
The correct answer is: hypophosphataemia
Most reptiles have a PCV of 30%. The veterinary surgeon may start to consider a blood transfusion once the PCV has reached:
Select one:
a. 10%
b. 20%
c. 25%
d. 15%
The correct answer is: 10%
The maximum recommended fluid maintenance rate for a reptile is:
Select one:
a. 20ml/kg/24 hours
b. 30ml/kg/24 hours
c. 50ml/kg/24 hours
d. 70ml/kg/24 hours
The correct answer is: 30ml/kg/24 hours
What Nutritional and husbandry diseases are common in reptiles?
Nutritional and husbandry diseases are common in reptiles including metabolic bone disease and hypocalcaemic tetany in egg bound mature lizards, such as the Green iguana. Emergency management may require administration of calcium gluconate; and possibly diazepam or midazolam for lizards having seizures
What problem may Garter and water snakes get that have been fed salt water fish that has previously be forezen?
Garter and water snakes fed salt water fish that has been previously frozen may suffer from a relative deficiency of vitamin B1 (thiamine): this can lead to a neurological condition (like cerebro-cortical necrosis in grain gorged cattle) which manifests as an inability to right itself and continual star gazing. If administered promptly, injections of vitamin B1 may be effective.
What treatment may be given to reptiles with Cardiovascular and respiratory disease that may have lead to pneumonia or pulmonary oedema?
Cardiovascular and respiratory disease are relatively common and may lead to pneumonia or pulmonary oedema. Diuretics such as furosemide and hydrochlorothiazide may be required. Oxygen therapy can be used but care should be taken as the stimulus for breathing in reptiles is lowered PaO2 rather than an elevated PaCO2, unlike mammals. Providing 100% oxygen, for even short periods of time, can mean the reptile stops breathing. Conscious intubation and a brief period of IPPV can be performed in collapsed reptiles- they are relatively easy to intubate and lack a cough reflex as they have no diaphragm.
What should be done if cardiac arrest occurs in a reptile?
If cardiac arrest occurs, intubation should be attempted followed by intratracheal administration of epinephrine/adrenaline/ adrenaline. Reptiles can cope with a degree of hypoxia beyond that tolerated by mammals. IPPV after intubation is essential- if this is not possible chest massage in lizards; and moving limbs in and out of the shell of chelonia may be successful at aiding air movement in and out of the lungs. Pollock (2012) states ‘the reptile heart rate varies with body temperature and will lower as body temperature falls. At lower temperatures, cardiac output is typically maintained by an increase in stroke volume, however it is prudent to never declare a reptile dead until it is WARM and dead’.
How should fluid be prepared prior to administration to a reptile?
Reptiles should be warm to maximise the absorption of fluids and the fluids should be warmed for administration 37.8-38.9°C (100–102°F) (Johnson 2011, cited by Pollock, 2011). Fluids can be administered by the IV or IO routes using a syringe driver or fluid pump that can deliver very small volumes as fluid overload is a risk.
What are the maintenance fluid value for reptiles?
These may be calculated as for cats and dogs. It is worth noting that a lot of the fluid intake is normally consumed as ‘food’ i.e. in the form of fresh vegetation for herbivorous species. This volume is difficult to quantify, therefore, it is safer to assume that the debilitated reptile will not be eating sufficient for it to affect the fluid calculation.
There is relatively little research on rates of fluid replacement. Consequently, for the vast number of reptiles and amphibians a calculated guess should be made. Frye (1991) recommends that levels of 20-25 ml/kg body weight per day be used for hydration purposes in both reptiles and amphibians; current literature suggests that rates vary from 10-50 ml/kg/day or 10-30 ml/kg/day across several species.
Wh at are the signs of excessive fluid administration in a reptile?
There is however another restriction to fluid rates of administration: as most fluids are given intracoelomically in the debilitated reptile although intravenous and intraosseous routes may also be used. Pollock (2014 b) describes catheter placement for delivery of fluids to reptiles. Reptiles do not possess true diaphragms, with the thorax and abdomen being interconnected as a common cavity or ‘coelom’. Administering fluids in this cavity is the equivalent of intra-peritoneal fluids in a mammal but as there is no diaphragm the fluid can pressure on the lungs, preventing their expansion. Excessive fluid administration may severely compromise respiration.
Excessive fluids may also:
overload the circulation
create pulmonary oedema
result in cardiac and renal over perfusion
cause solute wash-out with potassium excretion. This diuresis may cause a hypokalaemic crisis to develop, manifesting initially as an anorectic reptile but progressing to cardiac arrhythmias, coma and death.
How do you assess hydration in a reptile?
As with cats and dogs, it is possible to assume that 1% dehydration equates to a requirement for 10 ml per kilogram body weight fluid replacement, in addition to the maintenance requirements. It is also possible to make some qualitative assessment of the level of dehydration from the elasticity of the skin. Although reptile skin is not as elastic as mammalian, it should still be freely mobile and recoil, albeit slowly, after tenting. Other factors to assess are the brightness of the corneas in species with mobile eyelids. In those without mobile eyelids (e.g. snakes) collapse of the spectacle (the clear fused eyelids) is suggestive of dehydration. Other assessments of thirst and urate output can be made over a 24-hour period.
Assumptions must be made about the degree of dehydration of the reptile concerned based on clinical findings.
As a rough guide:
3% dehydrated – increased thirst, slight lethargy, decreased urates.
7% dehydrated – increased thirst leading to anorexia, dullness, tenting of the skin and slow return to normal, dull corneas, loss of turgor of spectacles in snakes
10% dehydrated – dull, comatose, skin remains tented after pinching, desiccated mucous membranes, sunken eyeballs, no urate/urine output.
What signs would indicate a 3% dehydration in a reptile?
3% dehydrated – increased thirst, slight lethargy, decreased urates.
What signs would indicate a 7% dehydration in a reptile?
7% dehydrated – increased thirst leading to anorexia, dullness, tenting of the skin and slow return to normal, dull corneas, loss of turgor of spectacles in snakes
What signs would indicate a 10% dehydration in a reptile?
10% dehydrated – dull, comatose, skin remains tented after pinching, desiccated mucous membranes, sunken eyeballs, no urate/urine output.
How much replacement fluids are needed in a reptile with a 1% increase in PCV?
The alternative is to compare packed cell volumes and total protein levels to assess dehydration: with 1% increase in PCV suggesting 10 ml/kg of replacement fluids are needed.
What is the PCV and TP of a green iguana?
Green iguana (Iguana iguana)
PCV l/l
0.25-0.38
Total Protein g/l
28-69
What is the PCV and TP of a Testudo spp. Tortoise?
PCV l/l
0.19-0.4
Total Protein g/l
32-50
What is the PCV and TP of a Ratsnake (Elaphe spp.)?
PCV l/l
0.2-0.3
Total Protein g/l
30-60
What is the PCV and TP of a Boa constrictor (Boa constrictor)?
PCV l/l
0.2-0.32
Total Protein g/l
46-60
When calculating fluid requirements for a reptile, What rate it is important not to administer more than and why?
When calculating fluid requirements, it is important not to administer more than 25-30 ml/kg/ daily for the reasons mentioned before. Therefore, rehydration of severely debilitated reptiles may take several days to weeks. As with avian patients the fluid deficit may need to be split over several days.
There is however another restriction to fluid rates of administration: as most fluids are given intracoelomically in the debilitated reptile although intravenous and intraosseous routes may also be used. Pollock (2014 b) describes catheter placement for delivery of fluids to reptiles. Reptiles do not possess true diaphragms, with the thorax and abdomen being interconnected as a common cavity or ‘coelom’. Administering fluids in this cavity is the equivalent of intra-peritoneal fluids in a mammal but as there is no diaphragm the fluid can pressure on the lungs, preventing their expansion. Excessive fluid administration may severely compromise respiration.
Excessive fluids may also:
overload the circulation
create pulmonary oedema
result in cardiac and renal over perfusion
cause solute wash-out with potassium excretion. This diuresis may cause a hypokalaemic crisis to develop, manifesting initially as an anorectic reptile but progressing to cardiac arrhythmias, coma and death.
What crystalloid fluids are suitable for reptiles?
Crystalloids
Lactated Ringer’s/Hartmanns
As with cats and dogs, this isotonic fluid is useful as a general-purpose rehydration/maintenance fluid. It is particularly useful for reptiles suffering from metabolic acidosis, such as those with chronic gastro-intestinal problems, but can also be used for fluid therapy after routine surgical procedures
There is some evidence (Rudloff, 2005) that in reptiles, and probably amphibians, the isotonicity of the extracellular fluids is lower than that of mammals. Studies in non-marine reptiles suggests that isotonicity for most reptiles is 0.8% rather than 0.9% as assumed for mammals. To this end several fluid combinations based on the above have been proposed.
Two types of crystalloid support have been suggested:
1. One third 5% glucose with 0.9% saline; one third lactated Ringer’s solution; one third sterile water.
2. Nine parts 5% glucose with 0.9% saline to 1 part sterile water.
Many texts, however, still advise that undiluted lactated Ringer’s solution or 4% glucose with 0.18% saline may be used. It is of course important that the fluids administered are warmed to the reptile/amphibian’s preferred body temperature (approximately 30-35˚C) before being given.
Why are saline glucose combination fluids useful for reptiles?
Glucose/Saline Combinations
These are useful for reptiles and amphibians who have often been anorexic for a period, therefore borderline hypoglycaemic, prior to treatment and. In addition, these are the fluids of choice for reptiles with renal disease and elevated potassium levels
There is some evidence (Rudloff, 2005) that in reptiles, and probably amphibians, the isotonicity of the extracellular fluids is lower than that of mammals. Studies in non-marine reptiles suggests that isotonicity for most reptiles is 0.8% rather than 0.9% as assumed for mammals. To this end several fluid combinations based on the above have been proposed.
Two types of crystalloid support have been suggested:
1. One third 5% glucose with 0.9% saline; one third lactated Ringer’s solution; one third sterile water.
2. Nine parts 5% glucose with 0.9% saline to 1 part sterile water.
Many texts, however, still advise that undiluted lactated Ringer’s solution or 4% glucose with 0.18% saline may be used. It is of course important that the fluids administered are warmed to the reptile/amphibian’s preferred body temperature (approximately 30-35˚C) before being given.
When would colloids be indicated in reptiles?
Colloids
These have been used in reptilian practice when direct venous or intra-osseous access has been achievable (Rudloff, 2005). They may be used as with cats and dogs, for serious hypovolaemia e.g. serious blood loss of blood occurs to maintain perfusion and support central blood pressure. This may be the only means of supporting such a patient or a temporary measure whilst a blood donor is found
When are Protein amino acid/B vitamin supplements indicated in reptiles?
Protein amino acid/B vitamin supplements
These are useful for nutritional support e.g. Duphalyte® (Zoetis) at the rate of 20% of fluid administered. They are particularly good to help replace some of the compounds needed for replenishment in cases where the patient is malnourished; has a protein losing enteropathy (often due to heavy parasitism); or a protein losing nephropathy. They are also a useful supplement for patients with hepatic disease or severe exudative skin diseases, such as heater burns.
When would oral administration of fluid be useful in reptiles and how can this be carried out safely?
Oral
This may be used in reptile and amphibian practice for those patients experiencing mild dehydration, and for ‘home’ administration. Many products are available for cats and dogs that may be used for reptiles. As with crystalloid fluids, it is advisable to dilute these oral electrolytes by approximately 10%, otherwise their concentration will be greater than the reptile’s extracellular fluid and so water will move from the body into the gastrointestinal tract. One electrolyte solution which may be useful is Repto Boost® by Vetark Professional Ltd. This may be given orally by gavage, or placed in bathing water, so that it is absorbed per cloaca. Warm water baths are a good way of encouraging mildly debilitated reptiles to drink. Even if the reptile does not drink the water, some may be absorbed via the cloaca and so mild levels of dehydration can be reversed if this technique is performed for 15-20 minutes daily. It is important to ensure that the water is not so deep as to place the reptile in danger of drowning.
When would enteral fluids via a pharyngostomy tube be indicated in a snake and how can this be placed?
Enteral fluids are not the best choice for seriously debilitated snakes but are useful for those with pharyngostomy feeding tubes in place; or if the owner/handler is experienced in stomach tubing. They may be a useful choice for mild cases of dehydration where owners wish to home treat their pet. A stomach tube is easily passed by restraining the snake’s head gently, but firmly, and then inserting a rubber/soft plastic/wooden tongue depressor to open the mouth. A lubricated feeding tube is then passed through the labial notch (the area at the most rostral aspect of the mouth without teeth) down to one third of the snake’s length. This route, however, is not advisable for severely dehydrated snakes and those with pre-existing gut pathology, due to the poor rate of fluid uptake from the GIT.
How do you tube feed a lizard?
Lizards
Pollock (2011a) describes feeding the hospitalised lizard.
Gavage tubes, avian straight crop tubes or feeding tubes can be used to place fluids directly into the oesophagus or stomach of lizards. The reptile needs to be firmly restrained to keep the head and oesophagus in a straight line. The mouth is opened with a plastic or wooden tongue depressor and the tube inserted to a depth of one third to one half the length of the reptile’s torso. This method, however, is often stressful for the reptile: the alternative is to syringe fluids into the mouth but there is a risk of inhalation in a debilitated reptile. A pharyngostomy or oesophagostomy tube may be placed for nutritional support, and so may be used for fluid therapy.
What is a chelonian?
the family of tortoises and turtles
How do you tube feed a chelonian?
The oral route can be used as for lizards and snakes. A pharyngostomy tube or oesophagostomy tube may be placed, as described later, and, up to, 10 ml/kg administered at a time. This is an excellent method of enabling an owner to give food, fluid and medication to their tortoise at home. Within 4 days of anorexia developing, a reptile is likely to be suffering from hepatic lipidosis making ongoing anorexia more likely. Such patients will often not eat again voluntarily for several weeks or months: a pharyngostomy tube enables food to be administered. Alternatively, a stomach tube may be inserted for each feed, if this will only be for a few days. The feeding tube is first pre-measured from the tip of the extended nose to the line where the pectoral and abdominal ventral scutes connect. It can then be lubricated and passed, after extending the head and gently prising the mouth open with a wooden or plastic speculum. There is marked variability in the ability to gavage chelonia. Testudo ssp. can usually be gavaged conscious, whilst many Geochelone individuals need sedation to allow oral medication. Similarly, some aquatic species (e.g. Trachemys ssp.) are easy to gavage; whilst others (e.g. Chelydra ssp.) pose a serious threat to the handler. Post-dosing regurgitation, through the mouth or nostrils is common, but rarely leads to glottal aspiration.
How do you place an oesophagostomy tube in a reptile?
Oesophagostomy tube placement
McCormack (2015) describes and demonstrates this procedure.
A suitable sized tube is selected for the patient: a suitable food, which can be passed down the selected tube, must also be available (e.g. Lafeber Emeraid® Herbivore). The tortoise is anaesthetised (the author (Forbes, N.A) favours Alfaxalone 6-9mg/kg, via the sub carapacial sinus). The tube is measured from the side of the neck (entry point for the tube) to the central plastron area and marked at this point. The side of the neck is prepared aseptically. A pair of artery forceps is passed per os, and the skin is tented up well, to ensure the jugular vein is away from the incision site, on the left side half way between the angle of mandible and thoracic inlet. The skin and oesophageal wall, over lying the point of the forceps, is carefully incised. The forceps are pushed through the incision and used to grasp the end of the tube. The tube is pulled up into the mouth, until the mark, rests at the incision site. Elastoplast is wrapped either side of the tube, then sutured to the skin. The distal tip of the tube is then reversed and carefully fed back down the oesophagus, being pushed down into the stomach with a large cotton bud. Tape is then placed over the medial aspect of the left foreleg and the cranial edge of the carapace (shell), to prevent the patient pushing the tube out. The tube is taped onto the carapace, leaving sufficient free length, for the head to extend and retract fully. The tortoise is fed daily via the tube. It can feed around (the tube) once its appetite returns. The tube is removed once the tortoise has been eating sufficiently by itself for one week. It is of course essential to ensure that the tortoise husbandry at home (especially temperature) is appropriate. Oesophagostomy tubes can also be placed in other reptiles, e.g. lizards, in a similar manner.
Where do you give subcutaneous fluids in a snake?
Snakes
The lateral aspect of the dorsum of the snake, in the caudal third of its body, is the ideal site for administration of subcutaneous fluids. This is a good technique for routine post-operative administration of fluids, for longer recovery patients undergoing minor surgical procedures, such as skin mass removal. There is a lymph sinus situated subcutaneously, lateral to the epaxial muscles on either side, which, with correct positioning, can be used for moderately large volumes. It may, however, still be necessary to use several sites.
Where do you give subcutaneous fluids in a lizard?
Lizards
The lateral thoracic area is easily utilised for smaller volumes of fluids at any one site. There is a risk of the reptile developing a dark pigmented area over the injection site, particularly in the chameleon, gecko and iguanid family. Owners must be warned of this
Where do you give subcutaneous fluids in a chelonian?
Chelonia
This is an easy route for post-operative fluids and mild dehydration in this species. They may be administered just cranial to the hind limbs or in the skin folds, just lateral, to the neck. Relatively large volumes may be given via this route.
How do you give intracoelomic fluids in a snake?
Intracoelomic
Snakes
This route is especially good for more seriously dehydrated reptiles, as there is a larger vasculature for absorption at this site. The needle or butterfly needle is inserted two rows of lateral scales dorsal to the ventral scales, in the caudal third of the snake, but cranial to the vent. The needle is inserted so that it just penetrates the body wall, the plunger of the syringe is pulled back to ensure no organ puncture has occurred and the fluids are administered. If correctly inserted there will be no resistance to the injection.
How do you give intracoelomic fluids in a lizard?
Lizards
As for small mammals, the lizard should be placed in dorsal recumbency, with its head downwards, to encourage the gut contents to fall cranially and away from the injection site. The needle, preferably a 25 gauge or smaller, is advanced slowly just through the abdominal wall in the caudal right ventral quadrant. The plunger should be pulled back to ensure that no organ penetration has occurred and the fluids should be administered without any resistance.
How do you give intracoelomic fluids in a chelonian?
Chelonia
This route can be used in tortoises, although the maximum amount that can be given is 20-25 ml/kg/day. The confines of the shell mean that the fluids place too much pressure on the lung fields. Access sites include pre-femoral fossa, the area cranial to the hind limbs. This is the same site as for subcutaneous administration but the needle is introduced to a greater depth. One main risk of this route is bladder puncture as it lies in this area. Another route is the cranial access site, located lateral to the neck and medial to the front limb. The needle is kept close and parallel to the plastron and a ¾ inch needle may be inserted to the level of the hub.
What intravenous routes can be used in snakes?
Snakes
There are no major vessels that are easily accessible for this technique in snakes. Therefore, if an intravenous route is to be used one of the following routes must be used:
1. The ventral tail vein – this is more of a plexus of veins and may be accessed from the ventrum. The needle is inserted in the midline, one third of the tail length from the vent, and advanced, at a 90-degree angle, until it touches the coccygeal vertebrae. The needle is then retracted slightly, whilst applying negative pressure to the syringe, until blood flows into the hub. Fluids may then be administered.
2. The palatine vein – this is present on the roof of the mouth and is paired. Cannulation may be performed with a 25-27 gauge butterfly needle, although the snake frequently has to be sedated or anaesthetised to gain access.
3. Intracardiac fluids – this can be used in emergencies. The heart may be catheterised under sedation or anaesthesia only. When the snake is turned onto its back, the heart can be seen beating against the ventral scale, approximately one quarter of its length from the snout. A 25-27 gauge over-the-needle catheter may be inserted, between the scales, in a caudo-cranial manner at 30 to the body wall into the single ventricle. A bolus may be administered, or the catheter may be taped, glued or sutured in place for 24-48 hours.
4. Jugular veins – these can also only be accessed in an anaesthetised/sedated snake. To gain access, a full-thickness skin cut-down procedure may be made 2-3 inches caudal to the angle of the jaw and two rows of scales dorsal to the ventral scales. The jugular vein can then be seen medial to the ribs. It is advisable to use an over-the-needle catheter with plastic wings as it should be sutured in place
What intravenous routes can be used in lizards?
Lizards
This route can be difficult in small lizards and frequently requires sedation or anaesthesia. Several routes may be tried.
1. Cephalic vein – this is approached in the anaesthetised lizard using a cut-down technique. The incision is made through the skin on the cranial aspect of the middle of the antebrachium, at a perpendicular angle to the long axis of the radius and ulna. The vessel may then be catheterised using an over-the-needle catheter and sutured in place. This technique is only useful for lizards over 0.25 kg in weight.
2. Jugular vein – this vessel may be accessed via a cut-down technique in the anaesthetised or sedated lizard. An incision is made in a cranio-caudal manner 1 inch caudal to the angle of the jaw. An over-the-needle catheter may then be sutured in place.
3. Ventral tail vein – this is more of a plexus of veins. This can be performed in the conscious lizard and is accessed from the ventral aspect of the tail. It is frequently only suitable for one-off bolus injections and special care should be taken with species that exhibit autotomy (spontaneous tail shedding), such as day geckos and green iguanas. The needle is inserted at 90 degrees to the angle of the tail and advanced until it touches the coccygeal vertebrae. It is then withdrawn slightly with negative pressure applied to the syringe. When blood flows into the syringe, the infusion may begin.
What intravenous routes can be used in chelonians?
Chelonia
The main routes are outlined below:
1. Dorsal tail vein – this is more of a plexus of veins so a catheter cannot be placed: it is often not possible to give large volumes of fluids. It is accessed in the midline, on the dorsal aspect of the tail. The needle is inserted until it hits the coccygeal vertebrae at a 90-degree angle. The needle is then pulled back whilst applying negative pressure until blood flows into the hub. 2. Jugular veins – these may be accessed for catheter placement in the sedated or anaesthetised tortoise. The neck is extended and the head tilted away from the operator to push the neck towards him/her. The jugular vein runs from the dorsal aspect of the eardrum along the more dorsal aspect of the neck. An over-the-needle catheter may be directly placed, or in thicker skinned animals, a cut down technique employed.
3. Sub-carapacial sinus – this may also be used to administer fluids, typically being an easy site to access, even in an uncooperative patient. It is located in the midline, directly above the head (easiest with the head pushed in). A long needle is inserted just ventral to where the skin meets the carapace. The needle is angled up to the top of the carapace, with suction being applied as it is advanced. Once blood appears, fluids may be administered, at a rate of 5-10ml/kg.
What intraossous routes can be used for snakes?
Intraosseous
Snakes: This route is not possible in the snake.
What intraossous routes can be used for lizards?
Lizards
1. This is a good route for the smaller species of lizards where venous access is restricted or difficult but asepsis is essential. Sedation or anaesthesia is required. Hypodermic or spinal needles (23-25 gauge) may be used.
The following sites can be used-
2. Proximal femur – this may be accessed from the fossa created between the greater trochanter and the hip joint. This route may be difficult due to the 90 degree angle the femur often forms with the pelvis.
3. Distal femur – this is relatively easy to access from the stifle joint. It does restrict the movement of the stifle joint but it is easier to bandage the needle into this site. Access to the medullary cavity of the femur is easier via this route.
4. Proximal tibia – this is possible in the larger species. The needle may be screwed into the tibial crest in a proximo-distal manner..
What intraossous routes can be used for chelonians?
Chelonia
Two main sites can be used-
1. Plastro-carapacial junction/pillar – this is the pillar of shell which connects the plastron to the carapace. It is approached from the caudal aspect, just cranial to one of the hind-limbs. The spinal/hypodermic needle (21-23 gauge) is screwed into the shell. The angle of insertion should be parallel with the outer wall of the shell so that the shell bone marrow cavity is entered. In larger, older species, the shell may be too tough to be penetrated using this method.
2. Proximal tibia – this may be approached as for lizards. The area is prepared aseptically and the hypodermic/spinal needle screwed into the tibial crest in the direction of the long axis of the tibia, distally.
What is the PCV for most reptiles?
The PCV for most reptiles is ~ 30%.
What PCV percentage in a reptile may require a blood transfusion?
There are various articles describing blood transfusion in reptiles with some authors advocating transfusions when the PCV drops to 10% and others suggesting when the PCV has dropped below 5%
What routes can be used to deliver a blood transfusion to a reptile?
Blood can be administered via the intravenous or intraosseous routes
Do you need to cross match blood for reptiles prior to a blood transfusion?
Cross-matching of blood groups does not appear to be necessary for one off transfusions, but the donor and recipient should be the same species e.g. green iguana to green iguana, boa constrictor to boa constrictor. In a dire emergency, however, it is possible to transfuse any one of a family group with another from the same group i.e. iguanid to iguanid and boiid to boiid.
Oxyglobin® may be administered at 1-2ml/kg if no donor is available.
What percentage of blood can be withdrawn from a reptile blood donor?
Blood volume, up to 2% of the body weight of a healthy donor, may be taken into a pre-heparinised syringe and it is immediately transfused into the recipient.
How do you perform a pulse assessment in a reptile?
Pulse assessment
This is best performed using a Doppler ultrasound probe. It should be attached to the skin with a generous amount of coupling gel. The main site of attachment is around the outflow of the heart. This is located immediately caudal to the neck inlet in chelonia; around the mid-point of the first third of a snake (measured from the snout); or just in front of the point of the shoulder in most lizards.
How do you perform a Cardiac and respiratory auscultation in a reptile?
Cardiac and respiratory auscultation
It is difficult to auscultate reptiles owing to their scaly skin. To aid the passage of sound, the reptile may be wrapped first in a damp cloth-applying the diaphragm of the stethoscope to the cloth can improve sound transference. The lungs of snakes are particularly challenging to auscultate due to their elongated structure. Chelonia lungs are situated in the dorsal aspect of the carapace and lizard lungs are encased in the ribcage. It is not possible to auscultate the heart in the same way as a bird or mammal’s heart due to the slow rate at which it beats and due to it being three chambered (there is only one ventricle). Doppler probes should be used, therefore, to gain an idea of cardiac output, strength and any variations in flow.
How do you perform a Neurological assessment in a reptile?
Neurological assessment
This can be very difficult in reptiles, particularly if they are not within their preferred optimum temperature zone. They will be sluggish and lethargic if too cold. One of the commonest neurological problems is loss of the righting reflex in snakes i.e. they continually flip onto their backs.
This may be associated with the following conditions:
1) Vitamin B1 deficiency (garter/water snakes fed defrosted frozen fish which contains large amounts of thiaminases);
2) Permethrin/organophosphate toxicity (over-zealous owners treating their snakes for mites);
3) Meningo-encephalitis (usually associated with Acanthomoeba invadens or Gram negative bacteria);
4) Inclusion body disease (a retrovirus, particularly prevalent in pythons and boas).
Blindness and head tilts may be seen in tortoises associated with frost damage.
Hypocalcaemic tremors are common in female iguanas on low calcium diets or where no/insufficient UV light has been provided. Panniculus reflexes can be tested in snakes and lizards as for cats and dogs, although of course with less success in chelonia!
What conditions may a loss of the righting reflex in snakes i.e. they continually flip onto their backs be associated with?
One of the commonest neurological problems is loss of the righting reflex in snakes i.e. they continually flip onto their backs.
This may be associated with the following conditions:
1) Vitamin B1 deficiency (garter/water snakes fed defrosted frozen fish which contains large amounts of thiaminases);
2) Permethrin/organophosphate toxicity (over-zealous owners treating their snakes for mites);
3) Meningo-encephalitis (usually associated with Acanthomoeba invadens or Gram negative bacteria);
4) Inclusion body disease (a retrovirus, particularly prevalent in pythons and boas).
How do you perform pulse oximetry in a reptile?
Pulse oximetry
This is not as useful in reptiles because reptile haemoglobin is significantly different to mammalian. However, trends of readings are of some help as with birds. The clips/probes may be applied to the vent, tongue (if anaesthetised) or the extremities, in thin skinned smaller reptiles.
What is it important to consider when performing a blood biochemistry on a reptile?
Blood biochemistry
It should be noted that, as with birds, uric acid is the only useful indicator of renal function, although more than three quarters of kidney function must be lost before uric acid levels become elevated. Urea and creatinine levels do not provide information on renal function in reptiles. As with birds, no one parameter is specific for liver damage, although AST is more useful than ALT.
What blood tube should be used to perform haematology on a reptile and why?
All reptile blood samples should be collected into heparin tubes as potassium EDTA lyses the red cells of many reptiles.
What is the range of white blood cells in reptiles and hwy is it important to analyse a blood smear?
In general, white cell counts are in the range of 2-8x109/L. During an infection, there is often no change in the overall white cell count and therefore, creating a blood smear is a vitally important diagnostic technique. The reptile equivalent of the neutrophil, as in birds, is the heterophil.
How do erythrocytes and platelets appear in reptiles?
Erythrocytes and platelets are nucleated in reptiles. Erythrocytes are rugby ball shaped, with a similar shaped nucleus, and platelets are oval and smaller. Other cells are like mammals. . In snakes, large numbers of ‘azurophils’ (darkly basophilic staining monocytic cells) may indicate chronic infection.
What may be seen when assessing haematology if there s a chronic infection?
Occasionally, however, a circulating plasma cell (a lymphocyte that has developed to produce antibodies) may be seen, particularly in the face of a chronic infection
When may an azurophil been seen in a snake?
In snakes, large numbers of ‘azurophils’ (darkly basophilic staining monocytic cells) may indicate chronic infection.
What may a urinalysis in a reptile reveal?
Urinalysis
This is less useful than for mammals owing to the faecal contamination that occurs with reptile urine. It is important, however to look at the urates (white portion of the dropping) to see if there is any blood; or if they have turned mustard yellow or lime green. If the latter has occurred, biliverdinuria is present, which in reptiles, as in birds, is an indicator of liver damage /inflammation. Biliverdin is the main excretory product of the liver as opposed to bilirubin in mammals. The volume of water/true urine should be small in a healthy reptile’s dropping. If droppings are very watery, as opposed to being diarrhoeic, this may indicate polyuria.
Why do reptiles often require long courses of antibiotics?
It is important to note that many reptiles will need to be on antibiotics for considerable periods of time (months rather than weeks). This is due to their slower metabolism and, usually, advanced state of infection on presentation.
How can you calculate energy requirements in reptiles?
Calculation of energy requirements can be made using the formula:
BMR = k x (weight (kg)) 0.75
k, the constant, is 10 for all reptiles.
MER (metabolic energy requirement) is generally 1.5-2x the BMR (basal metabolic requirement). If disease is present this further increases the required calories (e.g. sepsis and burns may increase MER by 2-3x).
How can you encourage anorectic snakes to eat?
To encourage anorectic snakes to eat, several techniques may be employed including:
1) Warming the prey, before offering, by heating in a pot of hot water
2) Breaking the prey’s cranium open to release the scent of blood
3) Teasing the snake by moving the dead prey item around the cage, with forceps, to mimic live prey
4) Trying a variety of colours of prey- some snakes will only take dark furred rodents
5) To get a snake used to eating rodent prey after only eating fish (such as garter and water snakes) or amphibians (Hog-nosed snakes), wipe the rodent food source with the previously taken food item to transfer scent
6) Ensure that there are plenty of areas to hide- some boids and pythons like to consume their prey in a box/hide
7) Leave the prey in overnight- some species prefer to hunt at night
8) Going to the next smallest size of rodent. If adult mice were previously offered try fuzzies; if juvenile rats, try adult mice etc.
NB: The term pinkies refers to nude neonatal rat and mice pups; fuzzies refers to week old rat and mice pups with a thin covering of fur; and furries refers to juvenile rat and mice pups of a few weeks of age (1-3) which have a soft, but longer, covering of fur.
How do you stomach tube most species of snakes and chelonians?
Most species of snakes and chelonia may be stomach tubed, relatively easily, if they still do not want to feed of their own accord. In snakes, a dog urinary catheter is used and inserted to approximately the caudal end of the first third of the snake (roughly where the stomach lies). The volume given depends on the size of the snake: a 100gm garter snake gets a maximum of 4-5 mls and a 30kg Burmese python gets up to 100-200ml.
Tortoises can be stomach tubed by measuring from the extended tip of the head to the caudal edge of the large abdominal scutes, on the plastron. Most tortoises of 2-3kg may be stomach tubed with 10-15ml of feed at one time
Lizards may be gavaged liquid feed, as for mammals, or stomach tubed. It is important to use a mouth gag when tubing reptiles to prevent the tube being bitten in half. In all cases, feeding by stomach tube should be the last procedure before the reptile is put back into its vivarium. Otherwise it will become distressed and regurgitate its feed.
What can you do prior to stomach tubing a reptile to reduce the risk of refeeding syndrome?
Due to the risk of the re-feeding syndrome developing, if a snake or other reptile has been anorexic for some time, it is essential to rehydrate the patient before attempting to feed a high protein meal. Initial feeding should be started off using small volumes e.g. only 50% of the calculated calorie requirement, based on the current weight of the reptile. If excess calories and proteins are administered, the rapid uptake of glucose from the bloodstream into the cells, taking potassium and phosphorous with it, can lead to life-threatening hypokalaemia/hypophosphataemia. Monitoring of blood phosphorus and potassium is therefore recommended when treating chronically anorectic reptiles, whether carnivorous or herbivorous.
What are open wounds usually caused by in reptiles?
Open wounds are generally of two main types in reptiles- thermal burns (Pollock, 2011b) and infections.
What nursing can bve provided to a patient with open wounds?
Open wounds are generally of two main types in reptiles- thermal burns (Pollock, 2011b) and infections. It is important therefore to ensure correct antibiotic coverage for these wounds. As most reptile infections are due to Gram negative bacteria, fluoroquinolones and third generation cephalosporins may be administered (see ongoing medication section).
Povidine iodine diluted to 1:30 with water may be used to clean infected wounds. Topical medications, such as silver sulfadiazine creams, have good efficacy against Gram negative bacteria; as do topical eye drops containing gentamicin. Where an infected wound is present, daily changing of any dressings is advised.
Dressings that may be sutured to the skin, around large wound deficits, include Granuflex®, Allevyn thin® and Veterinary BioSISt®. The former should be used where the infection is under control as it encourages granulation tissue to form. In many cases this primary dressing is all that is required, particularly when sutured to the patient, as reptiles rarely remove dressings. In snakes, especially however, dressings are difficult to attach unless sutured. Gels such as Intrasite® or Nugel® may be used to cover the surface of wounds and promote further healing.
How do you place an oesophagostomy tube in a reptile?
Placement of feeding tubes
Oesophagostomy tube placement
1 Sedation or anaesthesia is required and good analgesia post implantation.
2 The site is surgically prepared with povidine iodine. It is essential to be particularly scrupulous as reptile skin is notoriously dirty. In chelonia, the ventral aspect of the lateral neck is prepared, 3-4 cm caudal to the angle of the jaw. In snakes, the ventral aspect of the lateral ‘throat’ region is prepared, 5-10 cm caudal to the angle of the jaw. In lizards caudal to the jaw hinge joint is prepared, half way between the dorsal and ventral body surface.
3 A pair of haemostats is passed through the mouth and pushed laterally and ventrally, pushing the body wall out.
4 A sharp incision is made, with a scalpel blade, over the point of the haemostats, through the skin and the underlying muscle.
5 The feeding tube should be as a large a diameter as can comfortably pass down the oesophagus- Foley catheters are useful in tortoises. The tube should be pre-measured, prior to insertion, so the depth of insertion is known. In tortoises, this is from the site of the incision to the mid portion of the plastron, half way through the abdominal scutes. An additional length should be included to allow attachment of the end of the feeding tube to the dorsal aspect of the carapace. The feeding tube, is grasped with the haemostats as they protrude through the incision, then pulled into the pharynx and introduced into the oesophagus.
6 Once in place, two pieces of zinc oxide tape may be attached to the tube, close to the skin surface. Sutures may be placed through this tape and attached to the skin. The tube may then be attached to the midline cranial carapace in tortoises or taped to the side of the neck for snakes. A bung should be inserted. Care of the tube is the same as oesophagostomy tubes in cats and dogs. Sterile water should be flushed through the tube, prior to administering food, to ensure the tube is patent and still in the correct place. The tube should also be flushed after feeding to remove any food debris.
What is it important to consider and how do you carry out manual restraint on a snake?
Snakes
The snake family includes boa constrictors, corn snakes, Burmese pythons, and garter snakes.
There is a wide range of sizes from the anacondas and Burmese pythons, which may reach lengths of 30 feet or more: to the thread snake family which may only be tens of centimetres long. They are all characterised by their elongated form, with an absence of limbs. The danger areas are their teeth: in the case of the more venomous species, such as the viper family, their fang teeth are dangerous. The constrictor and python family pose a danger due to their ability to asphyxiate their prey by winding themselves around the victim’s chest/neck.
The following restraint techniques may be employed. Non-venomous snakes can be restrained by initially controlling the head- this is done by placing the thumb over the occiput and curling the fingers under the chin. Reptiles, like birds have only one occipital condyle so the importance of stabilising the neck at occipital/atlantal joint cannot be underestimated. It is also important to support the rest of the snake’s body, so that not all the weight of the snake is suspended from the head. This is best achieved by allowing the smaller species to coil around the handler’s arm, allowing the snake to support itself.
In larger species (longer than 10 feet), it is necessary to support the body length at regular intervals: so, several people will be needed. It is vital to adopt a safe operating practice with the larger constricting species of snake. For this reason, a ‘buddy system’ should be operated, as with scuba diving, whereby any snake, longer than 5-6 feet in length, should only be handled if there are two or more people available. This is to ensure that if the snake was to enwrap the handler, the assistant could disentangle him/her by unwinding from the tail end first. Above all it is important not to grip the snake too hard as this will cause bruising and the release of myoglobin from muscle cells. This will lodge in the kidneys causing damage to the filtration membranes.
Venomous snakes (such as the viper family, rattlesnakes etc.) or very aggressive species (such as anacondas, reticulated and rock pythons) may be restrained initially using snake hooks. These are 1.5-2-foot steel rods, with a blunt shepherd’s hook on the end and are used to loop under the body of the snake to move it, at arm’s length, into a container. The hook may also be used to trap the head flat with the floor, before grasping it with the hand. Once the head is controlled safely, the snake is rendered harmless, unless it is a member of the spitting cobra family. Fortunately, these are unlikely to be encountered in general practice. If they were, however, plastic goggles or a plastic face visor must be worn, as they spit poison into the prey/assailant’s eyes and mucous membranes, causing blindness and paralysis.
What is it important to consider and how do you carry out manual restraint on a lizard?
Lizards
Lizards come in many different shapes and sizes: from a four-foot-long adult green iguana to a 4-5-inch-long green anole. They all have roughly the same structural format with 4 limbs (although these may become vestigial e.g. the slow worm) and a tail. The main danger areas are their claws and teeth; in some species, such as iguanas, their tails can lash out, in a whip-like fashion, causing injury.
Geckos, other than Tokay geckos, are generally docile, as are lizards such as bearded dragons. Others such as green iguanas may be extremely aggressive, particularly sexually mature males. They may also be more aggressive towards female owners and handlers, as they can detect pheromones secreted during the menstrual cycle.
Restraint is best performed by grasping the pectoral girdle with one hand, from the dorsal aspect. One forelimb is controlled with the forefinger and the thumb and the other between the middle and fourth finger. The other hand is used to grasp the pelvic girdle, from the dorsal aspect. One limb is controlled with the thumb and forefinger, the other between middle and fourth finger. The lizard may then be held in a vertical manner, with its head uppermost. The tail should be out of harm’s way, underneath the handler’s arm. If the lizard is held in this way, the handler should allow some flexibility as the lizard may wriggle. Overly rigid restraint could damage the spine. It is then possible to present the head and feet of the lizard away from the handler to avoid injury. Some of the more aggressive iguanas may need to be pinned down, prior to this method of handling. As with avian patients, the use of a thick towel to control the tail and claws is often very useful. In some instances, gauntlets are necessary for particularly aggressive large lizards; and for those that may have a poisonous bite (e.g. the Gila monster and the beaded lizard). It is important to assure that too much force is not used when restraining a lizard. Those with skeletal problems, such as metabolic bone disease, may be seriously injured. In addition, lizards, like other reptiles, do not have a diaphragm so overzealous restraint will cause the digestive system to compress the lungs, increasing inspiratory effort.
Geckos can be extremely fragile. The day geckos, for example, are best examined in a clear plastic container, rather than physically restraining them. Other geckos have easily damaged skin- latex gloves and soft cloths should be used to cup them in the hand, rather than physically restraining them. Small lizards may have their heads controlled between the index finger and thumb, to prevent biting.
It is important that lizards are never restrained by their tails. Many will shed their tails and not all of them will re-grow. Green iguanas, for example, will only re-grow their tails as juveniles (less than 2.5-3 years of age). Once they are older, they will be left tail-less.
Why should lizards never be restrained by their tail?
It is important that lizards are never restrained by their tails. Many will shed their tails and not all of them will re-grow. Green iguanas, for example, will only re-grow their tails as juveniles (less than 2.5-3 years of age). Once they are older, they will be left tail-less.
What is it important to consider and how do you carry out manual restraint on chelonia?
Chelonia
This includes all land tortoises (which the Americans refer to as turtles), terrapins and aquatic turtles.
Size differences in this order are not as great as for the other two families. It is still possible, however, to see large variety. Chelonia vary from the small Egyptian tortoises (weighing a few hundred grams), to adult leopard tortoises at 40 kg and the Galapogean tortoise family, which can weigh several hundred kilograms. Most chelonia are harmless, although surprisingly strong. The exceptions include the snapping turtle and the alligator snapping turtle, both of which can give a serious bite. Most of the soft-shelled terrapins have mobile necks and can also bite. Even red eared terrapins may give a nasty nip!
Restraint can be achieved as follows. For the mild tempered Mediterranean species, the tortoise may be held with both hands, one on either side of the main part of the shell behind the front legs. To keep it still for examination, the tortoise may be placed onto a cylinder or stack of tins: this ensures that the legs are raised clear of the table allowing it to balance on the centre of the underside of the shell (plastron). For aggressive species, it is essential that the shell is held on both sides, behind and above the rear legs, to avoid being bitten. To examine the head region in these species, it is necessary to chemically restrain them. For the soft shelled and aquatic species, soft cloths and latex gloves may be required to avoid marking the shell.
Why is it important to maintain personal hygiene when handling reptiles?
It is important to bear in mind that many species of reptile and chelonia have a normal bacterial flora in their digestive systems, which frequently includes species such as the Salmonella family. These bacteria are found in abundance all over the body of the reptile. Personal hygiene is therefore very important when handling these patients, to prevent zoonotic diseases.
How do you carry out chemical restraint on a reptile?
Chemical restraint
Many species, such as snakes and lizards, may be induced using a face mask or induction chamber with 3-4% isoflurane in 100% oxygen. Chelonia, however, will breath-hold for hours, so it is not possible to induce them by this means.
Some lizards may also fight and a quicker form of induction can be achieved using either:
propofol IV/intraosseous (all reptiles). This must be done with care as extravascular administration has caused serious sloughs and paralysis, when close to the spinal cord
or
alfaxalone (Alfaxan®) IV (all reptiles, particularly chelonia) or sub carapacial
or
ketamine +/- medetomidine mixed in the same syringe and given IM (all reptiles)
The patient may then be intubated and maintained on either oxygen alone or, if insufficiently anaesthetised, by adding 1-2% isoflurane and performing IPPV at 4-6 breaths a minute. Most reptiles will not breathe for themselves during anaesthesia, so IPPV is essential.
