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
