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NO PAIN, NO GAIN:
UPDATES ON ANALGESIA IN BIRDS AND SMALL MAMMALS
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
“The lack of understanding of pain relief in any species should not preclude the use of analgesic drugs”
- Joanne Paul-Murphy, DVM, DACZM, DACAW
What is pain? Pain is considered to have two components: physical discomfort caused by injury or disease
and emotional suffering. Are animals capable of feeling emotional pain? This has been difficult to prove
scientifically as only humans and non-human primates have a neocortex, the so-called “thinking” area of the
brain. Most studies of pain in animals examine their responses to nociceptive stimuli, but does that also
indicate an emotional component? Therefore, some scientists prefer to use the term nociception and antinociception in reference to ‘pain’ in animals, other than non-human primates. How do you determine if a
drug provides analgesia? Pharmacokinetic studies can be performed which indicate a change in plasma
concentration over time (relationship between plasma concentration and dose) in a particular species. These
types of studies can also examine the effect of different routes of administration on plasma concentrations.
Pharmacodynamic studies measure the effect produced by the drug using various methods including surgical
pain models, thermal pain models, weight-bearing studies, and MAC (minimum anesthetic concentration,
minimum alveolar concentration) reduction studies. Analgesic efficacy of NSAIDs (non-steroidal antiinflammatory drugs) is difficult to determine as these drugs are highly protein bound, tend to accumulate in
areas of inflammation, and may have a longer duration of analgesic effect than predicted from plasma levels
alone. Finally, there is significant species variability (even within different avian orders), and it may not be
wise to extrapolate doses from other species.
RABBIT ANALGESIA
How can you tell if a rabbit is painful? There are no accepted, objective criteria for assessing pain in rabbits,
and there are no behavior-based pain assessment scores for rabbits. Rabbits (and rodents) are prey species,
and hide any signs of illness until it has become advanced. Therefore, it is challenging to detect subtle signs in
a stressful new environment (i.e. in the hospital). A study by Leach et al. videotaped rabbits pre and postspay to determine behaviors that may be indicators of pain, and they found “inactive pain behavior” was the
most useful sign. Other signs of pain include a hunched posture, reluctance to move, and grinding of the
teeth. To determine which analgesic is appropriate, first the clinician must try to characterize the pain (the
author uses mild, moderate, or severe grades). Other considerations include whether the patient will be
hospitalized, and if he/she has any concurrent diseases (renal, GI, etc.).
Many clinicians are concerned about administration of opioids leading to decreased GI motility in rabbits,
especially those with pre-existing ileus. Experimentally, both mu and kappa agonists can inhibit motility in the
isolated rabbit intestine (in vitro). Also, morphine has been shown to decrease duodenal peristalsis and
overall GI tract transit time in healthy rabbits. Another study examined potential side effects on GI motility
after a single intramuscular dose of buprenorphine (0.1mg/kg). No effects were noted in GI tract transit
time, and pyloric contractions were actually increased.
Opioids
• Butorphanol
 Mild pain, good sedation – duration of action: 2-3 hours
 0.5-1mg/kg SQ, IM, IV
• Buprenorphine
 Mild to moderate pain, minimal sedation – duration of action: 6-12 hours
 Ceiling effect
 0.03 – 0.1mg/kg SQ, IM, IV
• Oxymorphone
 Moderate to severe pain, mild sedation – duration of action: 3-4 hours
 0.1 – 0.3mg/kg SQ, IM, IV
• Fentanyl CRI
 Severe pain, intra-op or post-op - moderate sedation
 Loading dose: 5-10μg/kg IV, maintenance dose: 10-30μg/kg/hr IV
 Post-op: 1.25-5.0 μg/kg/h IV
A recent study by Barter, et al examined the effects of fentanyl administration on MAC (minimum alveolar
concentration) of isoflurane in 6 healthy rabbits. They found that fentanyl reduced the MAC by
approximately 60%. Another study by Schnellbacher, et al examined the effects of a lidocaine CRI on the
MAC of isoflurane in rabbits. They too found that lidocaine, both at 50 μg/kg/min and reduced MAC by 11
and 22%, respectively. Lidocaine CRI (100 μg/kg/min) has also been compared to buprenorphine
(0.06mg/kg SC every 8 hours) for post-ovariohysterectomy analgesia in rabbits. In that study, a lidocaine CRI
provided a superior outcome in respect to fecal output, appetite, and lower cortisone and glucose levels.
Tramadol provides analgesia by increasing release and decreasing reuptake of serotonin and norepinephrine in
the spinal cord. The parent drug and one of its metabolites have mu opioid receptor agonist effects. There
are several studies which have examined the use of tramadol in rabbits, both oral and parenteral (injectable
tramadol is not commercially available in the US). A PK study of a single dose of tramadol at 11mg/kg orally
revealed minimal plasma concentrations, which were lower than what was expected to provide analgesia in
other mammals. Tramadol was also used in a MAC reducing study in rabbits, which showed a single dose at
4.4mg/kg IV lead to a clinically unimportant reduction of MAC. Udegbunam et al found the use of 10 and 20
mg/kg SQ could provide adequate analgesia in post-surgical period (gastrotomy) and had no adverse effects
on selected biochemical parameters in rabbits.
Meloxicam is the most commonly used analgesic for rabbits in the author’s practice, and there have been two
PK and safety studies evaluating its use in this species. The initial PK study found a single (1mg/kg PO) and
multiple doses (1mg/kg PO q24h x 5 days) achieved adequate plasma concentrations in rabbits that were
similar to those of other species which provided analgesia. No significant bloodwork changes were noted in
any animal in that study. A subsequent study also found similar plasma concentrations when meloxicam was
administered at 1mg/kg PO q24h for 29 days. No bloodwork or necropsy changes were noted at the end of
that study in any animal.
RODENT ANALGESIA
Clinical signs of pain in rodents are similar to rabbits and include a hunched posture, lack of or increased
grooming, and decreased activity levels. There are also several studies that showed the presence of an
observer may alter the behavior of rodents, and thus compound the ability to accurately take behavioral
assessments of these prey species.
Recently, the analgesic efficacy and safety of buphrenorphine in chinchillas has been examined. No analgesic
efficacy (as measured by limb withdrawal latencies following a thermal noxious stimulus) was achieved at 0.05
or 0.1mg/kg SC. 0.2mg/kg SC buprenorphine did provide thermal antinociception, but only for < 6 hours.
A dose-dependent effect on fecal output and food intake was also seen. Sustained release buprenorphine has
been evaluated in both mice and rats. Significant analgesia for 48 hours was noted in mice administered a SR
buprenorphine at 1.5mg/kg SQ, and no adverse effects were noted. 1.2mg/kg of SR buprenorphine
administered SC provided significant analgesia for 2-3 days after a single dose in rats.
Many clinicians have utilized transmucosal (buccal, OTM) administration of buprenorphine in rabbits and
rodents. Rabbits and rodents are not used in studies of transmucosal analgesics in human trials due to
variable uptake of the drugs because of increased keratinization of of their oral mucosa. A recent study by
Sadar et al examined the PK of buprenorphine at 0.2mg/kg IV and transmucosally in guinea pigs. They
found that a single OTM dose of 0.2 mg/kg buprenorphine results in plasma buprenorphine concentrations
above those known to be analgesic in rats for approximately 3 hours. IV administration at the same dose in
guinea pigs lasted for approximately 5 hours. Therefore, a 1kg guinea pig would require ~0.67mL of
buprenorphine every 3 hours if administered via the transmucosal route. The large volume, frequent
administration, and expense make this route of administration less than ideal in this species.
AVIAN ANALGESIA
How can you tell if a bird is painful? Most birds are prey species, and tend to hide their signs of illness,
including pain. In chickens, a crouched posture and immobility has been associated with prolonged pain,
stress, and fear responses. If birds are housed in a flock, separation of a bird from the flock can indicate pain.
Birds may show reduced vocalization, fluffed appearance, a decrease or increase in grooming behavior
(particularly over the painful site), or even feather-destructive behavior. Clinicians should always assume a
particular condition in an avian patient is painful if they feel it would be painful in a mammal or human—
regardless of bird’s behavior or outward appearance.
Opioids
Most birds do not appear to respond to mu opioid agonists in the same manner as mammals. In pigeons,
76% of opioid receptors in the forebrain were determined to be kappa-type receptors. Currently, the kappa
opioid agonists, such as butorphanol and nalbuphine, are recommended for acute pain and preemptive
analgesia in psittacines. However, there are some recent studies that have proven analgesic efficacy with mu
agonists in several avian species. A study by Hoppes et al showed no analgesia after administration of low
dose fentanyl in cockatoos, and a higher dose (0.2mg/kg subcutaneously) provided mild analgesia. This
higher dose was a very large volume, and hyper excitability was seen in some birds. Pavez et al proved a
fentanyl CRI had a MAC-sparing effect in red-tailed hawks and had minimal effects on measured
cardiovascular parameters. Several studies by Guzman et al have examined both the pharmacokinetics and
antinociceptive effects of hydromorphone in American kestrels. They examined a variety of doses, but saw
analgesic effects from 3-6 hours after IM administration of the highest dose (0.6mg/kg). The drug had a
rapid elimination and high bioavailability after IM administration. Moderate to severe sedation was noted in
several birds with this dose.
Tramadol is a weak mu-receptor agonist, which also inhibits reuptake of serotonin and norepinephrine, which
activates the descending pain inhibitor system. There are several recent studies that have examined the
pharmacokinetic properties and antinociceptive effects of tramadol in psittacine patients. In Hispaniolan
amazon parrots, the drug had poor oral bioavailability (~25%), and doses of 30mg/kg PO reached plasma
levels associated with analgesia in other species. A separate study by Guzman et al showed effective
antinociception in this species for up to six hours. Souza et al examined the pharmacockinetics of tramadol
in Hispaniolan amazon parrots after repeated oral doses and found that the drug required administration
every 6-8 hours to maintain plasma levels thought to provide analgesia. Interestingly, the oral bioavailability
of tramadol in bald eagles is much higher; therefore, the recommended dose for this species is 5mg/kg orally
every twelve hours. The antinociceptive effects of tramadol were recently examined by Guzman et al in a
smaller raptor species, the American kestrel. These results indicated that a 5mg/kg oral dose resulted in
effective analgesia for 1.5-6 hours, but adverse effects, including vomiting, diarrhea, and polyuria, were noted
in half of the birds. These studies are a great example of the challenges a clinician faces when trying to
extrapolate dosages between different species of birds, and how this may not always be appropriate.
There are numerous older studies that examine the use of butorphanol in avian species; however, these will
not be reviewed in this presentation due to time constraints. Based on these studies, the published doses for
psittacines is anywhere from 1-5mg/kg. Currently, butorphanol is the preferred injectable opioid analgesic in
this order. A study by Guzman et al from 2011 examined the pharmacokinetics of butorphanol in
Hispaniolan amazon parrots after intramuscular, intravenous, and oral administration at a dose of 5mg/kg.
Those results indicated that IM or IV administration every 2-3 hours would be consistent with published
therapeutic levels—antinociception was not examined in this study. Importantly, the oral bioavailability of
this drug was very low (<10%), which proves that butorphanol should never be given orally to psittacines in
hopes of achieving analgesia. Escobar et al examined the use of butorphanol on the MAC of sevoflurane in
guineafowl. Butorphanol was found to effectively reduce MAC in this species at dosages up to 4mg/kg, but
the effect was small and the duration was short. The pharmacokinetics and thermal antinociceptive effects of
butorphanol in American kestrels was recently evaluated by Guzman et al, who found no significant analgesic
or sedative effects in this species at doses from 1-6mg/kg IM. In contrast with psittacines, these findings
suggest that butorphanol may not provide effective analgesia in American kestrels.
Buprenorphine is a unique opioid, as it has a high affinity for mu, delta, and kappa receptor subtypes. It has
been traditionally classified as a partial mu agonist and delta and kappa receptor antagonist. It is commonly
used in veterinary medicine because of its long duration of action and favorable safety profile. Use of
buprenorphine in psittacines was examined by Paul-Murphy et al which found no analgesic effect when
administered at 0.1m/kg intramuscularly to African Grey parrots. As subsequent pharmacokinetic study by
the same authors found that a dose of 0.1mg/kg both IM and IV did attain plasma concentrations known to
provide analgesia in other species, but only lasted for approximately 2 hours. They determined to achieve
plasma levels that would provide analgesia in people, psittacines would require a dose of ~2.5mg/kg every
three hours. Several recent studies have examined the pharmacokinetics and antinociceptive effects of
buprenorphine in American kestrels. After both intramuscular and intravenous injections of 0.6mg/kg,
plasma buprenorphine concentrations remained above a level deemed to provide analgesia in other species
for 9 hours. Also, at doses of 0.1, 0.3, and 0.6mg/kg, buprenorphine resulted in thermal antinociception for
at least six hours in this species.
Non-steroidal anti-inflammatories (NSAIDs)
Meloxicam is the most frequently prescribed NSAID for companion birds, and will be the focus of this
discussion. There are several recent studies which examined the use of this drug in psittacines, and the
dosages required for analgesia and plasma concentrations determined from those studies are ten-fold higher
than most mammalian doses. Cole et al examined intramuscular administration of meloxicam at doses of
0.05, 0.1, 0.5, and 1.0mg/kg for three doses in Hispaniolan amazon parrots with experimentally induced
arthritis by use of a weight-bearing perch. Of those doses, only the highest (1.0mg/kg) was effective at
relieving arthritic pain. A subsequent study by Molter et al examined the pharmacokinetics of a single
1.0mg/kg dose of meloxicam administered IV, IM, and PO in Hispaniolan amazon parrots. They found that
the oral dose had poor and highly variable bioavailability. Doses of no less than 1.0mg/kg orally twice daily
were recommended in this species. The post-operative analgesic effects of meloxicam in pigeons have also
been examined by Desmarchelier et al in 2012. The authors found that doses of 2mg/kg orally twice daily
provided quantifiable analgesia that appeared safe in this species—doses of 0.5mg/kg orally twice daily were
ineffective. A pharmacokinetic study of single dose of 0.5mg/kg meloxicam IV and PO in red-tailed hawks
and great horned owls did not maintain plasma concentrations that were likely to be therapeutic in either
species by either route of administration.
The adverse effects of NSAIDs in domestic dogs and cats are well known, and there are also several studies
which examine possible toxicity of NSAIDs in avian species. Repeated intramuscular injections of 2mg/kg
meloxicam in Japanese quail did lead to muscle necrosis at the injection sites, but no kidney lesions were
noted in any study bird. Liver and muscle necrosis was noted in pigeons after repeated intramuscular
injections of carprofen at dosages of 2, 5, or 10 mg/kg once daily for 7 days. A recent paper by Dijkstra et
al examined possible side effects of oral meloxicam on biochemical and hemostatic variables in healthy
Hispaniolan Amazon parrots. Birds were administered meloxicam orally at 1.6mg/kg twice daily for fifteen
days—no apparent negative changes were noted in renal, gastrointestinal, or hemostatic variables in any bird.
Clinicians should be aware that these studies were all performed in healthy animals, and the effects of
NSAIDs may be different in birds with compromised health.
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Escobar, André, et al. "Effects of butorphanol on the minimum anesthetic concentration for sevoflurane in
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AN INSIDER ON INCISORS: RABBIT AND RODENT DENTISTRY
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
Dental formulas of common exotic small mammals
Incisors
Canines
2/1
0/0
Rabbits
Premolars
Molars
3/2
Elodont/hypsodont
1/1
Elodont/hypsodont
3/3
Elodont/hypsodont
3/3
Elodont/hypsodont
Hystricomorphs
Elodont/hypsodont
1/1
Elodont/hypsodont
Myomorphs
1/1
Elodont/hypsodont
0/0
0/0
3/3
Anelodont/brachyodont
Sciuromorphs
1/1
Elodont/hypsodont
0/0
1-2/1
Anelodont/brachyodont
3/3
Anelodont/brachyodont
(guinea pigs,
chinchilla, degus)
(mice, rats, hamsters,
gerbils)
(prairie dogs, squirrels)
0/0
Common dental terms
• Aradicular = elodont: continuously growing teeth that do NOT develop anatomical roots
• Anelodont: teeth with limited period of growth
• Hypsodont: long-crowned teeth
• Brachyodont: short-crowned teeth
RABBITS
All rabbit teeth are hypsodont and elodont. Their teeth are composed of a clinical crown (visible
above the gingiva), reserve crown, and apex. As all of their teeth grow continuously and lack anatomic
roots, the phrase “tooth-root abscess” is incorrect. They are also anisognathic—their mandible is
narrower than their maxilla. They have a ~10 degree angle of occlusion for their premolars and
molars. This is important to maintain, if possible, with occlusal adjustments (dental procedures). When
the incisors are in contact, the premolars and molars are not in occlusion. Their incisors have a chiseled
edge—which also should be maintained when performing dental procedures. Normal rabbit incisors
grow quite quickly—a rate of 2 mm/week for the maxillary incisors and 2.4 mm/week for the
mandibular incisors. Their chewing motions vary for each type of substrate they ingest. When chewing
natural vegetation (hay, greens, grasses), the major chewing motion (“power stroke”) is horizontal. This
leads to a normal wearing pattern of all the teeth. However, when rabbits eat a pelleted diet or grains,
the major chewing motion is more vertical, which leads to uneven tooth wear over time.
Etiology of dental disease in rabbits
There are several proposed causes for dental disease in rabbits, and it is likely a multi-factorial disease.
Some of the more common hypotheses include genetics (dwarf breeds more predisposed), metabolic
bone disease, inappropriate diet (lack of hay and other abrasive dietary items), trauma, and aging.
Harcourt-Brown describes a progressive syndrome of acquired dental disease in rabbits with three main
stages: early changes, acquired malocclusion, and late changes. The early changes include apical
elongation of the teeth and enamel hypoplasia. Elongated mandibular premolars and molars can be
palpated on the ventral aspect of the mandibles, and elongated maxillary premolars and molars can
sometimes be palpated ventral to the eyes. This can be painful, and owners may notice the rabbit
refusing to eat harder foods, such as hay. Epiphora may also be noted due to partial or complete
occlusion of the nasolacrimal duct by apical elongation of the maxillary incisors and/or premolars.
Acquired malocclusion is the next stage, and is characterized by development of dental points or spurs
from inappropriate wear on the premolars-molars. The final stages involve fractured crowns, and
uneven teeth due to destruction of the germinal tissue which will lead to cessation of growth.
Common complications of chronic dental disease include apical tooth abscesses and dacryocystitis
(chronic infection of obstructed tear ducts).
Oral exam and diagnostics
A limited oral exam can be performed on awake patients, with either an otoscope cone or a human
nasal speculum (author’s preference). The view of the caudal-most teeth is limited, and a complete oral
exam often requires heavy sedation or general anesthesia. Due to their natural chewing motions, dental
points will most likely be found on the lingual surface of the mandibular premolars/molars and the
buccal surface of the maxillary premolars/molars. Any purulent material or foul odor in the mouth
should be noted. The clinician should also thoroughly palpate the ventral surface of each mandible and
ventral to each eye for any masses or asymmetry. Ideally, some form of imaging would be obtained
from each patient with suspected dental disease to help determine the extent of the sub-gingival disease.
Computed tomography is superior to skull radiographs for imaging of the skull; however, this is not
always possible for every patient.
Treatments
Numerous treatment methods for dental disease in rabbits have been published. Before treatment is
initiated, no matter which methodology, the owner should be notified that dental disease in rabbits
often requires life-long treatment, and a single procedure is not likely to be curative. Occlusal
adjustments should be performed on anesthetized rabbits—ideally they would also be intubated and
have IV access. The author uses a round-tipped bur on a slow-speed handpiece for premolar and
molar adjustments, and the high-speed handpiece for incisor adjustments. The clinician should attempt
to correct the dental abnormalities so they approximate, as closely as possible, the normal occlusion
rather than reducing the clinical crowns as short as possible.
Several treatments for apical/jaw abscesses in rabbits have been published including marsupialization,
complete abscess excision, and packing the abscess with a variety of antibiotic impregnated items
(PMMA beads, gauze, umbilical tape, etc.). The author prefers marsupialization, and this is the
treatment of choice for the dentistry and oral surgery department at the University of California, Davis.
With this technique, a large punch biopsy is used to create a hole in the skin and abscess capsule. The
abscess is debrided and flushed with dilute chlorhexidine and saline, and the skin edges sutured to the
edge of the abscess capsule with simple interrupted sutures. The abscess capsule is flushed, ideally,
twice daily at home with the owner, and the skin defect usually heals closed within 1-2 weeks. As rabbit
pus is not liquid, it will not drain like a cat/dog abscess. Therefore, lancing and drain placement is
inappropriate and often ineffective for this species.
Long-term antibiotics are often indicated in rabbits with dental disease, and especially those with apical
abscesses. Treatment of abscess with antibiotics alone may reduce the size of the abscess during
treatment, but is unlikely to lead to complete resolution of the infection. In a retrospective review of
cultures of rabbit abscesses, 100% of the strains tested were susceptible clindamycin (cannot safely be
given orally to rabbits), 96% were susceptible to penicillin (can be given parentally to rabbits), 54%
were susceptible to ciprofloxacin (enrofloxacin), and only 7% were susceptible to trimethoprimsulfamethoxazole. A recent proceedings evaluated the microbiology and antibiotic susceptibilities of
odontogenic abscesses in rabbits via retrospective analysis of rabbits that presented to the UC Davis
companion avian and exotics service over the last 14 years. The two most common anaerobic isolates
were Fusobacterium spp and Peptostreptococcus spp, and the most common aerobic isolates were Pseudomonas
aeruginosa and Pasteurella sp. The large number of anerobic bacteria culture was unexpected, and
possible safe options for anaerobic treatment in rabbits include penicillin, chloramphenicol and
metronidazole.
RODENTS
Guinea pigs and chinchillas are the most common rodent species presented for dental disease, and they
will be the focus of this presentation. Chinchillas (similar to rabbits) have a 0-10° angle to their occlusal
plane, versus guinea pigs have a 30° angle. The growth rate of chinchilla premolars and molars is
approximately 3-4 mm/month. Guinea pig incisors grow at a rate similar to rabbits. Guinea pig
mandibular incisors are 3 times longer than the maxillary incisors, and this should not be misinterpreted
as overgrowth. Guinea pigs and Russian hamsters lack ferric oxide (which looks yellow) in their enamel
contrary to other rodents.
Guinea pigs commonly have incisor malocclusion and mandibular premolar-molar overgrowth and
malocclusion. 80% of guinea pigs with dental disease have a low body condition score (BCS). In one
study, there was no correlation between bruxism and dental disease. Food impaction and gingivitis significantly
more common in chinchillas than guinea pigs and rabbits. Drooling and perioral staining was seen in 70% of
chinchillas affected by dental disease. Overall, rodents have a significantly lower prevalence of dental disease
than rabbits and they are less prone to periapical abscesses and osteomyelitis (Capello and Lennox, 2012).
Diagnostics and treatments for rodents with dental disease are similar to those for rabbits, as described above. A
recent paper by Schweda, et al describes the use of computed tomography (CT) for the assessment of dental
disease in 66 guiena pigs. 95% of the patients had a history of dysphagia, but only 2 of the guinea pigs showed
ptylism. Other common presenting complaints were reduced body condition score (50% of patients) and
diarrhea (30%). Obliquely worn incisors were a common physical exam finding, and a majority of these cases
were caused by periapical disease of the cheek teeth. Tooth fractures were noted in ~25% of all the guinea pigs.
REFERENCES
1.
2.
3.
4.
5.
6.
Verstraete FJ, Osofsky DA. Dentistry in pet rabbits. Compendium. 2005.
Osofsky A, Verstraete FJ. Dentistry in pet rodents. Compendium on continuing education for the practicing veterinarian.
2006.
Capello V. Diagnosis and treatment of dental disease in pet rodents. Journal of Exotic Pet Medicine.
2008;17(2):114-123.
Gracis M. Clinical technique: normal dental radiography of rabbits, guinea pigs, and chinchillas. Journal of
Exotic Pet Medicine. 2008;17(2):78-86.
Capello V, Cauduro A. Clinical technique: application of computed tomography for diagnosis of dental
disease in the rabbit, guinea pig, and chinchilla. Journal of Exotic Pet Medicine. 2008;17(2):93-101.
Tyrrell KL, Citron DM, Jenkins JR, Goldstein EJC, Group VS. Periodontal Bacteria in Rabbit Mandibular and
Maxillary Abscesses. Journal of Clinical Microbiology. March 1, 2002 2002;40(3):1044-1047.
7.
8.
9.
10.
11.
Reiter AM. Pathophysiology of Dental Disease in the Rabbit, Guinea Pig, and Chinchilla. Journal of Exotic
Pet Medicine.17(2):70-77.
Boehmer E, Crossley D. Objective interpretation of dental disease in rabbits, guinea pigs and chinchillas.
Tierrztliche Praxis Kleintiere. 2009;4:250-260.
Legendre LÃF. Oral disorders of exotic rodents. Veterinary Clinics of North America: Exotic Animal Practice.
2003;6(3):601-628.
Crossley DA. Oral biology and disorders of lagomorphs. Veterinary Clinics of North America: Exotic Animal
Practice. 2003;6(3):629-659.
Jekl V, Redrobe S. Rabbit dental disease and calcium metabolism: the science behind divided opinions.
Journal of Small Animal Practice. 2013;54(9):481-490.
GUINEA PIG AND CHINCHILLA HUSBANDRY
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
TAXONOMY
Guinea pigs and chinchillas belong to the order Rodentia, which is derived from the latin word rodere
meaning to gnaw. Rodentia is the largest mammalian order, comprising 40% of all mammalian
species (approximately 2277 total species). All rodents have 2 continuously growing (elodont)
mandibular and maxillary incisors. The order rodentia contains three suborders: myomorpha (mice,
rats, hamsters, gerbils), sciuromorpha (dormice, squirrels, chipmunks, prairie dogs), and
hystricomorpha (guinea pigs, capybaras, degus, chinchillas). There are several anatomic differences
that separate these three suborders—mainly the bones of the skull, jaw muscles, and dentition. Only
rodents within the suborder hysricomorpha have continuously growing incisors, premolars, and
molars.
GUNIEA PIGS
Guinea pigs (Cavia porcellus) are native to South America and inhabit a variety of landscapes. They
live in groups of up to 10 individuals and are mainly nocturnal. They were domesticated around 900
B.C., and became a popular pet in Europe in the late 1500s. In South America, they are commonly
kept both as pets and for for food (cuy). Common breeds of guinea pigs include short haired (also
called American or English), Abyssinian (with rosettes of hair), and Peruvian (long haired).
Males = boars
Females = sows
Young = pups
Guinea pigs are precocial, meaning they are born with hair, open eyes, and teeth. They start eating
hay and other foods within 1-2 days of birth.
Lifespan
5-7 years
Male adult weight
900-1200 grams
Female adult weight
700-900 grams
Sexual maturity
Males: 3 months
Females: 2 months
Gestation
59-72 days (68 days average)
Litter size
1-13 kits (2-4 average)
Guinea pigs have no tail, hairless pinnae, 4 digits on their forelimbs, and 3 on their hindlimbs. They
possess one pair of inguinal nipples, and a sebaceous gland over their lumbar spine, often referred to
as the “grease gland.” The palatal ostium is composed of mucosa at the base of their tongue,
ventrally, and the soft palate dorsally. Their epiglottis and opening to the trachea is caudal to this
ostium, which makes endotracheal intubation very difficult in this species. The pubic bones of
female guinea pigs do not separate in nulliparous animals after 6 months of age; therefore, if a
guinea pig is bred for the first time after this age, there is a high risk of dystocia. The young are
delivered rapidly, and are usually weaned at about 3 months of age.
Guinea pigs are very social species, and do best when housed in pairs or trios. Adult, intact males
may be aggressive, and shouldn’t be housed together. This species reaches sexual maturity at about
2-3 months of age, and the sexes should be kept separate to avoid unwanted pregnancies. Guinea
pigs should not be housed with or interact with rabbits, as rabbits commonly carry Bordetella
bronchiseptica, which can lead to serious illness in guinea pigs.
Housing requirements:
• Minimum size: 7 ft2 for the 1st cavy, 2-4 ft2 for each additional
• Solid sides for several inches from bottom, then wire sides +/- top
• Temperature range: 65-79°F
Sipper bottles should be used, as many guinea pigs will quickly soil any water bowls that are left in
their cage. Wood shavings and corn cob bedding should be avoided, and paper-based bedding is
recommended. Wire bottom cages are also not recommended at this can lead to pododermatitis.
Frequent cleaning (every 1-2 weeks) is recommended to avoid buildup of moisture and waste which
can lead to upper and lower respiratory tract disease. As most guinea pigs are shy animals, they
should also be provided with hide boxes.
The digestive system of a guinea pig is very similar to a rabbit—they have a large cecum and are
hind-gut fermenters. High fiber diets are important for proper dental health (occlusion of the teeth),
normal GI motility, and maintaining normal cecal flora. Guinea pigs should have access to free
choice grass hay, timothy-based guinea pig pellets (6gm/100gm body weight), and 1-2 cups of fresh
vegetables per day. Vegetables high in vitamin C are recommended, and include turnip greens,
mustard greens, dandelion greens, kale, parsley, collards, and bell peppers. Similar to people, guinea
pigs lack the enzyme L-gulonolactone oxidase, and cannot synthesize ascorbic acid (vitamin C).
Adults require 10mg/kg/day in their diet. Vitamin C is highly labile and light sensitive. Most guinea
pig pellets are fortified with Vitamin C, but the best source is fresh fruits and vegetables.
CHINCHILLAS
Chinchillas (Chinchilla laniger) are also native to South America, but from areas of higher elevation in
the Andes. They are gregarious, and live in groups of several hundred animals. They are considered
crepuscular feeders, and eat native shrubs and grasses, which have a very high silica content.
Chinchillas were domesticated more recently than guinea pigs, and were initially brought into the US
in the 1920s for the fur trade. In fact, the North American domestic population of chinchillas was
descendent from only 11-13 individuals. Currently, domesticated chinchillas are used for pets,
research (auditory disease in people), and the fur trade. There are no recognized breeds, but there
are a variety of coat colors.
Male = buck
Female = doe
Young = kits
Chinchilla kits are also precocial at birth, and are weaned at 6-8 weeks. They have large, hairless
pinnae, and even larger tympanic bullae (hence their use in auditory research). They have four toes
on both front and rear feet. Similar to guinea pigs, they too have a palatal ostium, which makes
intubation challenging.
Lifespan
10-15 years (20 years max)
Male adult weight
400-500 grams
Female adult weight
450-700 grams
Sexual maturity
7-9 months
Gestation
105 - 118 days (111 days average)
Litter size
1-6 pups (2 average)
Chinchillas are very active rodents, and need regular handling and socialization to make good pets.
They are considered crepuscular/nocturnal. They should be handled carefully to prevent fur slip,
which is when large amounts of hair epilate in patches due to inappropriate handling and struggling
of the animal. Many people advocate to grasp the base of the tail with one hand and hold the front
feet with the opposite hand.
Chinchillas also require large cages, where vertical space is as important as horizontal space. A hide
box should be provided and solid flooring is best. Different levels should be provided for climbing
and jumping. Substrate recommendations are similar to guinea pigs. Dust baths should be provided
several times per week. These help maintain their coat condition, but excessive use could lead to
conjunctivitis. Commercial dust mixes are available at most pet stores, and are usually a mix of
silver sand and Fuller’s earth. Chinchillas do not handle heat or humidity well—their ideal
temperature is 64-72°F, and humidity less than 50%. They are prone to heat stroke at temperatures
as low as 80°F.
They should be housed in pairs or groups, and breeders often keep chinchillas in polygamous
groups. Chinchillas, like guinea pigs, are strict herbivores and hind-gut fermenters. A diet
containing 20-35% fiber is recommended, and often grass hays and greens help meet this need.
Pellets can also be provided, but care should be taken to avoid alfalfa-based pellets (which are
common), as these are too high in calcium and proteins. On average, an adult chinchilla should
receive 1-2 Tbs/animal/day. Chinchilla pellets are similar to rabbit pellets, but are usually longer in
shape to facilitate prehension by the animals. Salt and mineral blocks are not necessary for this
species.
CAN I CATCH THAT?
AN UPDATE ON EXOTIC ANIMAL ZOONOSES
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
REPTILE AND AMPHIBIAN ZOONOSES
Salmonella spp.
Salmonella is a gram-negative facultative aerobic bacteria with more than 2400 recognized
serotypes.1 Some public health officials consider all serotypes pathogenic, but it is important to
remember that in certain species, including reptiles and amphibians, Salmonella is considered a
normal inhabitant of their gastrointestinal tract. Most reptiles and amphibians that culture positive
for this bacteria have no clinical signs. Approximately 1.4 million cases of human salmonellosis are
reported annually in the United States—the vast majority of these infections are food-borne. In
1996, the CDC estimated that reptile-associated salmonellosis accounted for 3-5% of all human
cases.1 Despite this low number, reptile-associated salmonellosis is still an important zoonosis, and
clients should be educated about this disease and prevention strategies.
To help reduce the incidence of turtle-associated salmonellosis (particularly in children), the FDA
passed a regulation in 1975 that restricted the interstate shipment and sale of all turtle eggs and live
turtles with a carapace length of less than 10.2cm (~4 inches). However, enforcement of this law is
sporadic, and cases still occur. In 2013, the CDC reported an outbreak of salmonellosis in 473
people from 41 states. 69% of ill persons reported exposure to turtles before their illness began, and
88% of infected people reported exposure specifically to small turtles (turtles with a carapace length
of < 4 inches). 70% of ill persons were children < 10 years of age. The turtles were purchased from
a variety of sources including pet stores and street vendors.
Clients may think of Salmonella originating mainly from chelonians, but all reptiles should be
considered possible carriers for this bacteria. In July of 2014, the CDC reported an outbreak
salmonellosis linked to contact with pet bearded dragons purchased from multiple stores in different
states. 166 persons were infected, and 59% of those were children 5 years of age or younger.
Amphibians can also be sources of infection for salmonella in people. In 2011, an ongoing
nationwide outbreak of human Salmonella Typhimurium infections were linked to a single African
dwarf frog breeding facility in Madera County, California. 2 241 people were infected from 42 states,
with 69% of infected persons less than 10 years of age.
Burkholderia pseudomallei
Burkholderia pseudomallei is a gram negative bacteria and the causative agent of melioidosis. It is
considered a Tier 3 foreign animal disease in the United States, and is also a Biosafety level 3 agent.
Endemic to Southeast Asia, Australia, and occasionally reported from Central and South America,
most cases seen in the US (in animals and people) have a travel history to these locations.
Pneumonia is the most common clinical manifestation in people, and this can progress to multiorgan abscessation, sepsis, and death if not promptly treated. Infections in domestic animals are
most commonly reported in cattle, goats, and swine; however, B. pseudomallei has a wide host range.
Three cases of infection with B. psudomallei in pet iguanas were reported in 2014—two from
California3 and one from the Czech Republic.4 All iguanas were diagnosed with abscesses—one over
the shoulder, one had multiple hepatic abscesses, and one had a recurrent abscess at the base of the
tongue. All iguanas were purchased from a pet store at a young age, none of the iguanas had access
to the outdoors, and none had left their state/country after purchase. Euthanasia of all iguanas was
recommended due to the zoonotic risk. No pet-related B. pseudomallei infections in humans have
ever been reported, including these recent cases. B. pseudomallei was cultured from the environment
of one of the iguanas, suggesting environmental contamination is possible. Owners and
veterinarians should take proper precautions not only when handling a potentially infected animal,
but also with anything it has contacted. The infected iguanas from California likely originated from
Central America, which may be where the initial infection occurred.
Campylobacter spp.
Campylobacter spp. are the most common cause of bacterial gastrointestinal disease in humans, and C.
jejuni is the most frequently reported. It is primarily considered a food-borne pathogen, but human
infections can also occur via direct contact with a variety of animals. C. fetus infects livestock (cattle
and sheep) but is considered an opportunistic human pathogen that mainly causes systemic
infections in immunocompromised people. In a recent study, 109 cloacal swabs from privately
owned and zoo-housed reptiles in Italy (chelonians, lizards, and snakes) were cultured for
Campylobacter. 7% of the samples were positive for several different species of Campylobacter,
including C. fetus. None of the snakes were positive.5 A similar study was performed in Taiwan—
179 fecal samples from pet store, privately owned, and wild reptiles were cultured for the presence
of Campylobacter. 6.7% of the samples were positive, all for C. fetus. Interestingly, the strains of C.
fetus isolated from the reptiles in this study were genetically distinct from the classical mammalian C.
fetus.6
A newly proposed subspecies, C. fetus subsp, testudinum subsp. nov has been isolated from both
humans and reptiles, and is believed to be of reptile origin. A recent paper reviewed the 9 reported
human cases in the United States—all were men with an average age of 73 years with a variety of
clinical signs.7 Most of the infected men were of Asian descent and reported eating dishes that may
have contained turtles or frogs at different Chinese restaurants.
AVIAN ZOONOSES
Chlamydia psittaci, recently renamed from Chlamydophilia psittaci, is a gram-negative, obligate
intracellular bacteria that causes chlamydiosis in birds and psittacosis in people. Infected birds may
show non-specific clinical signs (ocular-nasal discharge, lethargy, yellow-green urates), but many
remain asymptomatic and intermittently shed organisms, especially during times of stress. A recent
study examined risk factors associated with infection in psittacine birds sold at pet markets and kept
as pet birds in Brazil.8 The overall infection frequency was 10.6% and birds kept in households were
less frequently infected than those at markets (3% vs 17%). Of all the epidemiologic factors
analyzed, only population density and cage hygiene of the birds kept in markets were significantly
associated with C. psittaci infection. Another recent study tested 26 hyacinth macaws for C. psittaci
after they had been confiscated from the illegal pet trade in Brazil.9 Not surprisingly, these birds
were maintained under poor husbandry conditions, and 65% of them tested positive for infection.
These reports highlight the importance of client education regarding C. psittaci when a veterinarian
suspects a bird was obtained from the illegal pet trade, or had been group-housed in a bird market.
SMALL MAMMAL ZOONOSES
Salmonella spp.
Pet small mammals, are an under recognized source of human salmonella infections. In a 2007
publication, 28 cases of rodent associated multi-drug resistant Salmonella infections (S. enterica
serotype Typhimurium) were documented in the US from December 2003 to September 2004. 10
59% of cases reported exposure to pet hamsters, mice or rats. The median age of affected persons
was 16 years and 40% of patients required hospitalization.
African pygmy hedgehogs can harbor several serotypes of Salmonella, and animals with subclinical
infections can intermittently shed the bacteria. Hedgehogs also have a tendency to walk in their
feces, which can facilitate spread throughout their environment. In 2013, the CDC reported a multistate outbreak of Salmonella Typhimurium, and epidemiologic investigations linked this outbreak to
contact with pet hedgehogs purchased from multiple sources in several states. 11 35% of infected
people were hospitalized, and one death associated with Salmonella infection was reported in
Washington. In contrast with the recent Salmonella outbreaks in reptiles, only 35% of infected
persons were children 10 years of age or younger. As with reptiles, all hedgehogs should be
considered potentially infected with Salmonella, and the same precautions should be taken to
prevent infection. Also similar to reptiles, intermittent shedding can lead to false-negative fecal
cultures, and treatment to eliminate the carrier state is not recommended as antibiotic resistance
could result.
In June 2014, the CDC reported 41 human cases of Salmonella Typhimurium linked to contact with
frozen feeder rodents packaged by Reptile Industries, Inc. 12 These included rats, mice, and “other
rodents” of all ages. 16% of ill persons were hospitalized, but no deaths were reported. This same
outbreak strain was also was implicated in a 2009 outbreak in the United Kingdom and a 2010
outbreak in the United States, both linked to frozen feeder rodents from a single U.S. supplier. 13
These outbreaks highlight an important point—contact with both live and frozen-thawed rodents
are risk factors for Salmonella infections. Everyone should always wash their hands thoroughly
following handling of these animals.
Streptobacillus moniliformis
Streptobacillus moniformis is a gram-negative rod and the cause of rat bite fever in the United States.
Spirillum minus is the causative agent of rat bite fever in African and Asian countries. Both wild and
domestic rats harbor this bacteria in their nasopharynx, middle ear, and respiratory tract but it also
can be present in their blood and urine.14 Rats are asymptomatic for this disease. Mice and gerbils
can also carry this pathogen, but less commonly. Most human cases arise following a bite or scratch
from an infected rodent, but infections can occur following routine handling and fecal-oral
contamination. Initial clinical signs in humans occur within 7 days of exposure and include fever,
chills, headache and vomiting. A maculopapular rash on the extremities or septic arthritis may also
develop if the disease is allowed to progress. The reported mortality rate is 7-10% in untreated
patients.15
In August 2013, a 10 year old boy from San Diego, who also owned two rats, died of S. moniliformis.16
Two days prior to his death, he was examined by his physician for fever, vomiting, headaches, and
leg pain. A diagnosis of viral gastroenteritis was made, and he was prescribed anti-nausea
medication. These signs progressed in the next 24 hours to weakness, confusion, and collapse.
Oropharyngeal swabs from one of his pet rats were positive for the same strain of S. moniliformis that
was found on the child’s autopsy.
Rat bite fever is a rare disease in the United States, with only 17 cases identified in San Diego
County over the last 12 years. The national infection rate is unknown, as it not a reportable disease.
The median patient age was 10 years and 94% of infections were pet-associated. Sixteen of the
seventeen cases reported exposure to pet rats, and interestingly, only 38% of those patients reported
being bitten by the rat(s). Handwashing is always recommended after handling pet rats, and medical
care should sought if any symptoms of rat bite fever are noted after contact with pet or wild rats.
Giardia
Giardiasis causes gastroenteritis in numerous vertebrates, including humans. Potentially zoonotic G.
duodenalis has been reported in rabbits, guinea pigs, and chinchillas. 17 A recent study examined a total
of 1180 fecal samples from pet rabbits, chinchillas, and guinea pigs in multiple European countries
from 2006-2012 for Giardia by coproantigen ELISA.18 7.6% of rabbit, 61% of chinchilla, and 4% of
guinea pig samples were found to be positive. The majority of isolates from chinchillas had Giardia
duodenalis sequences similar to those associated with human infections. Although this report does
not make any direct connections between the positive rodent fecal samples and human disease, it
does indicate these species, especially chinchillas, could act as a reservoir of Giardia.
Rabies
There are no reported cases of rabies in humans linked to exposure to rabbits and rodents.
However, small numbers of these species do test positive for the virus on an annual basis. In 2005,
a case series described 7 pet domestic rabbits, and one guinea pig positive for the rabies virus. Half
of these animals had known exposure to raccoons and all cases originated from the northeastern
United States.19 A recent cross-sectional study reviewed all records of rodents and lagomorphs (both
wild and domestic) submitted to all state laboratories for rabies testing from 1995 through 2010. 20
737 rabid rodents and lagomorphs were reported, which was a 62% increase from the previous 15
years. The most commonly reported species was the groundhog, and all cases were positive for the
raccoon rabies variant. The positive cases also included one chinchilla, one guinea pig, and 25
domestic rabbits. This report should function as a reminder that all mammals can carry rabies, and
any owner bitten by a pet (or wild) mammal should seek medical attention from a physician or
public health official.
REFERENCES
1.
Mitchell MA. Zoonotic diseases associated with reptiles and amphibians: an update.
Veterinary Clinics of North America: Exotic Animal Practice. 2011;14(3):439-456.
2.
Notes from the field: update on human Salmonella Typhimurium infections associated with
aquatic frogs--United States, 2009-2011. MMWR. Morbidity and mortality weekly report.
2011;60(19):628.
3.
Zehnder AM, Hawkins MG, Koski MA, et al. Burkholderia pseudomallei isolates in 2 pet
iguanas, California, USA. Emerging infectious diseases. 2014;20(2):304.
4.
Elschner MC, Hnizdo J, Stamm I, El-Adawy H, Mertens K, Melzer F. Isolation of the highly
pathogenic and zoonotic agent Burkholderia pseudomallei from a pet green Iguana in
Prague, Czech Republic. BMC veterinary research. 2014;10(1):283.
5.
Giacomelli M, Piccirillo A. Pet reptiles as potential reservoir of Campylobacter species with
zoonotic potential. Veterinary Record. 2014.
6.
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Wang C-M, Shia W-Y, Jhou Y-J, Shyu C-L. Occurrence and molecular characterization of
reptilian Campylobacter fetus strains isolated in Taiwan. Veterinary Microbiology.
2013;164(1):67-76.
Patrick ME, Gilbert MJ, Blaser MJ, Tauxe RV, Wagenaar JA, Fitzgerald C. Human infections
with new subspecies of Campylobacter fetus. Emerging infectious diseases. 2013;19(10):1678.
Santos F, Leal D, Raso T, et al. Risk factors associated with Chlamydia psittaci infection in
psittacine birds. Journal of medical microbiology. 2014;63(Pt 3):458-463.
Raso Tania Freitas, et al. Chlamydophila psittaci infections in hyacinth macaws
(Anodorhynchus hyacinthinus) confiscated in Brazil. Journal of Zoo and Wildlife Medicine.
2013;44(1):169-172.
Swanson SJ, Snider C, Braden CR, et al. Multidrug-resistant Salmonella enterica serotype
Typhimurium associated with pet rodents. New England Journal of Medicine. 2007;356(1):21-28.
Notes from the field: Multistate outbreak of human Salmonella typhimurium infections
linked to contact with pet hedgehogs-United States, 2011-2013. MMWR. Morbidity and
mortality weekly report. 2013;62(4):73.
Notes from the field: Infections with Salmonella-linked to exposure to feeder rodentsUnited States, August 2011-February 2012. MMWR. Morbidity and mortality weekly report.
2012;61:277.
Harker K, Lane C, De Pinna E, Adak G. An outbreak of Salmonella Typhimurium DT191a
associated with reptile feeder mice. Epidemiology and infection. 2011;139(08):1254-1261.
Hill WA, Brown JP. Zoonoses of rabbits and rodents. Veterinary Clinics of North America:
Exotic Animal Practice. 2011;14(3):519-531.
Elliott SP. Rat bite fever and Streptobacillus moniliformis. Clinical microbiology reviews.
2007;20(1):13-22.
Adam JK, Varan AK, Pong AL, McDonald EC. Notes from the Field: Fatal Rat-Bite Fever
in a Child‚ San Diego County, California, 2013. MMWR. Morbidity and mortality weekly report.
2014;63(50):1210-1211.
Lebbad M, Mattsson JG, Christensson B, et al. From mouse to moose: multilocus
genotyping of Giardia isolates from various animal species. Veterinary parasitology.
2010;168(3):231-239.
Pantchev N, Broglia A, Paoletti B, et al. Occurrence and molecular typing of Giardia isolates
in pet rabbits, chinchillas, guinea pigs and ferrets collected in Europe during 2006-2012. The
Veterinary record. 2014;175(1):18-18.
Eidson M, Matthews SD, Willsey AL, Cherry B, Rudd RJ, Trimarchi CV. Rabies virus
infection in a pet guinea pig and seven pet rabbits. Journal of the American Veterinary Medical
Association. 2005;227(6):932-935.
Fitzpatrick JL, Dyer JL, Blanton JD, Kuzmin IV, Rupprecht CE. Rabies in rodents and
lagomorphs in the United States, 1995-2010. Journal of the American Veterinary Medical
Association. 2014;245(3):333-337.
CLINICALLY IMPORTANT AVIAN AND REPTILIAN ANATOMY
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
AVIAN ANATOMY
Most psittacines are not sexually dimorphic—some classic exceptions include budgerigars,
cockatiels, and eclectus parrots. Male anseriform (ducks and geese) species posses a phallus. If you
cannot determine the sex of the bird externally, endoscopic/surgical visualization of the gonads or a
blood test are other methods. Most birds possess a keel which is a ventral enlargement of the
sternum for the attachment of the flight muscles. These muscles can be used to determine the bird’s
body condition score (BCS). Birds have several other skeletal modifications for flight, including
pneumatized bones, fused vertebrae, and fused wing bones. The muscular attachments to the keel
and vertebral ribs move these bones cranially and ventrally during inspiration. This helps to expand
the air sacs, as there is no diaphragm. Restriction of this keel movement will hinder breathing, and
this should be taken into consideration with restraint of any avian species.
Dorsal recumbency is preferred for anesthetized birds, as this will not inhibit movement of the keel,
and leads to the greatest possible air sac volume. Birds lack an epiglottis, which predisposes them to
aspiration. They have complete tracheal rings, and very long tracheas relative to body size. Birds
compensate for this increased length and dead space (4x greater than a mammal of comparable size)
by decreasing respiratory rate (1/3 that of a mammal of comparable size) and increasing tidal
volume (4x as high as a mammal of comparable size). The syrinx is the site where the trachea
bifurcates, and is responsible for voice production in birds. Cole or non-cuffed tubes should be
used for intubation. Transtracheal membranes and tracheal strictures are possible complications of
intubation, particularly in long-necked birds. Psittacines, and most birds, have 9 air sacs—2 cervical,
1 clavicular, 2 cranial thoracic, 2 caudal thoracic, and 2 abdominal. No gas exchange occurs here,
and the air sacs function as a bellows to help move air through the respiratory system. The air sacs
are connected to the lungs via ostia and to the pneumatized bones, such as the humerus, femur, and
vertebrae. Positive pressure ventilation is recommended during anesthesia to assist with ventilation,
as the weight of the bird’s viscera, regardless of how they are positioned, will compress and
eventually collapse the thin-walled air sacs.
Subcutaneous fluids can be safely administered in the inguinal area in most birds and over the
dorsum in waterfowl and galliformes (chickens, turkeys). IO fluids can be administered to the ulna
and tibiotarsus but should never be given into pneumatized bones such as the humerus and femur.
Birds breath in a four-part cycle—air is inhaled into the trachea and then enters the caudal air sacs,
to the lungs for gas exchange, to the cranial air sacs, and then exhaled out through the trachea. The
lungs are affixed to the ribs, and do not expand. There is no pleural space, and the lungs are not
divided into lobes or lobules. Secondary bronchi divide into parabronchi (functional unit of the
lung), and then into air capillaries, which are the site of gas exchange and equivalent to mammalian
alveoli. Air capillaries are 3-10 micrometers in diameter, whereas the smallest mammalian alveoli is
35 micrometers. The avian lung is the most efficient gas exchange system among all air-breathing
vertebrates. Why? They have continuous, and unidirectional air flow through the lung which means
less anatomic dead space in their lungs. There is a cross-current exchange between blood and air
capillaries, which means birds need less ventilation than mammals to achieve the same level of
oxygenation in the lungs. The diameter of their terminal airways is very small, and they have an
increased total anatomic diffusing capacity of the lung for oxygen (approximately 20% greater than
mammals).
Approximately 1% of a bird’s total body weight can be safely removed during routine venipuncture,
assuming the bird is healthy and euvolemic. (1mL per 100gm) A recent study examined the effects
of pre-heparinzing syringes prior to venipuncture. They found that this procedure significantly
lowers the mean PCV and total solids compared with controls (up to 23%). This dilution effect was
inconsistent between samples. The jugular vein (right jugular for most species), medial metatarsal
vein, or ulnar vein are commonly used for venipuncture. Blood should never be obtained from
clipping a toe nail. This is very painful, and can lead to spurious bloodwork results. IV catheters
can be placed in the ulnar, medial metatarsal, or jugular veins. Considerations for IV placement
include the sedation/anesthesia that is often required for placement, and an E-collar to maintain the
catheter (which can lead to anorexia in most species).
Birds have paired kidneys, embedded within the renal fossa of the synsacrum (fusion of pelvic
bones). In most birds, including psittacines, each kidney is comprised of a cranial, middle, and
caudal division. These are each supplied by one of three renal arteries. This makes nephrectomy
very difficult if not impossible. The sciatic nerves run through the parenchyma of each kidney;
therefore, lameness or neurologic deficits to a pelvic limb can be due to renal masses. Avian kidneys
lack a renal medulla and pelvis. They contain two types of nephrons—mammalian (10-30% of total
nephrons) and reptilian, which lack a loop of Henle. All birds lack a urinary bladder and urethra.
The ureters connect directly to the cloaca, and urine is then refluxed into the colon for water
reabsorption. Due to all of these anatomic differences, birds cannot concentrate their urine to the
same degree as mammals, and there is significant post-renal modification of urine (urine samples are
never sterile). Birds are uricotelic, meaning their main nitrogenous waste is uric acid which is seen
clinically as the white urates within the dropping. Birds do produce urea (BUN) but in very small
amounts. When uric acid becomes significantly elevated, as in renal disease, it can precipitate out of
the blood and form gout, both visceral (on organs) and articular (within the joints).
REPTILIAN ANATOMY
Sex determination in reptiles varies by species. Snakes are commonly sexed by probing into the
hemipene pockets that open into the cloaca—if the probe extends >5 scutes distal to the cloaca, it is
likely a male. Most male lizards will have a bulge, from their hemipenes, on either side of the
proximal tail, just caudal to the vent. Male lizards including the green iguana, Chinese water dragon
and bearded dragon also have larger femoral pores (located on the medial/ventral surface of the
proximal pelvic limbs) than females. Male box turtles have bright red irises, whereas females have
brown irises. Some species of chelonians (turtles and tortoises) have changes in their plastron
(concave for males), tail length (males usually longer), and some male aquatic turtles (red-eared
sliders, etc.) have longer claws on their front limbs when compared to their hind limb claws.
Certain lizard species (most iguanas and geckos, but not chameleons and bearded dragons) have tail
autotomy, which is the ability to consciously “drop” their tails when stressed or when their tail is
grasped (either by a human or a predator). Pre-determined fracture planes are located within the
vertebral bodies of the tail. The tails often grow back, but may be smaller and discolored compared
with the remaining tail. Reptiles display three different types of dentition: acrodont (teeth attached
to the crest of the bone; chameleons, dragons), pleurodont (teeth attached to the labial surface;
snakes, some lizards [iguanas]), and thecodont (teeth held within a deep bony socket, crocodilians).
All of the teeth are replaced rapidly throughout life, with the exception of reptiles with acrodont
dentition, which will not regrow.
The kidneys of most lizards are located within the pelvic canal—varanids (monitor lizards) have
kidneys located in the dorsal-caudal coelom. Some reptiles have bladders including all chelonians,
some lizards, but no snake species. The ureters connect the kidneys to the urodeum in the cloaca,
and then urine is moved to the bladder, if present. Therefore, reptile urine, even if obtained from a
cystocentesis is not sterile. Terrestrial reptiles produce uric acid as their main form of nitrogenous
waste, similar to birds. In comparison, aquatic turtles and other highly aquatic species do produce a
substantial amount of BUN. The reptilian nephron lacks a loop of Henle, renal pelvis, and they only
have several thousand nephrons (most mammals have millions of nephrons). Similar to birds,
reptiles cannot significantly concentrate their urine and rely on other methods for water
conservation. Reptiles have a renal portal system, which blood from the hind limbs and tail can flow
through the kidneys prior to reaching systemic circulation. There are valves within the vessels that
can shunt blood to or away from the portal system. This is thought to help reduce GFR (glomerular
filtration rate) during reduced water availability. Clinically, this is significant because drugs injected
in the hind limbs or tail may be filtered through the kidney prior to reaching systemic circulation.
This is particularly important for drugs with potential nephrotoxicity, such as aminoglycosides. In
addition, drugs delivered in the caudal half of the body may diverted to the hepatic portal system.
Shunting of blood through these two portal systems is variable species to species, but as a general
rule, one shouldn’t give any injections to reptiles in the caudal portion of their body.
The hearts of snakes, lizards, and chelonians contains two atria and a single ventricle. Therefore,
there is mixing of arterial and venous blood. They also possess two aortic arches. In most lizards,
the heart is located cranially, between the pectoral girdle. The varanid heart is located more caudal,
similar to a mammal. The chelonian heart is located on ventral midline, where the humeral, pectoral,
and abdominal plastron scutes intersect. Only 5-8% of blood volume can be safely drawn from a
healthy reptile during venipuncture, equating to 0.5-0.8% of body weight, or 0.5-0.8mL per 100gm
body weight. Several species, including most chelonians, exhibit RBC lysis in EDTA; therefore,
lithium heparin is the anticoagulant of choice. Blood can be obtained from the ventral tail vein or
via cardiocentesis in snakes, through the ventral tail, ventral abdominal, or jugular vein in lizards, and
from the brachial plexus, dorsal tail vein, jugular vein, or subcarapacial sinus. Clinical abnormalities
including forelimb, hindlimb, or tail paresis have been seen following subcarapacial venipuncture
and injections; therefore this venipuncture site should be used with caution in chelonians. IV
catheters can be placed in the ventral tail veins of snakes and lizards, or the jugular vein of
chelonians.
Crocodilians and chelonians have complete tracheal rings, and squamates (lizards and snakes) have
incomplete tracheal rings. Their lungs are very simple and saclike—the internal structure is a
honeycomb of faveoli, which are similar to mammalian alveoli. Reptiles have a large lung volume,
but only about 1% of the lung surface area compared to mammals. Reptiles lack an epiglottis, and
their glottis is located rostral in the oral cavity, at the base of their tongue. Non-cuffed ET tubes are
often used for intubation.
Reptilian cardiovascular physiology is much different that mammals, and these differences should be
taken into consideration for general anesthesia. There is mixing of oxygenated and deoxygenated
blood in the heart, and they are ectothermic (proper heat support is required). Many species of
reptiles, especially aquatic species and chelonians, can hold their breath for extended periods of time.
During which, they can utilize anaerobic metabolism and can shunt blood away from their lungs.
Their stimulus to breath during these breath-holding events is low oxygen concentrations, rather
than elevated carbon dioxide concentrations, as in mammals. During anesthesia, an ultrasonic
Doppler flow device is often used to monitor heart rate and rhythm. An ECG and venous blood
gas can also be utilized. The following monitoring techniques are NOT accurate in any reptilian
species: mucous membrane color, indirect blood pressure, pulse oximetry, and capnography.
During anesthetic recovery, it is important to keep a reptile close to their POTZ (Preferred
Optimum Temperature Zone), as a cold reptile will recover much slower. If intubated, they should
be transitioned to room air, and positive pressure ventilation provided with an Ambu bag, rather
than oxygen for reasons listed above.
PET RABBIT HUSBANDRY
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
Rabbits are in the order Lagomorpha, and are differentiated from rodents by the presence of a
second pair of incisors, called peg teeth. This order contains three genera: oryctolagus (the
European or true rabbit, contains all pet rabbits), lepus (hares, jackrabbits), and sylvilagus
(cottontails). There are approximately 50 breeds of rabbits, including those for pets, meat, wool, and
research. Rabbits are currently the third most popular pet in the US, thanks in part to the House
Rabbit Society, which was established in 1988. The average rabbit lifespan is 8-10 years, with smaller
sized rabbits living longer than their larger cousins.
Females = does
Males = bucks
Young = kits
Giving birth = kindling
ANATOMY
The ears comprise 12% of a rabbit’s total body surface, and have a thermoregulatory function.
Rabbits lack footpads on their hindlimbs, which can predispose them to pododermatitis if housed
on rough or hard surfaces. They possess inguinal, anal, and chin scent glands, but do not have any
sweat glands. Rabbits are obligate nasal breathers and have a very narrow oropharyngeal cavity,
similar to horses. Compared to their total body size, they have a small thoracic cavity. Rabbits have
an extremely light skeleton and vertebral column compared with their large muscle masses of their
legs. This helps them to escape from predators in the wild, but also predisposes them to fractures
(sometimes iatrogenic in the case of vertebral fractures). Their skeleton represents only 7-8% of
their total body weight, whereas the skeleton of a cat comprises 12-13% of body weight.
All of their teeth are hypsodont (high-crowned) and open-rooted, which means they grow
continuously throughout life. Their first incisors grow between 2-3mm per week. Rabbits are
considered hind-gut fermenters, and they have an enlarged cecum and colon to facilitate
fermentation of feed-stuffs. They produce two types of feces—normal, dry pelleted feces and soft,
dark cecotrophs, which are meant to be ingested. If the cecotrophs are not ingested, they will often
adhere to the perineal area, and may be mistaken for diarrhea. Rabbits have a unique calcium
metabolism, in that, they absorb all of the calcium they ingest, a majority of which is excreted in the
urine. Therefore, their urine can vary in color and clarity—it is often opaque.
Rabbits have a biocornutate uterus, with two cervices—one for each uterine horn. Adult female
intact rabbits have very high incidence of uterine diseases including uterine adenocarcinoma,
endometrial hyperplasia, and endometritis. Rabbits of certain breeds which are >4 years of age have
an incidence of uterine adenocarcinoma of 50-80%. Therefore, all female rabbits should be spayed
when they are less than 1 year of age to prevent disease, as well as unwanted pregnancy. All rabbits
have open inguinal rings, which is very important to note during castration. The testes descend
when they are 10-12 weeks of age, and they may attempt to breed as young as 3.5-4 months of age.
Does become sexually mature at 4-6 months of age.
RESTRAINT
Many people are hesitant to restrain rabbits due to the risk of vertebral fracture. While uncommon,
this still is a risk if a rabbit is improperly restrained. One should always support the lower back, and
ensure that the rabbit’s powerful hindlimbs are not able to come into contact with a surface with
which to jump from. Performing procedures or even a physical exam in a quiet room without the
smell of predators greatly decreases stress for this prey species. Also, covering the eyes of the rabbit
(such as with a light towel), will also reduce the stress of handling/restraint for most patients. When
returning the rabbit to its enclosure (cage, carrier, etc.), it is recommended to place the rabbit in head
last (hind-end first), so it cannot see where it is going. Often, if they see an escape (such as their
carrier), rabbits will jump or struggle, sometimes without warning, to get to that area. This can lead
to injuries of both the rabbit and the handler if he/she is unprepared. If the rabbit is fractious, or
resisting manual restraint, then chemical restraint and sedation should be utilized to prevent injury
and undue stress to the patient.
DIET
Rabbits require a high fiber, low calcium diet. The majority of their daily diet should be comprised
of a high quality grass hay (such as timothy hay) and fresh greens. The House Rabbit Society
recommends to feed one cup of greens for every 2 lbs. of body weight, once daily or divided into
multiple feedings. Pellets are not required, but can be added to the diet at a rate of ¼ cup per ~5lbs
of body weight, divided over two meals and offered for 5 days per week. Excessive pellets often
leads to obesity, as pellets were designed to feed meat rabbits, so they would gain weight quickly.
Juveniles and pregnant/lactating does should always be provided pellets due to their increased
energy demands.
Many rabbit owners are hesitant to provide certain greens and vegetables, such as cruciferous
veggies (broccoli, cabbage, kale, bok choy, collard greens, and cauliflower) due to concerns with
increased gas production and formation of goiter. If ingested in large quantities, goitrogens within
these plants can inhibit incorporation of iodine into thyroid hormone, and lead to goiter (in people).
This has never been proven in rabbits, and they are considered safe to feed in moderation. Also,
certain greens are high in oxalic acid, which can be a problem in people with hypercalcemia or oxlate
bladder stones. Although never proven in rabbits, it is recommended not to feed these greens in
large quantities or exclusively. These include: spinach, Swiss chard, beets (roots), beet greens
(leaves), collards, okra, and parsley. Another owner concern is providing vegetables that cause gas in
people, such as broccoli. Rabbit’s GI tracts are much different than humans, and many of the foods
that may cause excessive gas production in people may not do so in rabbits. Food items high in
starch and sugar (including carrots and fruits) can lead to GI disease and stasis secondary to
alterations in cecal flora. Therefore, these items should only be given sparingly to rabbits.
Examples of healthy greens for rabbits
Kale
Mint
Clover
Endive
Parsley
Cilantro
Chicory
Escarole
Raddichio
Carrot tops
Radish tops
Beet greens
Swiss chard
Watercress
Wheat grass
Collard greens
Broccoli leaves
Mustard Greens
Romaine lettuce
Dandelion Greens
Green/Red leaf lettuce
HOUSING
All rabbits should have a minimum of 4 hours of exercise per day, as obesity is common in house
rabbits. The larger house the better! Their primary enclosure should be no less than than 4-5 times
the rabbits length (for the length of the cage) and no less than twice the rabbits length (for the
width). Metal and plastic cages are preferred over wood, as it is impossible to disinfect and rabbits
will chew on it, weakening its structural integrity. Solid flooring is always the best. Rabbits, especially
obese rabbits sitting on wire mesh have a much higher incidence of pododermatitis (“sore hocks”)
than rabbits housed on solid flooring. Toenails can get caught and avulsed easily on mesh bottom
caging. The HRS has a nice listing of appropriate sized cage companies that are easy to understand
for your clients.
If a rabbit is free roaming in the house, it should be “bunny proofed.” Special attention should be
made to items which they could ingest (carpeting, cardboard, etc.) which may lead to GI obstruction
House plants (which may be toxic if ingested), should also be removed from the rabbit’s reach.
Electrical wires and cords should be protected to prevent electrocution. As with other small
mammals, recycled paper bedding is preferred over wood shavings. Rabbits can be trained to use a
litter box, and owners are encouraged to provide one and train their rabbit to help with waste cleanup. Some rabbits will eat paper bedding, and hay or straw can be used as an alternative in these
cases.
If a rabbit is housed outside, there are several considerations for the hutch design. First and
foremost, it should provide protection against predators (both from the air and on the ground!).
These enclosures should provide shade and good ventilation. Rabbits tolerate cold better than
heat—they have limited sweat glands, and are obligate nasal breathers, therefore, they cannot
effectively pant to release body heat. Ideally, rabbits should be brought indoors during the night,
inclement weather, and if they are ill.
FERRET MEDICINE 101:
PHYSICAL EXAM AND CLINICAL TECHNIQUES
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
The ferret is a domestic animal descended from either the European or Siberian polecat. Ferrets belong to
the family Mustelidae which also includes other weasels, mink, otters and skunks. The fitch ferret as it
became known, was used for hunting rodents and rabbits and farmed for its fur. There are many color
variations, including, but not limited to: sable, “fitch,” with the clearly defined mask and dark eyes, pastel,
cream with light colored mask and dark eyes, Siamese with brown guard hairs, cinnamon with reddish
guard hairs, Albino (white) with no mask or patterning and red eyes, Silver mitt (white or silver fur, black
eyes, usually white feet), and Gnome (brown or pastel with four white paws and throat patch). A recent
study found an association between certain coat colors and deafness in this species. Most ferrets now sold
for pets in the U.S. have been spayed/neutered and descented at 4-6 weeks of age, these ferrets can be
identified as having 2 tatoos in the right ear. Ferrets are not legal in all states and municipalities. For
example, it is still illegal to own a pet ferret in California and Hawaii as well as in municipalities such as
New York City and Salt Lake City. In other areas a permit is required for ownership.
GENERAL BIOLOGY
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Life span for pet ferrets in the US is approximately 5-7 years (UK appx. 8-10 yrs)
Adult males weigh 0.8-3.0 kg, adult females 0.6-1.0 kg
Heart rate range 180-240 bpm, respiration rate is 33-36 bpm, blood volume is 60-80 ml/kg
Body temperature ranges from 100-104.1oF (excited), average = 102oF.
First breeding usually occurs the first spring after reaching adult body weight
Females are polyestrous and induced ovulators, breeding year round and rebreeding is immediate
after parturition
Length of gestation is 41-43 days. Litter size ranges from 7-14 (avg. 8)
Birthweight is 8.5 grams, ferrets wean at 6 weeks at approx. 0.2-0.4 kg but ferrets begin to eat solid
food at 14-21 days of age
SELECT ANATOMY/PHYSIOLOGY
Gastrointestinal: The dental formula of the ferret is 2(I 3/3 C 1/1 P3/3 M 1/2) and all the teeth are
brachydont (closed rooted). Ferrets are strict carnivores and are designed to eat whole prey. They have a
simple stomach that can enlarge to hold a large food bolus. They also have a very short GI tract, no
cecum, and minimal GI flora and brush border enzymes so they cannot digest fiber or efficiently use
carbohydrates. The maximum GI transit time in the ferret is 3-4 hours! Because of the minimal GI flora,
gastrointestinal upset with antibiotics is rare in ferrets.
Thoracic Cavity: The heart lies between the sixth and eighth ribs. There are six lung lobes: left and right
cranial and caudal, and right middle and accessory. The thymus is found within the thoracic inlet in the
cranial mediastinum and can be very prominent in the young ferret. The thymus is a common site of
neoplasia in ferrets < 1 year old.
Spleen/Liver/Pancreas: The spleen is located on the greater curvature of the stomach and can vary in
size depending upon age and health status. With splenomegaly, the size of the spleen extends from the
upper left to the lower right abdomen. The liver has six lobes and a gallbladder is present between the
quadrate lobe and right medial lobe. The pancreas is V-shaped with right and left limbs connecting at the
level of the pylorus. The right limb is longer than the left extending along the descending duodenum and
the left limb extends between the stomach and the spleen.
Adrenal Glands: The left adrenal gland lies in fatty tissue just medial to the cranial pole of the left kidney.
The right adrenal gland lies more dorsal and cranial to the left and is covered by the caudate lobe of the
liver. The right gland is intimately attached to the caudal vena cava. Accessory adrenal tissue is commonly
found in ferrets and often is directly adjacent to the glands. Several reports suggest that the size of these
glands can vary with side and by sex of the animal.
Urogenital tract: The cranial pole of the right kidney is covered ventrally by the caudate lobe of the liver.
Male ferrets have a small prostate gland located at the base of the bladder that surrounds the urethra.
Prostatic cysts are common in male ferrets and paraurethral cysts can occur in both sexes. Male ferrets
have an easily palpable J-shaped os penis which can complicate catheterization. Ferrets are easily sexed
based upon anogenital distance. Female ferrets have a slit-like vulvar opening in non-estrous females; this
tissue swells and protrudes during estrus and has a doughnut-like appearance.
Musculoskeletal: The ferret has a long, flexible spine with a vertebral formula of C7, T15, L5-7, S3,
Cd18. Some ferrets have 15 ribs on one side and 14 on the other. In ferrets, the thoracic inlet is bordered
by the first pair of ribs and the sternum and is very small. All four feet have five digits with non-retractable
claws.
Integument: Ferrets molt in the fall and spring with a concomitant change in body weight. The ferret
gains weight in the fall and will lose the weight in the spring as the mating season begins. Coat color
change can also occur with typically a lighter coat color in winter and darker in summer and ferrets may
lose their mask configuration from season to season and from year to year. Thus, photographs for
identification are unreliable in the ferret. Most ferrets in the US are ear-tattooed by the breeder. Males are
tattooed in the right ear; females in the left. The skin of the ferret is especially thick over the dorsal neck
where it protects the ferret during fights or during mating when aggressive biting of the back of the neck
occurs. Ferrets have active sebaceous glands, which accounts for their musky body odor. Ferrets have
well-developed paired anal glands which produce a liquid with a powerful odor. However, the serous
secretions from these glands are not responsible for the musky body odor of the ferret. Nonetheless,
virtually all the ferrets in the US are “descented” (anal sacculectomies) at 4-6 weeks of age.
DIET
It is recommended that the ferret diet contains 30-50% high-quality animal (not plant) protein, 20-40% fat,
<4% fiber and minimal carbohydrates. Excess dietary carbohydrates may affect the pancreas and it has
been suggested that this might contribute to Beta cell disease. Ferrets in nature would only encounter
carbohydrates found in the partially digested GI tracts of their prey. A whole prey diet or a freeze-dried
carnivore diet is fed in many parts of the world. However, the most common diets fed to pet ferrets in the
US are dry kibble. In the past, high quality cat foods were fed to ferrets. Unfortunately, most cat dry kibble
contains large amounts of grain for binders to hold the extruded pellet shape. Commercial diets specifically
formulated for ferrets have been developed (Totally Ferret® - Performance Foods, Inc.; Forti-Diet® Kaytee Products, Inc., Marshall Ferret Diet ® - Marshall Pet Products, Inc., Purina/Mazuri Ferret Chow® PMI Feeds/Mazuri Zoo Feeding), which minimize carbohydrates, but still contain some grains. Newer
diets (Pretty Pets Nature’s Gold Ferret ®, Natura’s Innova Evo Ferret®) are marketed as “grain-free”.
Because of the rapid GI transit time of the ferret, fasting of no more than 3 hours should be adequate for
blood glucose testing and no more than 6 hours should empty the GI tract for surgical procedures.
Caution must be taken with fasting ferrets older than 2 years as often there is pancreatic disease present
and any fast may result in a serious hypoglycemic condition.
RESTRAINT AND PHYSICAL EXAMINATION
Most ferrets are very docile and can be examined without assistance. However, training treats such as
laxatone, Ferratone, and Nutrical are indispensable tools for the physical exam. Ferrets have a very short
attention span, become wiggly, and may become impatient and bite. A small amount of Nutrical on a
tongue depressor will often distract the ferret long enough to perform your physical exam or give a
vaccination. However, these “treats” are pure carbohydrates, and should be used judiciously, especially
with an animal with a potential pancreatic beta islet cell tumor (insulinoma).
Two basic methods of restraint are utilized in the ferret. For tractable animals, these ferrets can be
restrained upright while supporting their hindquarters for most of the examination. For active or biting
ferrets, the “scruff” restraint is very effective. The ferret is scruffed at the same point of the neck as a cat
however, you should restrain the ferret with all four feet off the table. Ferrets relax in this hanging position
and will commonly yawn allowing for a full oral examination.
The physical examination follows that of other mammals. Closely examine the vulva in the female ferret
and point out any abnormalities to the client. It is also recommended to show the client the normal vulva
as well so that they can detect early signs of swelling or enlargement that might be associated with adrenal
disease. Also examine the tail tip for evidence of subtle alopecia. Seasonal alopecia of the tail tip can be a
normal, incidental finding but can also be the first signs of alopecia associated with adrenal disease.
Remember, hyperadrenocortisim in ferrets in associated with elevated sex hormones (zona reticularis in
the adrenal cortex) rather than glucocorticoids; therefore, the treatment in this species is completely
different than in dogs. Their heart is located more caudal in their chest cavity, rather than just behind the
shoulder, as in dogs and cats. Cutaneous neoplasia, especially mast cell tumors, are very common.
Therefore, close attention should be paid to their integument during the physical exam. Lymphoma is one
of the top three diseases in ferrets, and all the peripheral lymph nodes should be palpated in every exam.
Ferrets often have large accumulations of fat around their lymph nodes, which sometimes makes it
challenging to determine if lymphadenopathy is present. In the author’s experience, abnormal lymph
nodes in this species are usually firm in addition to being enlarged.
CLINICAL TECHNIQUES
Imaging
Ferret radiographic positioning and technique follows the principles of small animal radiology with the
exception that it is necessary to use high speed film and cassette combinations. Also, because of the
ferret’s small size, whole body radiographs can be taken allowing evaluation of both the soft tissues (in
both the thorax and abdomen) and the skeletal structures.
Thoracic radiology should always include the entire thorax as mediastinal neoplasia is common, especially
in the young ferret. The shape of the normal ferret heart is more globoid than a dog or cat and is located
between the sixth and eighth intercostal spaces. The heart silhouette is not in contact with the sternum.
Loss of this is often an early sign of heart disease. The vertebral heart score used in dogs and cats to
determine heart size has been modified to a ferret specific scale, to take into account the small, laterally
compressed thorax of the ferret.
Abdominal radiography is evaluated as in other carnivores such as the domestic cat. If prostatic or
paraurethral cysts are suspected, they are often identified on a lateral radiograph as a round mass just
dorsal to the neck of the bladder. The spleen is commonly enlarged in ferrets and this is not always
associated with disease. Ultrasound is utilized as with other small mammals. This imaging modality is very
useful in the diagnosis of adrenal disease, gastrointestinal neoplasia, cardiac disease and other disorders in
the ferret. Computed tomography, MRI and nuclear scans have been utilized for various ferret diseases.
Blood Collection
Blood collection can be challenging in a wiggling, overweight ferret. The jugular, cephalic, and lateral
saphenous veins can be accessed for venipuncture. Small volumes (0.3mL or less) can be obtained from
the cephalic and lateral saphenous, but if larger amounts of blood are required, the jugular or cranial vena
cava are recommended. The technique for jugular venipuncture is similar to the cat. The forelegs are
extended over the edge of a table and the neck extended dorsally. The neck may need to be clipped to
locate the jugular vein, and it is more laterally located than in most cats and dogs. Unfortunately, the awake
ferret wiggles considerably and if the neck is fat, this may be difficult to perform without sedation. The
cranial vena cava has also been utilized extensively for venipuncture in ferrets. Sedation or general
anesthesia is required to ensure iatrogenic trauma does not occur. Access to the cranial vena cava is via the
“sternal notch” to the jugular as it joins with the opposite jugular and brachial veins to become the cranial
vena cava. The ferret is restrained on its back with the forelegs held against its sides and the head
extended. The needle is inserted into the thoracic cavity between the first rib and the manubrium at a 30450 angle to the body at an angle toward the opposite rear leg. Risk of inadvertently hitting the heart is
minimal due to its caudal position within the chest.
Injection Sites
With the exception of the tough skin over the dorsal scapular area, injection sites in ferrets are similar to
those in other small mammals. However, subcutaneous administration of many medications is preferred
over intramuscular because of the limited overall muscle mass. Depending on the volume of intravenous
medication, the cephalic or the lateral saphenous are the veins of choice for indwelling intravenous
catheters. The jugular vein is difficult to catheterize percutaneously. Intraosseous catheters in the greater
trochanter of the femur may be used if larger volumes or prolonged use is anticipated. Sedation and/or
anesthesia is often required for catheter placement, and a small cut down in the skin overlying the vessel is
recommended. Fluids can be administered subcutaneously or intravenously. Vascular access ports (VAPs)
can be placed into the jugular vein of the ferret for long-term blood collection or for administration of
medications, such as chemotherapy.
Nutritional Support
Assisted feeding is required in any ferret that has been anorectic for 24 hours or more. It is imperative to
ensure the anorectic ferret maintains a positive energy balance as negative energy balances begin to occur
rapidly due to their short GI tract transit time. Syringe feeding in ferrets can be performed relatively easily,
with formulations made depending on illness. Oxbow Carnivore Critical Care® or Lafeber’s carnivore
powdered diet are recommended for syringe feeding ferrets. Partial parenteral nutrition (PPN) has been
formulated for ferrets as well. Percutaneous gastrotomy tubes have been utilized in ferrets but there is little
information available as to tolerance or practicality. Multiple meals (3-4) of 5-10 mL are often accepted
comfortably.
Urine Collection, Urinary Catheterization
Urine samples can be obtained either via cystocentesis, free catch or gentle manual expression. These
techniques are the same as in cats. Anesthesia is recommended for fractious ferrets to prevent trauma to
the thin bladder wall. Urinary catheterization is often indicated in male ferrets; indications in female ferrets
are rare. Placing a urinary catheter in a male ferret is difficult as the urethral opening is very small and
located on the ventral surface of the penis, below the hook in the end of the os penis. Often a magnifying
loupe is necessary to visualize the opening. 3.0 and 3.5 Fr urinary catheters are often used.
PREVENTATIVE CARE
Vaccinations
Canine distemper: Ferrets are exquisitely susceptible to this disease, with a fatality rate of ~100%.
Clinical signs are the same as in canids and include diarrhea, nasal/ocular discharge, dermatitis and
hyperkeratosis of the footpads and chin. There were only two approved distemper vaccines for ferrets:
Merial: PureVax and Fervac-D (United Vaccine). PureVax is a recombinant vaccine, is more
expensive than Fervac, but appears to have potentially fewer vaccine reactions. United vaccine, the
company that manufactured Fervac-D, closed in 2007 and the vaccine became unavailable indefinitely.
PureVac ferret distemper vaccine has been on indefinite backorder for years. Currently, many ferret
rescues are using a modified live Canine distemper-parvo combination vaccine (Nobivac), and have not
had any reversions to virulence documented to date. Use of a modified live distemper vaccines in any
species other than domestic dogs is always a risk, but this has been used out of necessity due to lack of
other available recombinant vaccinations. The author has used this vaccination in several ferrets with no
apparent side effects, but this species is not seen commonly in California.
Rabies: There is only one approved rabies vaccine for ferrets- Imrab. The first vaccine is given at 3
months SC then an annual booster is recommended. Studies are ongoing as to efficacy in ferrets and
shedding patterns, although it appears that ferrets rarely shed the virus in saliva and the public health risk
from a ferret infected with rabies appears to be negligible. Guidelines for injection sites are following those
being recommended for cats.
An anaphylactic-like reaction to Fervac-D has been frequently reported, and more recently, a few have
been noted with PureVax and Imrab. If mild, clinical signs may only be erythema and pruritus but if
more severe, can include dyspnea, hypersalivation, tremors or seizures, urination/defecation, diarrhea,
pyrexia and panting. Because of the significant numbers of vaccine reactions documented in ferrets, it is
recommended that all ferrets should also be kept at the clinic and observed for at least 20-30 minutes postvaccination. Treatment for a vaccine reaction includes parenteral diphenhydramine, steroids (prednisone
or dexamethasone), supplemental oxygen, fluids and other supportive care as needed. The clinician must
weigh the risk:benefit from vaccinating animals that have had a vaccine reaction. The choices are to not
vaccinate an animal with a potential low exposure to the virus however these animals are extremely
sensitive to this virus. Most clinicians do not give distemper and rabies vaccines on the same day so that a
reaction can be specifically documented if it should occur. Ferrets that are known reactors regardless of
vaccine brand should be given diphenhydramine 20-30 minutes prior to vaccination, and the vaccine
changed if possible.
Heartworm Prophylaxis
Ferrets are susceptible to canine heartworm disease (Dirofilaria immitis). The clinical disease resembles that
in cats. More studies are needed in diagnosing this disease in ferret. Approximately 50% of positive ferrets
have microfilaria and often have a low female worm burden so may be falsely negative on antigen tests.
The IDEXX Feline Snap test appears to be the most sensitive diagnostic test. In areas with endemic
disease, ferrets are prophylactically treated with Ivermectin (Heartgard heartworm prevention at ¼ tablet
of the smallest sized canine tablet, or ½ feline tablet) or selamectin.
ANESTHETICS AND ANALGESICS
Sedation is often used for minor procedures, such as venipuncture and ultrasonography. The author
prefers 0.1mg/kg butorphanol and 0.2mg/kg midazolam, in combination, IM. This provides
approximately 30 minutes of sedation, and can be partially reversed with 0.01-0.02mg/kg flumazenil IM.
Most ferrets are anesthetized with inhalant anesthetics but it is also important to remember that premedications for analgesia should be given for any potentially painful procedure. Both sevoflurane and
isoflurane administered via chamber, then maintained by mask or via intubation are commonly used
inhalant anesthetics. Ferrets are most commonly intubated using 2-3.5 endotracheal tubes. Intubation is
straight-forward; ferrets are commonly used for training physicians in appropriate neonatal intubation.
The use of inhalant anesthestics has been shown to reduce the hematocrit of the ferret, sometimes by up
to 20% due to splenic sequestration of the red blood cells.
Ferrets are often very stoic and mask signs of pain. Opioid medications offer excellent analgesia for
moderate-to-severe pain and can reduce inhalant anesthetic concentrations during surgery. Commonly
used opioid analgesics in ferrets include oxymorphone, buprenorphine and butorphanol. Hyperthermia is a
common sequelae to using hydromorphone in this species. Gastrointestinal stasis associated with opioid
medications can occur in ferrets but is not as life-threatening as this can be in the small herbivores, such as
the rabbit. Non-steroidal anti-inflammatory medications (NSAIDs) can also be used in ferrets. It is
important to monitor the ferret carefully during NSAID therapy for signs of GI ulceration. Meloxicam is
the most commonly used NSAID in ferrets today.
COMMON DISEASES OF BACKYARD POULTRY
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
Backyard chickens, or “urban chickens” are becoming very popular in the United States. These are
often considered family pets, but are also utilized as food animals, in that the owners often consume
their eggs. Even if the eggs are not consumed, these birds are still considered food-producing
animals by the USDA, and are governed by the many of the same rules (feed additives, drugs) and
regulations that apply to pigs, cattle, and commercial chickens. Clinicians who treat chickens,
especially with any form of antibiotic, are encouraged to visit the FARAD website (Food Animal
Residue Avoidance Database, www.farad.org ) for additional details on what is and is not legal to
administer to this species. If there is not an FDA approved medication to treat that particular
condition, then is the client willing/able to make this bird a NON-food producing animal?
Clinicians are encouraged to obtain a signature from the owner which document this discussion and
their understanding. As in all other food producing animals, there are certain drugs that are
prohibited from use including flouroquinolones and anti-viral medications.
Many homeowners/neighborhood associations and city ordinances have rules regarding the housing
and ownership of chickens. Roosters in particular are commonly prohibited, mainly due to concerns
with noise violations. A recent survey (Elkhoraibi, et al) of backyard chicken owners found that
most owned fewer than 10 chickens, and had kept chickens for a relatively short period of time (<5
years). 95% were kept for egg/meat production, and 57% stated they were solely kept as pets.
Many of the respondents knew either little or nothing about exotic Newcastle or Marek's disease,
and most (61%) had not vaccinated their chickens against Marek's. The Internet was the main source
of information (87%), and minimizing predation was the most cited challenge (49%).
HISTORY AND PHYSICAL EXAM
As most chickens are flock animals, it is important to get information on the flock health in addition
to the individual chicken that is brought in for examination. In addition to the basic historical
questions for all birds, there are some that are specific to chickens including:
- What is the flock hierarchy and have there been any recent changes?
- Has this or any of the chickens been vaccinated for Marek’s disease?
- When did she lay last, and was the egg(s) normal in appearance?
- Does anyone ingest the eggs from this (or any) chicken in the flock?
Before a physical exam, it is important to observe the bird in the carrier or walking around the exam
room. Note the bird’s mentation, droppings (if provided), and the presence of a limp or wing
droop. Body conditions score (BCS) is measured the same way as in other birds—pectoral
musculature. Assess hydration status with the venous refill time of the ulnar vein (similar to other
birds) or the presence of stringy saliva. Auscultation and palpation of the coelom is performed
similar to other birds. The ventriculus is often more prominent in galliformes than in psittacines,
and should not be misinterpreted as a coelomic mass. Evaluation of the integument for
ectoparasites is also important, as mites are very common in chickens. If present, they are often
found around the tail base, vent, and uropygial gland.
COMMON DISEASES
Reproductive tract disease
This is the most common disease of chickens in the author’s current practice. Chickens can present
with a number of clinical signs related to reproductive tract disease, some of which are non-specific
including lethargy, decreased appetite, and diarrhea. They often will have varying degrees of
coelomic fluid accumulation and distension. Ovarian neoplasia, often with carcinomatosis, is quite
common in aged laying hens, and will lead to coelomic effusion. Salpingitis and oviductal impactions
are also common in older hens. If the oviduct is large enough, it is often palpable as a firm tubular
structure in the left dorsal coelom. Salpingohysterectomy will successfully remove the oviduct, but
the ovary is rarely removed due to risks for fatal hemorrhage. If coelomic effusion is present, the
risks of intracoelomic surgery are great due to the high likelihood of aspiration of coelomic fluid if
the airsacs are disrupted during the procedure. The are several publications describing the use of
deslorelin acetate, a GnRH receptor agonist, in chickens to stop egg production, and as a
chemopreventative treatment for ovarian neoplasia. One 4.7mg implant is reported to last
approximately 6 months in this species. This drug is not approved for any avian species, nor for any
food producing species in the US.
Trauma
This is the second most common disease of chickens in the author’s practice, and bites/attacks by
predators are the most common type of trauma. Many wounds are deep and the clinician should
investigate all wounds for potential penetration into the coelomic cavity or the respiratory system
(air sac) as these carry a much more guarded prognosis. Lavage and cleaning of the wounds with
any liquids should be performed with caution if the air sacs are thought to be involved. Systemic
antibiotics are indicated in these cases, ideally based on culture and sensitivity of the wounds.
Analgesic therapy is also extremely important, but the clinician should be aware that most are not
approved for use in chickens. Primary closure of the larger wounds is often required, and the
clinician may need to get creative with bandaging techniques! Following any trauma, it is imperative
the chicken is kept indoors until the wounds are healed to prevent myiasis.
Ectoparasites
Chickens may or may not be symptomatic for ectoparasites, and they are often incidental findings
on a physical exam. If present, clinical signs include scratching, feather loss, and skin erythema.
Mites and lice are the most common ectoparasites encountered, with fleas, bedbugs, and ticks being
less common. Treatment is difficult, especially if the owner wishes to keep the chickens organic and
not use pesticides. Diatomaceous earth is an organic alternative for treatment of ectoparasites in
poultry, but will not completely rid the birds of disease. Treatment of the whole flock and the
environment is also imperative to help reduce the overall parasite burden.
Upper respiratory tract disease
In addition to nasal discharge, chickens with upper respiratory tract disease often have infraorbital
sinusitis which presents clinically as firm swellings ventral to the eyes. Differnetials for swelling in
this location include bacterial infection (Pasteurella, Haemophilus, Pseudomonas, Klebsiella, Mycoplasma,
Chlamydia), fungal infection (Aspergillus sp, Cryptococcus), viral (avian influenza), or neoplasia. If
bacterial etiology is confirmed, surgical debulking is often required due to the caseous nature of
these lesions. Antibiotic therapy alone will not likely resolve the clinical signs once the sinusitis is
advanced.
Diarrhea
Differentials for a chicken with diarrhea include stress (transient, such as traveling to a veterinarian’s
office), dietary (indiscretion, increased fruits/veggies/greens), bacterial overgrowth (Clostridium spp),
parasitic (coccidiosis most common), or systemic disease. As in all species, ideally diagnostics would
be performed to rule in/out these causes; these include fecal flotation, and if negative then
bloodwork, +/- whole body radiographs. Supportive care should also be provided in the form of
fluids (subcutaneous or intravenous), and gavage feeding if anorexic.
Pododermatitis
This is a debilitating disease of many avian species, including poultry. There are a number of
proposed etiologies, and ultimately, it is likely a multi-factorial disease. The most important risk
factors seem to be obesity, uneven weight bearing on the feet (due to arthritis, trauma, etc), trauma
to the feet/digits, and inappropriate substrates.
Grading systems
• Grade I/III: Redness/flattening of scale
• Grade II/III: Ulceration +/- infection of deeper tissue
• Grade III/III: Chronic fibrosis/loss of normal use
Radiographs are recommended to rule out osteomyelitis for Grades II and III, and DEEP cultures
of the wounds are needed if there is a lack of response to antibiotics. There are numerous published
and proposed treatment options for pododermatitis in multiple avian species. In general, treatment
for Grade I lesions includes improved husbandry such as changing the substrate and ensuring a
clean environment. Grade II and III lesions often require surgical debridement, bandaging, systemic
antibiotics, and anti-inflammatories. Supportive bandaging and environmental changes are also
important in the more severe cases. Surgical options include implantation of antibiotic-impregnated
items (beads, gauze), regional limb perfusion, and VAC systems in addition to traditional
debridement. In general, prevention is a much better and easier option than treatment after the fact.
REFERENCES AND SUGGESTED READING
Greenacre, Cheryl B., and Teresa Y. Morishita, eds. Backyard Poultry Medicine and Surgery: A Guide for
Veterinary Practitioners. John Wiley & Sons, 2014.
Elkhoraibi, C., et al. "Backyard chickens in the United States: A survey of flock owners." Poultry
science 93.11 (2014): 2920-2931.
WHEN TO FLY TO THE ER:
COMMON AVIAN EMERGENCIES
Olivia A. Petritz, DVM, DACZM
ACCESS Specialty Animal Hospitals
Culver City, California 90232
[email protected]
HEMORRHAGE
Acceptable blood loss, either through venipuncture or hemorrhage, is approximately 1% of body weight in
a healthy bird, or 1mL per 100gm of body weight. The average cockatiel is 80-100gm. While most avian
species seen in private practice are significantly smaller than dogs and cats, birds in general have several
anatomic differences which compensates for their small size. On average, their PCV is higher (45-55%),
and their red blood cell lifespan is much shorter, with a half-life of 30-45 days. Experimentally, birds have
been shown to rapidly recover from experimentally induced blood loss. For example, after the removal of
about 30% of the estimated blood volume of Japanese quail (Coturnix coturnix japonica), erythrocyte
numbers recovered to baseline values within 72 hours.
One of the most common emergency presentations of hemorrhage in the avian patient is a broken blood
feather. A blood feather is a growing feather than contains an artery and vein within the shaft; therefore, if
this feather shaft is damaged, it can lead to significant hemorrhage. On initial presentation, the first step is
to control the hemorrhage either with digital pressure or with a bandage. Alternatively, you can apply
quick-stop or silver nitrate to the end of the feather for temporary hemostasis. To prevent recurrent
hemorrhage, the blood feather should be completely removed. This can be accomplished by grasping the
base of the fractured feather with fine-tipped hemostats, and provide gentle but firm traction. As the
primary and secondary remiges (flight feathers) are attached to the periosteum, the author recommends to
administer meloxicam (1mg/kg PO BID) for 1-2 days following removal. Complications, although rare,
include incomplete removal of the feather follicle and fracture of the bone if a primary or secondary flight
feather is involved.
DYSPNEA
The initial physical exam of any dyspneic patient is often brief. Therefore, it is important to maximize the
data which can be collected in the first interaction with the patient. Brief observation of the bird in the
carrier is always important to help characterize the dyspnea (see below) and obtain a respiratory rate. The
bird’s mentation and stance (normal or wide-based) should also be recorded. Are the wings held out from
the body or in a normal position? Is there open beak breathing or a tail bob? With experience, the
clinician may then determine the stability of the patient for restraint.
If possible, flow by oxygen supplementation should be delivered during manual restraint for the physical.
The bird should always be maintained in an upright position throughout the physical exam. The key
points to focus on include auscultation (heart, lungs, ventral air sacs), palpation of body condition, and
coelomic palpation. Remember, as birds do not have pulmonary alveoli, they cannot have “crackles” as
you could auscultate a mammal with pulmonary disease. Crackles and other similar sounds may be
auscultated in the air sacs if fluid is present within or around these structures. As with other species,
audible wheezes or other abnormally loud respiratory sounds are often attributed to upper respiratory
disease. A bird with a thin body condition score is more likely to have a chronic condition even if the
owner presented the patient for an acute illness. The presence of coelomic effusion may require more
immediate treatment than just oxygen supplementation including coelomocentesis. If the bird remains
stable, the clinician can proceed with a more thorough physical exam. If not, the bird should be placed in
a warm incubator with oxygen supplementation. It is also important to monitor the bird briefly after
manual restraint to help determine its stability for future restraint—how quickly did the bird recover?
There are several published criteria for divisions of avian dyspnea. 1,9 The author prefers to use a
combination of these criteria in clinical practice. One of the first clinical decisions to make for a dyspneic
bird is whether or not to place an air sac tube. To help determine if this is an appropriate treatment, the
bird’s respiratory pattern can be classified as obstructive or restrictive.
Obstructive respiratory pattern:
• Acute to peracute onset, often in good body condition
• Audible wheezing, +/- loud inspiratory stridor, respiratory rate may be normal
Restrictive respiratory pattern:
• Acute to chronic condition, normal to thin body condition score
• No respiratory noise
• +/- coelomic mass, +/- coelomic effusion
An obstructive breathing pattern is associated with such conditions as a tracheal obstruction or syringeal
mass (i.e. fungal granuloma). In these cases, placement of an air sac tube will help alleviate the dyspnea
and allow the clinician to treat the inciting cause. If any coelomic effusion is present, regardless of the
breathing pattern, air sac tube placement is contraindicated.
Air sac tubes are placed in the caudal thoracic air sac or abdominal air sac using a similar approach as
coelomic endoscopy. General anesthesia is necessary for placement. The clinician should select a tube
that is of similar diameter to the patient’s trachea. The author has used red rubber feeding tubes,
endotracheal tubes, and air sac cannulas for this purpose. The patient is placed in right lateral recumbency,
with the leg pulled cranially. The skin overlying the placement site is routinely prepped, and a small skin
incision is made just ventral to the flexor cruris medialis muscle and caudal to the last rib. A pointed
hemostat is used to create a defect in the coelomic wall with gentle downward pressure using a short finger
stop on the instrument. The tube is secured routinely with a purse string suture followed by a Chinese
finger trap suture. The author also then secures the tube on the bird’s dorsum to prevent self-trauma to
the tube. E-collars may be necessary to protect the integrity of the tube depending on the bird’s
temperament. Whole body radiographs and confirmation of airflow through the tube will help determine
if the air sac tube is in the correct location.
Restrictive breathing patterns are seen for diseases of the small airways, lung parenchyma, and in cases
of coelomic cavity effusion or masses. Additional classification of dyspnea will help localize the disease
process. This will allow the clinician to formulate an appropriate diagnostic and therapeutic plan for each
patient.
Upper airway (nares, infraorbital sinus)
• Nasal discharge, +/- infraorbital swelling or “puffing” with each breath
• Increased rate without increased effort  unless bilateral nasal obstruction is present
Differential diagnosis: URI secondary to hypovitaminosis A and squamous metaplasia, bacterial infections
(gram negative bacteria, Mycoplasma sp, Mycobacterium sp., Chlamydia psittaci), fungal infections (Aspergillus
species, yeast), viral infections (poxvirus, avian influenza), and parasitic infections (Knemidokoptes sp)
Large airway (trachea, syrinx)
• Loud inspiratory stridor
• Increased rate and effort, usually with open beak breathing
Differential diagnosis: tracheal foreign body,3 tracheal stenosis,4 postintubation tracheal obstruction,10
traumatic tracheal collapse,5 tracheal masses,7 syringeal masses (infectious and non-infectious)
Lungs and air sacs
• Soft expiratory wheeze or no audible sounds
• Extreme respiratory distress with open beak breathing
Differential diagnosis: airborne respiratory toxins/irritants (smoke, aerosols, polytetrafluoroethylene
[Teflon]), bacterial infections (gram negative bacteria, Mycobacterium sp., Chlamydia psittaci), fungal infections
(Aspergillus sp, yeast), viral infections (poxvirus, avian influenza, polyomavirus, paramyxoviruses), and
parasitic infections (air sac mites, Syngamus sp)
Coelomic cavity disease
• Increased respiratory rate and effort, exacerbated by handling/stress
• No audible respiratory sounds, +/- distended coelom
Differential diagnosis: transudate (liver failure, heart failure), exudate (septic peritonitis, egg yolk
peritonitis), neoplasia, organomegaly (numerous), egg-binding/dystocia
Oxygen therapy is essential and ideally will be administered to the patient in a quiet, incubator with heat
support. Some authors advocate administration of a bronchial dilator, terbutaline (0.01mg/kg IM), initially
prior to obtaining additional diagnostics.9 Sedation with butorphanol (1-2mg/kg IM) is also indicated as an
initial treatment if the patient is anxious or appears painful. If heart failure is suspected, some authors
recommend administration of furosemide (2-4mg/kg IM or IV).4 Use of this drug in avian species is
controversial as only 10-30% of avian nephrons have a loop of Henle—the remainder are reptilian-type
nephrons that lack this anatomical structure.2
Nebulization of either sterile saline or antibiotics are also beneficial for certain cases of avian dyspnea. A
recent study examined the distribution of fluorescent microspheres after nebulization in domestic pigeons
as a model for nebulization of birds with respiratory disease in clinical practice. 12 Nebulization of 30
minutes to one hour in the pigeons resulted in limited numbers of the microspheres in the secondary
bronchi and pulmonary parenchyma—2-4 hours of nebulization therapy resulted in a more significant
number in the ostia of the lungs and air sac membranes. A similar study also determined that traditional
medical nebulizers designed for use in humans do not create aerosol droplets small enough to reach the
avian lungs and air sacs—these nebulizers create aerosols with an average diameter of 5-10μm, and only
aerosols 3 μm and smaller were noted throughout the pigeon respiratory tract.11
SEIZURES
Just as in other species, it is first important to distinguish a seizure episode from a syncopal episode.
Owners may witness the seizure, or may hear the bird fall off their perch. Common causes of seizures in
birds include:
• Hypocalcemia (secondary to poor diet—seeds and idiopathic in African grey parrots [Psittacus sp])
• Heavy metal toxicity (lead)
• Head trauma
• “Yolk stroke” in egg-laying females (yolk emboli)
• Atherosclerosis – cerebral infarction
Idiopathic epilepsy is uncommon in birds, and is often a diagnosis of exclusion. Initial treatment for status
epilepticus is midazolam 0.5-1mg/kg IM or 2mg/kg intranasal. If hypocalcemia is suspected, 25-50mg/kg
of calcium gluconate can be administered SQ or IM. After stabilization, initial diagnostics should be
performed including whole body radiographs, complete bloodwork, and ideally an ionized calcium if
possible. Even if the bird is not stable for these diagnostics, a single, non-restrained DV (“bird in box”)
radiograph should be taken to look for evidence of metallic foreign bodies, which could lead to seizures
secondary to heavy metal toxicity.
The most important component of treatment is to identify and treat the underlying cause for the seizures.
Supportive care should be provided in the interim which includes fluid therapy, thermal support,
nutritional support, and analgesics if deemed appropriate. Prophylatic antibiotics could also be considered
pending a diagnosis. Antibiotics with good CNS penetration include chloramphenicol, fluoroquinolones,
azithromycin, trimethoprim-sulfa, and third-generation cephalosporins. Currently, there are no effective,
long-term antiepileptic drugs licensed for use in avian medicine. An initial pharmacokinetic study of
phenobarbital in African Grey parrots13 indicated low plasma concentrations when given to birds at
traditional dog/cat doses. Another recent pharmacokinetic study of levetiracetam in Hispaniolan amazon
parrots14 found acceptable plasma levels of 50mg/kg orally every eight hours or 100mg/kg orally every
twelve hours.
FRACTURES
Long bone fractures in birds are often open due to the thin skin and reduced musculature on the distal
extremities. All fractures should be closely examined for puncture wounds through the overlying skin, and
prophylactic antibiotics should be considered for most fractures. This is especially important for fractures
of pneumatic bones (humerus, femur), as pneumonia and air sacculitis can develop secondary to fractures
of these bones. Another initial consideration for any bird with a fracture, especially of the wing, is its
‘occupation’, i.e is it a psittacine with clipped wings or a hunting falcon. Also, remember that bruises in
birds will turn a greenish color 2-3 days post-trauma due to lack of bilirubin reductase and presence of
biliverdin.
Often birds with a fracture will present with a drooped wing, or non-weight bearing on a limb. The
clinician should try to determine where the wing is drooping—at the shoulder or distally. If there is a wing
droop and no apparent fractures, consider a fractured coracoid, proximal humeral fracture (difficult to
palpate), or a brachial plexus injury. As with other species, it is important to check the neurologic status of
the affected wing/leg. The patient should be stabilized with fluids, analgesics (see other presentation on
this topic), +/- sedation. Two view radiographs should be obtained of the affected limb, and the contralateral limb could be used for comparison. Fractures of the radius, ulna, metacarpals, and digits can be
stabilized with a figure-of-8 bandage. Fractures of the humerus, coracoid, and clavicle can be stabilized
with a figure-of-8 bandage with a body wrap. Leg fractures should be stabilized with similar rules as a
mammal, including a joint above and below. Birds that weigh < 200 grams with distal tibiotarsal and
tarsometatarsal fractures can be placed in a tape splint. Consider placement of an E-collar when a wing or
leg bandage is placed, especially in psittacine patients.
THE “FLUFFED” BIRD
The most common ER presentation of the avian patient is a “fluffed bird” or a bird at the bottom of the
cage. Elevation of the feathers, giving a “fluffed” appearance helps to retain body heat and is a general
sign of illness. Normally, birds will perch in the highest location possible; therefore, a bird that stays on
the bottom of the cage is not normal, and this is also a general sign of illness. A complete physical exam
and additional diagnostics are required to determine the etiology.
As with a seizing bird, a single DV view (“bird in box”) should be taken to check for heavy metal toxicity
and the presence of shelled egg(s). Two view whole body radiographs often require sedation for proper
positioning in the psittacine patient. Depending on the case, it may be more appropriate to provide
supportive care for 12-24 hours prior to sedation for diagnostics. A complete blood count can be
accomplished in any hospital by a PCV tube and a blood smear. Abaxis produces an avian and reptile
rotor that measures common biochemistry parameters of these species (VetScan VS2). Fluid therapy is an
important mainstay of generalized supportive care. The daily fluid requirement for most psittacines is 75100mL/kg/day, which can be divided into BID treatments. SQ fluids are most commonly administered as
sedation/anesthesia is often required for placement of IV or IO catheters and fluids given via SQ
administration are rapidly absorbed. If the bird is able to stand, and is not eating, gavage feeding should
be considered to help maintain their nutrient requirements. RER/MER can be calculated, but the clinician
should consider the crop volume of each patient to help determine how much liquid diet could be safely
administered at one time. On average, the crop volume of a typical adult psittacine is 1mL/30 grams of
body weight. If the bird is unable to stand or hold their head up, do not gavage feed them as it can easily
lead to life-threatening regurgitation/aspiration. The clinician may consider prophylactic antibiotics,
depending on the case history and physical exam, prior to obtaining any additional diagnostics.
REGURGITATION
As birds do not possess a diaphragm, technically they cannot vomit. Differentials for regurgitation in birds
are as long and varied the differentials for a vomiting dog. One important differential for the regurgitating
bird is behavioral causes, as this is their natural behavior to feed their young or bond with their mate.
Ideally diagnostics would be performed to help determine the cause of regurgitation including a crop swab
to submit for cytology, gram staining and +/- culture, two-view whole body radiographs, and bloodwork.
As heavy metal toxicity can be a cause of regurgitation, a “bird in the box” radiographic view may be
beneficial in the critical patient. Diagnostics will help determine if there is a localized GI problem or a
underlying systemic disease. Fluoroscopy with GI contrast could also be considered if any abnormalities
in GI motility is suspected. Treatment involves determination of the underlying cause for regurgitation,
and treating that disease. There are no effective anti-emetics in birds, due to the presence of a crop. This
author has used parenteral metoclopramide (0.5mg/kg IM) 10 minutes prior to gavage feeding, but the
efficacy is controversial.
CLOACAL PROLAPSE
Cloacal prolapse may involve the oviduct, ureters, intestines, or copradeum. Prolapses occur secondary to
egg laying, excessive sexual/masterbatory behavior, constipation/tenesmus, or idiopathic/behavioral
(especially in cockatoos). Cloacitis, cloacaliths, severe enteritis, or GI obstruction have all been
documented as causes for cloacal prolapse secondary to tenesmus. Ideally, the primary cause of the cloacal
prolapse should be identified and corrected in addition to replacement of the cloacal tissue (if still viable).
Two lateral vent sutures can be placed to help maintain the cloacal tissue in the proper position, but
recurrent prolapses may require more permanent surgical fixation including ventplasty, cloacopexy, or
salpingectomy (if the oviduct is affected).
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