Urgent Care for Reptiles

Supplemental oxygen is a key part of urgent care for the snake with pneumonia …True or False?

Go to Respiratory Disease in Snakes for quick access to the answer.

Shown here, a corn snake by ‘goingslo’.

Cytology Tips for the Busy Practitioner

When do you perform cytology in your avian patients? When should you?

Read Cytology of the Avian Gastrointestinal Tract by Terry Campbell for practical information and helpful tips.

Macaw chick photo provided by Dr. Greg Rich.

Test Your Knowledge

Did you know that many pet rabbits are “mutts”?

Test your knowledge of varieties commonly seen in practice with the new article: Rabbit Breeds.

Shown here, English lop rabbit.

Responsible Herptile Ownership

Promote responsible ownership of reptiles & amphibians!

Download our new client handout on salmonellosis available in black/white or color versions.

Author: Paul A. Flecknell, VetMB, PhD; Royal College of Veterinary Surgeons, European College of Veterinary Anesthesia and the Comparative Biology Centere, University of Newcastle-upon-Tyne, United Kingdom

Date: April 28, 2010

Key words:  Analgesia, pain, mammal, pre-emptive, multimodal, non-steroidal, NSAID, opioid

Table of Contents

1. Introduction
2. Pre-emptive analgesia
3. Multi-modal analgesia
   1. Opioids
   2. Tramadol
   3. Non-steroidal anti-inflammatory agents
   4. Local anesthetics
4. Practical recommendations for pain prevention
5. Recovery and monitoring
6. Conclusion
7. References

Key Points

• The goal of analgesia is to minimize pain and distress.
• Carefully consider the choice of analgesic drug and timing of administration.
• Prevent “pain windup” by administering opioid analgesia before injury.
• Combinations of analgesics from different classes are more likely to be effective than a single agent used alone.
• Ranges of different analgesics are available, including non-steroidal anti-inflammatory drugs (NSAIDs), opioids, and local anesthetics.
• Gentle handling of tissues can reduce the degree of postoperative pain.
• Pain assessment is more challenging in small mammals. Signs of pain may include immobility, reduced grooming, hunched posture, abdominal press, aggression, and reduced appetite.

Introduction

As in other species, to manage pain successfully, one must know when pain might occur. Several common medical disorders can result in acute pain such as otitis, conjunctivitis, and acute gastrointestinal disease. Chronic pain can arise from conditions such as arthritis, which commonly develops in older rodents, particularly rats (Rattus norvegicus), dental disease [link to article on dental disease in rabbits & rodents], and neoplasia. Management of traumatic and postsurgical pain also requires analgesia. In rabbits and rodents, unrelieved postsurgical pain can be a major factor in the anorexia that is often seen during the postoperative period. Reducing or preventing inappetance is important in small mammals because their energy reserves are significantly lower than in larger species such as dogs and cats. Anorexia can also promote potentially fatal gastrointestinal stasis [inactive link].

Pain management is challenging in small mammals. There are practical difficulties associated with recognizing pain and monitoring the efficacy of pain relief therapy. Nevertheless virtually every available analgesic drug has undergone extensive testing in small mammals. Safe dosages are available for a range of drugs in several common laboratory species (Table 1). The main problem is extrapolating available doses from one species to another and translating doses that are effective in anti-nociceptive tests into those that are appropriate for clinical use. Important concepts in pain prevention include multimodal analgesia and pre-emptive analgesia.

Pre-emptive analgesia

The goal of pre-emptive analgesia is to prevent the noxious stimuli that occur in response to peripheral injury from reaching the central nervous system (CNS). In addition to preventing this “windup” of pain perception, pre-emptive analgesia also aims to reduce or eliminate peripheral inflammation, which increases input into the CNS thereby aggravating central hypersensitivity. Changes in the central processing of noxious stimuli can be suppressed to a greater extent by administration of opioid analgesic before, rather than after injury.

Administration of an opioid as a preanesthetic medication potentiates the effects of the anesthetic agents administered. Volatile anesthetics, such as isoflurane or sevoflurane, are generally preferable because they are simple, relatively safe to administer to most small mammals, and it is easy to adjust the maintenance concentration. If buprenorphine (Buprenex^®, Reckitt & Colman) is administered 30 to 60 minutes before anesthetic induction, the time taken to reach a surgical plane of anesthesia is shorter and recovery is smoother because the animal does not experience postsurgical pain.

Multi-modal analgesia

Multimodal analgesia is based on the premise that because there are numerous pain pathways involving a variety of different neurotransmitters, it is unlikely that a single class of analgesic will be completely effective in controlling pain. Analgesic combinations also allow agents to be used at lower doses, thereby reducing the risk of undesirable side effects.

Multimodal analgesia also has practical advantages. Pre-emptive analgesia is not always appropriate in small, stressed patients and analgesics must sometimes be administered post-induction or postoperatively. Many opioids act rapidly within 10 to 15 minutes of subcutaneous injection, whereas it can be an hour before significant analgesic effects are noted after NSAID administration. Therefore administer a relatively short-acting opioid such as butorphanol (Torbugesic^®, Fort Dodge) or pethidine/meperidine (Demerol^®, Sanofi-Aventis) during recovery to provide immediate pain relief, and concurrently administer an NSAID to provide more prolonged analgesia.

Opioids

Opioids are often required for control of major trauma or postsurgical pain.

Butorphanol and buprenorphine are widely popular agonist/antagonists that have been used to provide analgesia in a variety of species. Buprenorphine has the advantage of having a prolonged duration of action of 6-12 hours in many species including rabbits and rodents. The opioid agonist, pethidine (meperidine) has been used widely in the United Kingdom but has a relatively short duration of action in many species (<2h). The analgesic effects of butorphanol typically last 3-4 hours.

All opioids can produce some degree of respiratory depression, but when administered at clinically effective doses this is rarely a serious problem in small mammals. Opioids may also cause sedation or excitement, but these effects varying considerably with the species. Finally, opioids may cause vomiting, delayed gastric emptying, and increased intestinal peristalsis. These effects may preclude their use in certain experiments but generally these effects are of minimal clinical significance.

Tramadol

Tramadol (Ultram^®, Ortho-McNeil-Janssen) is an opioid agonist that also has analgesic action mediated by inhibition of serotonin and noradrenaline reuptake in the spinal cord. Tramadol has been advocated as an alternative to more potent opioids because of its second mode of action, however it is metabolized relatively rapidly in several species.

Non-steroidal anti-inflammatory drugs

When pain is considered to be mild to moderate, non-steroidal anti-inflammatory drugs (NSAIDs) such as ketoprofen (Ketofen^®, Fort Dodge), carprofen (Rimadyl^®, Pfizer), and meloxicam (Metacam^®, Boehringer Ingelheim) are often sufficient. The palatable oral preparation of meloxicam makes it particularly useful when additional drug doses are required.

The most significant problems associated with NSAID administration are gastrointestinal disturbances, notably ulceration and hemorrhage, nephrotoxicity, and interference with platelet function. Other problems such as blood dyscrasias and liver toxicity can also occur. Adverse effects are rarely significant when these drugs are used to control postsurgical pain in otherwise healthy, young adult animals. Exercise caution when using NSAIDs in animals that may have preexisting organ damage. Chronic renal disease is very common in hamsters (Mesocricetus auratus), rats, and guinea pigs (Cavia porcellus).

Local anesthetics

Local anesthetics can be used to provide post-operative pain relief and as adjuncts to general anesthesia. A longer lasting local agent such as bupivicaine (Marcaine^®, AstraZeneca) may be infiltrated into the wound margins or used to provide a local or regional nerve block. Epidural or spinal administration of drugs has been described in ferrets (Mustela putorious furo), rabbits (Oryctolagus cuniculus), guinea pigs, and small rodents.

Practical recommendations for pain prevention

Surgical pain management

Gentle handling of tissues can reduce the degree of postoperative pain. Stretching and pulling on sensitive structures triggers nociceptive nerve endings. Even though the pet is anesthetized, these nerve impulses trigger changes in the central nervous system (CNS) that result in increased pain during the postoperative period. Rough handling and crushing of tissues also releases substances that intensify the inflammatory response thereby increasing the degree of postsurgical pain.

In my research facility, if the procedure is relatively minor, then only a single dose of analgesic is administered. In some circumstances, a potent NSAID such as meloxicam may be used. For any surgical procedure, an opioid such as buprenorphine is administered either pre-operatively or immediately after anesthetic induction if an inhalant anesthetic is used. Buprenorphine is routinely given in combination with an NSAID for 8-24 hours. This is followed by an NSAID alone for 24-36 hours. Following more invasive procedures, such as laparotomy or orthopedic surgery, opioid administration is continued for up to 48 hours depending upon the species and the expertise of the surgeon. Continue analgesic administration for 72 hours after major surgery.

Management of arthritic pain

Take special care if analgesics are administered long term to manage painful conditions such as arthritis. Fortunately in my experience, oral administration of low doses NSAIDs for relatively long periods can have very positive effects on the quality of life for arthritic rodents. A regimen cycle of 2-3 weeks of treatment, followed by 7-day breaks seems to reduce the risk of undesirable side effects.

Also look for other means of reducing arthritic pain such as providing additional soft bedding. Feed a soft diet rather than a hard pellet to improve food intake, especially if mandibular joint arthritis is present. Also consider the water bottles and change their design or location if they require the animal to adopt a posture that could exacerbate joint pain.

Recovery and monitoring

Pain assessment in small mammals is more difficult than in dogs and cats for a variety of reasons. Changes in behavior provide the most useful indicators of pain; yet veterinarians are often less familiar with the normal behavior of small mammals. Aside from a lack of familiarity with these species, assessment is also difficult because the changes associated with pain are often quite subtle. Additionally many small mammals are prey species, and even caregivers can be viewed as potential predators. In the presence of predators, the response of prey species can be to freeze and remain immobile thereby making an assessment of normal behavior virtually impossible. Even when the animal has been well socialized to human contact, unfamiliar surroundings can be extremely stressful again resulting in a masking of pain-related behavior.

Patients must be provided with a postoperative recovery area appropriate to their particular needs. Allowing a small mammal to recover in the same room as predator species such as cats can be stressful and can also change their behavior patterns, masking signs of pain. Lighting should be subdued but adequate to allow easy observation. Higher intensity lighting must be readily available to allow more detailed examination. The recovery area should be warm and quiet. Provide an ambient temperature should be 27-30ºC (80-86ºF) for adult animals and 35-37ºC (95-98.6ºF) for neonates. Maintain a general room temperature at 21-25ºC (70-77ºF) and provide supplemental heating using warming lamps or heating pads. Although an incubator is ideal, use caution in patients less than 2 kg since many commercially available incubators do not maintain stable temperatures and these patients can suffer from transient hypothermia or hyperthermia. If available, incubators designed for use in human neonates provide excellent conditions for small rodents.

In an effort to assess the patient, carefully observe the animal while undisturbed. Fear and pain can appear similarly. As with other species, many of the changes described here are associated not merely with the presence of pain, but they must be interpreted with the clinical history of the animal.

Both rabbits and small rodents can become completely immobile when experiencing moderate to severe postoperative pain. They will usually position themselves in the farthest corner of their enclosure or try to hide under bedding. When housed in small groups, rodents and rabbits may separate themselves from their cage mates. The animal will usually remain immobile even when approached but will attempt to escape when handled. While moving, the gait may be altered. To complicate matters, nocturnal species such as the rat, hamster, and gerbil are normally relatively inactive during the day. Painful rabbits and rodents may also show a hunched posture.

The painful animal may also groom less frequently. Patients may exhibit piloerection and soiling of the coat. In rats this can lead to build up of reddish-brown secretions or porphyrin tears around the eyes and nose.

Ferrets and rabbits with abdominal pain may loudly grind their teeth, and rats exhibit a characteristic stretching and back-arching behavior following abdominal surgery. Other signs of abdominal pain in rabbits and rodents may include frequent twitching and pressing the ventral abdomen onto the ground.

The painful animal may become unusually aggressive, especially if it is unable to escape from restraint or examination. The animal may also vocalize, sometimes at an abnormal pitch. Remember some of these species are capable of vocalizing at frequencies outside the range of human hearing.

Finally, small mammals often demonstrate reduced appetite if pain is not alleviated effectively. Reductions in food intake are particularly important in rabbits, guinea pigs, and chinchillas because they can trigger gastrointestinal disturbances that can potentially prove fatal. Always weigh animals prior to surgery, so that weight reduction can be detected. [1]

Conclusion

Provided that a few simple principles are observed, providing analgesia in small mammals presents few practical difficulties. The greater challenge is determining the adequacy of the analgesia provided. It is also crucial to keep abreast of changes in analgesia therapy. There are relatively few studies on the efficacy of analgesics in clinically relevant situations. Most information has therefore been obtained from clinical experience. For instance dose rates of analgesics may be extrapolated from other species. Continued review of analgesic use is therefore important because recommendations will change more frequently than they will in companion animal species.

Table 1. ANALGESICS FOR USE IN SMALL LABORATORY ANIMALS

All doses listed are in mg/kg. IM: intramuscular, SC: subcutaneous, IV: intravenous, PO: per os. h: hours

References

Corre TJ, Katz J, Vaccarino AL, Melzack R. Contribution of central neuroplasticity to pathological pain: Review of clinical and experimental evidence. Pain 52():259-285, 1993.

Fairbanks CA. Spinal delivery of analgesics in experimental models of pain and analgesia. Adv Drug Deliv Rev 55(8):1007-1041, 2003.

Flecknell P. Analgesia and post-operative care. In: Flecknell P (ed). Laboratory Animal Anaesthesia, 3^nd ed. London, Academic Press, 2009. Pp. 139-179.

Flecknell PA. Analgesia of small mammals. Vet Clin North Am: Ex Anim Pract 4(1):47-56, 2001.

Flecknell PA, Roughan JV. Assessing pain in animals—putting research into practice. Animal Welfare 13:571-575, 2004.

Hughes PJ, Doherty MM, Charman WN. A rabbit model for the evaluation of epidurally administered local anesthetic agents. Anaesth Intensive Care 21(3):298-303, 1993.

Leach MC, Allweiler S, Richardson C, et al. Behavioural effects of ovariohysterectomy and oral administration of meloxicam in laboratory housed rabbits. Res Vet Sci 87(2):336-347, 2009.

Richmond CE, Bromley LM, Woolf CJ. Preoperative morphine pre-empts postoperative pain. Lancet 342(8863):73-75, 1993.

Souza MJ, Greenacre CB, Cox SK. Pharmacokinetics of orally administered tramadol in domestic rabbits (Oryctolagus cuniculus). Am J Vet Res 69(8):979-982, 2008.

Stokes EL, Flecknell PA, Richardson CA. Reported analgesic and anaesthetic administration to rodents undergoing experimental surgical procedures. Lab An 43():149-154, 2009.

Thomasson B, Ruuskanen O, Merikanto J. Spinal anaesthesia in the guinea pig. Lab Anim 8(2):241-244, 1974.

Turner PV, Chen HC, Taylor WM. Pharmacokinetics of meloxicam in rabbits after single and repeat oral dosing. Comp Med 56(1):63-67, 2006.

Author: Heidi Hoefer, DVM, Dipl. ABVP-Avian; Island Exotic Veterinary Care
Huntington, New York
Date:  February 10, 2010
Key words:  Cardiac, ferret, heart, cardiomyopathy, heartworm, lymphoma, weakness, lethargy, dyspnea

Table of Contents
I.Introduction
II.Clinical picture
III.Differential diagnoses
a.Dilated cardiomyopathy
b.Acquired valvular disease
c.Hypertrophic cardiomyopathy
d.Heartworm disease
e.Arrhythmias
f.Miscellaneous conditions
g.Lymphoma
IV.Diagnosis
V.Therapy
VI.Prognosis
VII.Conclusion
VIII.References
Key Points
Along with insulinoma, cardiac disease is an important differential for weakness and lethargy in the ferret.
Dilated cardiomyopathy is the most common heart disorder in middle aged to older ferrets.
Use caution in the diagnosis of congestive heart failure in ferrets with severe pleural effusion. Pleural fluid may mask the presence of a mediastinal mass caused by lymphoma, particularly in ferrets 2 years of age or younger.
Management of heart disease in the ferret generally follows the same therapeutic guidelines used in dogs and cats, however ferrets seem to be very sensitive to enalapril and can become lethargic secondary to hypotension.
The long-term prognosis for ferrets with cardiac disease is guarded to poor.

Introduction
Cardiac disease is common in middle-aged and older domestic ferrets (Mustela putorius furo). Dilated cardiomyopathy is most common but hypertrophic cardiomyopathy and acquired valvular disease can also be a problem.

Clinical picture
Heart disease is a common finding in ferrets over 4 years of age. Some ferrets are asymptomatic, particularly early in the disease process. Heart murmur or tachyarrhythmia may be incidental findings during physical examination.
Ferrets with advanced heart disease display anorexia and generalized weakness, which may initially present as rear leg weakness or ataxia. Do not mistake weakness or lethargy for insulinoma [link to Pancreatic beta cell tumors in the ferret] before tests are conducted. Common clinical signs of congestive heart failure (CHF) may also include dyspnea or tachypnea, especially if thoracic effusion is present. Pale or cyanotic mucus membranes with prolonged capillary refill time may also be seen. Coughing is rare but may occur secondary to pulmonary edema, heartworm infection, or main-stem bronchi compression. Heart murmurs are common with dilated cardiomyopathy and valvular insufficiency, but relatively rare with hypertrophic cardiomyopathy. Tachyarrhythmias (>250 bpm) may be seen with either form of cardiomyopathy, however a prominent sinus arrhythmia is a common finding in healthy ferrets.

Differential Diagnoses
Dilated cardiomyopathy
Dilated cardiomyopathy (DCM) is the most common cardiac disorder in older ferrets. The heart loses its normal shape, becoming larger and rounder (Fig 1.), and the myocardium becomes progressively weaker until it cannot keep up with bodily demands for oxygen-rich blood. Affected ferrets may show no clinical signs initially but as edema builds up in the lungs, breathing becomes progressively labored. Heart murmurs are common and tachyarrhythmias may also be seen. With proper diagnosis and treatment, affected ferrets may live a year or more from the time of initial diagnosis.

Figure 1.  Dilated heart shape associated with dilated cardiomyopathy in a ferret (Mustela putorius furo). Photograph provided by Dr. Heidi Hoefer.

Acquired valvular disease
Acquired valvular disease caused by endocardiosis is the second most common heart disease seen in ferrets. Heart murmurs are common and may be holosystolic secondary to valvular regurgitation. Left apical murmurs are usually caused by mitral valve disease, and right parasternal murmurs may be caused by tricuspid valve insufficiency.
Hypertrophic cardiomyopathy
Hypertrophic cardiomyopathy (HCM) occurs occasionally in ferrets. While the heart may retain its normal shape, the heart wall becomes progressively thicker until the heart chambers cannot efficiently pump blood. Ferrets with HCM often show no clinical signs until they rapidly de-compensate and enter a crisis. Heart murmurs are relatively uncommon although tachyarrhythmias may be detected. Ferrets with HCM have a poor prognosis with sudden death possible and an expected survival time of only months after diagnosis.
Heartworm disease
Ferrets of any age are susceptible to heartworm disease (HWD) caused by the nematode, Dirofilaria immitis. Although incidence is uncommon, even one or two worms can result in potentially fatal cardiac disease because of the relatively small size of the ferret heart. Clinical signs may include dyspnea, tachypnea, anorexia, pulmonary rales, coughing, holosystolic heart murmur, ascites, pleural effusion, and sometimes sudden death. Early diagnosis is important since clinical disease progresses rapidly.
Radiographs reveal pleural effusion and tortuous, dilated pulmonary vessels. Sonography reveals an enlarged right atrium and ventricle with tricuspid regurgitation. Eosinophilia is rare.
Ferrets with natural infections are not microfilaremic. Use the enzyme-linked immunosorbent assay (IDEXX Snap) antibody test to diagnose HWD. Echocardiography may show nematodes within the right ventricle or vena cava, however this an unreliable diagnostic test. DNA-based polymerase chain reaction assays are capable of sensitive and specific identifcation of D. immitis genetic material in blood specimens, and even a single heartworm can be detected.
Treatment of dirofilariasis in ferrets is possible but carries a guarded prognosis. Successful treatment depends on early diagnosis and anti-thrombotic therapy with prednisone for up to 4 months. The adulticide melarsomine hydrochloride (Immiticide®, Merial) can produce severe reactions including death and should be used with caution. A single dose of moxidectin (ProHeart®, Fort Dodge) 0.1 ml SC per ferret has reportedly been used as an effective and safe adulticide
Small animal products containing ivermectin, selamectin, moxidectin, and milbemycin oxime have all been used in ferrets as heartworm preventive medication in endemic regions. Give ivermectin at 0.02 mg/kg PO or SQ monthly. Dilute the 1% bovine preparation (Ivomec®, Merck AgVET) with propylene glycol to create a 0.1 mg/ml solution, which may be given orally. The suspension should maintain potency until the expiration date of the ivermectin if kept in an amber bottle and protected from light. The 68µg oral ivermectin tablet (Heartgard-30®, Merck AgVET) can be dosed at 1/4 tablet PO monthly, however these tablets cannot be reused once removed from the wrapper and what is left over must be discarded.

Conclusion
Cardiac disease is common in middle-aged and older domestic ferrets. Dilated cardiomyopathy is the most common heart disorder in older ferrets, however hypertrophic cardiomyopathy and valve conditions also occur in the ferret. Clinical signs range from asymptomatic disease to fulminate heart failure with problems such as anorexia, weakness, and dyspnea. In cases where pleural effusion is present and so extensive that the heart cannot be visualized radiographically, be tentative with the initial diagnosis because there may be an imperceptible thoracic mass. Utilize regular veterinary visits to monitor the heart and adjust drug doses as needed. Also monitor ferrets for other common geriatric conditions, such as insulinoma and adrenal gland disease, which can complicate diagnosis and therapy of heart disease.

References
Antinoff N. Clinical observations in ferrets with naturally occurring heartworm disease and preliminary evaluation of treatment with ivermectin with and without melarsomine. Recent Adv Heartworm Dis 2002;45-47.
Fox JG. Other systemic diseases. In Fox JG, ed.: Biology and Diseases of the Ferret, 2nd ed. Philadelphia, Lea & Febiger, 1998, pp 307-320.
Hoefer HL. Thoracic disease in the ferret. Proc Atlantic Coast Veterinary Conference 2001.
Hoefer HL. Heart disease in ferrets. In Bonagura JD, Kirk RW (ed). Current Veterinary Therapy XII. WB Saunders, Philadelphia, 1998. Pp. 1144-1148.
Wagner RA. Ferret cardiology. Vet Clin North Am Exot Anim Pract 12(1):115-134, 2009.

PASTE FROM WORD

Author: Heidi Hoefer, DVM, Dipl. ABVP-Avian; Island Exotic Veterinary Care
Huntington, New York
Date:  February 10, 2010
Key words:  Cardiac, ferret, heart, cardiomyopathy, heartworm, lymphoma, weakness, lethargy, dyspnea,

Table of Contents
I.Introduction
II.Clinical picture
III.Differential diagnoses
a.Dilated cardiomyopathy
b.Acquired valvular disease
c.Hypertrophic cardiomyopathy
d.Heartworm disease
e.Arrhythmias
f.Miscellaneous conditions
g.Lymphoma
IV.Diagnosis
V.Therapy
VI.Prognosis
VII.Conclusion
VIII.References
Key Points
Along with insulinoma, cardiac disease is an important differential for weakness and lethargy in the ferret.
Dilated cardiomyopathy is the most common heart disorder in middle aged to older ferrets.
Use caution in the diagnosis of congestive heart failure in ferrets with severe pleural effusion. Pleural fluid may mask the presence of a mediastinal mass caused by lymphoma, particularly in ferrets 2 years of age or younger.
Management of heart disease in the ferret generally follows the same therapeutic guidelines used in dogs and cats, however ferrets seem to be very sensitive to enalapril and can become lethargic secondary to hypotension.
The long-term prognosis for ferrets with cardiac disease is guarded to poor.

Introduction
Cardiac disease is common in middle-aged and older domestic ferrets (Mustela putorius furo). Dilated cardiomyopathy is most common but hypertrophic cardiomyopathy and acquired valvular disease can also be a problem.

Clinical picture
Heart disease is a common finding in ferrets over 4 years of age. Some ferrets are asymptomatic, particularly early in the disease process. Heart murmur or tachyarrhythmia may be incidental findings during physical examination.
Ferrets with advanced heart disease display anorexia and generalized weakness, which may initially present as rear leg weakness or ataxia. Do not mistake weakness or lethargy for insulinoma [link to Pancreatic beta cell tumors in the ferret] before tests are conducted. Common clinical signs of congestive heart failure (CHF) may also include dyspnea or tachypnea, especially if thoracic effusion is present. Pale or cyanotic mucus membranes with prolonged capillary refill time may also be seen. Coughing is rare but may occur secondary to pulmonary edema, heartworm infection, or main-stem bronchi compression. Heart murmurs are common with dilated cardiomyopathy and valvular insufficiency, but relatively rare with hypertrophic cardiomyopathy. Tachyarrhythmias (>250 bpm) may be seen with either form of cardiomyopathy, however a prominent sinus arrhythmia is a common finding in healthy ferrets.

Differential Diagnoses
Dilated cardiomyopathy
Dilated cardiomyopathy (DCM) is the most common cardiac disorder in older ferrets. The heart loses its normal shape, becoming larger and rounder (Fig 1.), and the myocardium becomes progressively weaker until it cannot keep up with bodily demands for oxygen-rich blood. Affected ferrets may show no clinical signs initially but as edema builds up in the lungs, breathing becomes progressively labored. Heart murmurs are common and tachyarrhythmias may also be seen. With proper diagnosis and treatment, affected ferrets may live a year or more from the time of initial diagnosis.

Figure 1.  Dilated heart shape associated with dilated cardiomyopathy in a ferret (Mustela putorius furo). Photograph provided by Dr. Heidi Hoefer.

Acquired valvular disease
Acquired valvular disease caused by endocardiosis is the second most common heart disease seen in ferrets. Heart murmurs are common and may be holosystolic secondary to valvular regurgitation. Left apical murmurs are usually caused by mitral valve disease, and right parasternal murmurs may be caused by tricuspid valve insufficiency.
Hypertrophic cardiomyopathy
Hypertrophic cardiomyopathy (HCM) occurs occasionally in ferrets. While the heart may retain its normal shape, the heart wall becomes progressively thicker until the heart chambers cannot efficiently pump blood. Ferrets with HCM often show no clinical signs until they rapidly de-compensate and enter a crisis. Heart murmurs are relatively uncommon although tachyarrhythmias may be detected. Ferrets with HCM have a poor prognosis with sudden death possible and an expected survival time of only months after diagnosis.
Heartworm disease
Ferrets of any age are susceptible to heartworm disease (HWD) caused by the nematode, Dirofilaria immitis. Although incidence is uncommon, even one or two worms can result in potentially fatal cardiac disease because of the relatively small size of the ferret heart. Clinical signs may include dyspnea, tachypnea, anorexia, pulmonary rales, coughing, holosystolic heart murmur, ascites, pleural effusion, and sometimes sudden death. Early diagnosis is important since clinical disease progresses rapidly.
Radiographs reveal pleural effusion and tortuous, dilated pulmonary vessels. Sonography reveals an enlarged right atrium and ventricle with tricuspid regurgitation. Eosinophilia is rare.
Ferrets with natural infections are not microfilaremic. Use the enzyme-linked immunosorbent assay (IDEXX Snap) antibody test to diagnose HWD. Echocardiography may show nematodes within the right ventricle or vena cava, however this an unreliable diagnostic test. DNA-based polymerase chain reaction assays are capable of sensitive and specific identifcation of D. immitis genetic material in blood specimens, and even a single heartworm can be detected.

Conclusion
Cardiac disease is common in middle-aged and older domestic ferrets. Dilated cardiomyopathy is the most common heart disorder in older ferrets, however hypertrophic cardiomyopathy and valve conditions also occur in the ferret. Clinical signs range from asymptomatic disease to fulminate heart failure with problems such as anorexia, weakness, and dyspnea. In cases where pleural effusion is present and so extensive that the heart cannot be visualized radiographically, be tentative with the initial diagnosis because there may be an imperceptible thoracic mass. Utilize regular veterinary visits to monitor the heart and adjust drug doses as needed. Also monitor ferrets for other common geriatric conditions, such as insulinoma and adrenal gland disease, which can complicate diagnosis and therapy of heart disease.

References
Antinoff N. Clinical observations in ferrets with naturally occurring heartworm disease and preliminary evaluation of treatment with ivermectin with and without melarsomine. Recent Adv Heartworm Dis 2002;45-47.
Fox JG. Other systemic diseases. In Fox JG, ed.: Biology and Diseases of the Ferret, 2nd ed. Philadelphia, Lea & Febiger, 1998, pp 307-320.
Hoefer HL. Thoracic disease in the ferret. Proc Atlantic Coast Veterinary Conference 2001.
Hoefer HL. Heart disease in ferrets. In Bonagura JD, Kirk RW (ed). Current Veterinary Therapy XII. WB Saunders, Philadelphia, 1998. Pp. 1144-1148.
Wagner RA. Ferret cardiology. Vet Clin North Am Exot Anim Pract 12(1):115-134, 2009.

Introduction

The lungs of birds are small, compact, spongy structures that are fitted closely against the contours of the ribs on either side of the spine. Avian lungs weigh as much as the lungs of mammals of similar body weight, but because avian lungs have much greater tissue density, they occupy about one-half the volume.

The air sac system is a vital part of the avian respiratory system. Air sacs are thin-walled structures, one to two cell layers thick, that extend throughout the body cavity and into the wing and leg bones. Most birds have nine air sacs: two cervical, two cranial thoracic, two caudal thoracic, two abdominal, and a single interclavicular air sac. The abdominal air sacs carry air to leg and pelvic bones; the interclavicular sac branches into the wing bones, sternum, and syrinx.

The air sacs connect directly to the bronchi and make possible an effective continuous flow of air through the lungs. Air flows into the avian lungs through a set of bronchi into the air-capillary system, which is the primary site of oxygen and carbon dioxide exchange. There is a constant flow of air through the avian lung, allowing a more efficient extraction of oxygen than occurs in the mammalian lung. Half of the inhaled air during the first inhalation in this cycle passes through the primary bronchi to the posterior air sacs. During the exhalation of the first breath, the inhaled air moves from the posterior air sacs into the lungs, where it flows through the gas-exchange areas. The next time the bird inhales, this oxygen-depleted air moves into the anterior air sacs. During the second exhalation, the carbon dioxide-rich air is then expelled from the anterior air sacs, bronchi, and trachea back into the atmosphere. The constant airflow through the avian lung allows more efficient extraction of oxygen than occurs in the mammalian lung.

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Indications

Air sac cannulas or tubes are used routinely to ventilate birds by a method other than endotracheal intubation. Air sac cannulas are used for oxygenation and anesthesia, especially during head or trachea surgery where tracheal intubation would be cumbersome. Air sac cannulation is also used therapeutically to aid dyspneic birds. These are often birds with upper respiratory tract foreign bodies, or tracheosyringeal obstruction from granulomas or tumors. Less commonly, air sac cannulas are placed because of tracheal trauma, collapse, or obstruction. In addition, air sac cannulas provide a means to medicate air sacs directly.

Air sac cannulas have also been used in research. Small songbirds are used for studies on neural mechanisms that underlie vocal learning. Researchers have performed survival surgeries for localization of specific brain regions and have used air sac cannulation to deliver anesthetic gas to zebra finches (Taeniopygia guttata) during neurosurgery. Included among the advantages of this method are that it leaves the bird’s head free for stereotaxic targeting and does not interfere with the beak clamps that are often used to position and stabilize the animal’s head. In addition, this method allows for the use of isoflurane, allowing fast, minimally stressful induction, and preventing lengthy postoperative recovery times.

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Complications

Air sac cannulas can be left in place for up to 7 days, however cannulas should ideally be removed within 4 days because of increased risk of bacterial or fungal infection and subsequent air sacculitis associated with prolonged tube placement. Other possible complications of air sac tubes include severe air sac damage, occlusion of the cannula with exudate or fluid, and abdominal organ damage. Life-threatening blood loss can occur during placement of the cannula if an internal organ is penetrated. Subcutaneous emphysema commonly occurs when the cannula is removed, but is self-limiting and will diminish with time.

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Equipment

Minimal equipment is necessary for air sac cannula placement. A surgical blade (size no.11 or no. 15), a fine mosquito hemostat, a needle holder, a pair of scissors, and suture material are all that is required to perform the procedure (Fig 1). A sterile endotracheal tube cut to the appropriate length, trimmed rubber tubing, a commercially available air sac cannula, or plastic tubing from an IV extension set can be used as an air sac cannula.

Figure 1. Minimal equipment is needed for air sac cannula placement. In addition to a tube, select a 15 or 11 blade, fine-tipped mosquito forceps, needle holders, scissors, and appropriately-sized suture material.

The surgical site should be aseptically prepared. Sterile, water-soluble lubricant can be used to keep surrounding feathers from getting into the surgical site. A small, sterile, self-adhesive drape is used to isolate the surgical site. The size of the cannula and suture material will vary depending on the size of the bird.

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Method

In parrots, pigeons and doves, and passerine birds, the aim is to insert the air sac cannula into the caudal thoracic or abdominal air sac (Fig 2). The bird is placed in lateral recumbency and the upper leg is pulled cranially. The entry site is at the junction of the caudal edge of the last rib and the flexor cruris medialis muscle. The pubic bone is an additional landmark that outlines a triangular shaped region for cannula placement (Fig 3).

Figure 2. The air sac cannula is inserted into the caudal thoracic or abdominal air sac space just behind the last rib.

Figure 3. Anatomical landmarks for entry of air sac cannula into the caudal thoracic or abdominal air sac. Place the bird in lateral recumbency with its upper leg pulled cranially (a). The white triangle marks the site for air sac cannula placement at the junction of the caudal end of the last rib (b), the flexor cruris medialis muscle (c), and the pubic bone (d).

The site is prepared by gently plucking feathers and performing a sterile surgical prep of the skin. A sterile drape is placed, a small incision is made through the skin, and the mosquito forceps are used to bluntly dissect through the muscle wall and enter the air sac. The jaws of the mosquito forceps are opened and the cannula is inserted between them into the air sac (Fig 5 and Fig 6).

Figure 4. To insert an air sac cannula, begin by performing a stab incision through the skin, followed by blunt dissection through the muscle with mosquito hemostats and into the air sac lining.

Figure 5. Insert the air sac cannula through the hemostat jaws.

Figure 6. Insert the air sac cannula through the hemostat jaws into the air sac.

If the cannula is placed appropriately, condensation will appear within the cannula tube or on a glass slide placed at the end of the cannula. A down feather can also be held at the end of the cannula to check for gentle air movement as the bird respires (Fig 7). Use caution not to allow the feather to be inhaled into the air sac. To secure the tube in place, a piece of tape can be affixed around the proximal aspect of the cannula, which is then secured to the skin. Alternatively, a finger trap suture technique can be used to secure the tube in place (Fig 8). It is important to re-assess patency after the cannula is sutured in place, as the cannula’s position may shift, resulting in an obstructed airway if the cannula is compressed against an internal structure. Radiographs are often used to assess proper tube placement by determining the length of the tube, its location in the air sac, and whether the tube is affecting other internal organs. Until experience is developed with this procedure, it is advisable to obtain both ventrodorsal and lateral radiographs of the bird once the tube is in place.

Figure 7. One method for checking air sac cannula placement is to hold a down feather at the end of the tube to check for gentle air movement as the bird breathes. Be sure to hold the feather securely to prevent the feather from being inhaled into the air sac.

Figure 8. To secure an air sac cannula in place, use the finger-trap suture technique. Be sure to reassess the patency of the air sac cannula after it is sutured in place.

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Clinical Notes

1. Culture the air sac tube after removal. Cytologic analysis of the tube tip may also be warranted.
2. If an endotracheal tube is used as an air sac cannula, keep the adaptor nearby for emergency oxygen supplementation.
3. Radiographs may be necessary before air sac tube placement to ensure the tube is not placed into a severely diseased air sac. With diffuse airsacculitis, air sac tube placement is not generally recommended.
4. Air sac cannulas can be placed endoscopically to ensure they are placed correctly.

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Harcourt-Brown N, Chitty J (eds). BSAVA Manual of Psittacine Birds. Quedgeley: British Small Animal Veterinary Association; 2005.

Gill FB. Ornithology. New York: WH Freeman and Co.; 2006.

Graham JE. Approach to the dyspneic avian patient. Sem Avian Exotic Pet Med. 13:154–159, 2004.

Nilson PC, Teramitsu I, White SA. Caudal thoracic air sac cannulation in zebra finches for isoflurane anesthesia. J. Neurosci Methods 143(2):107–115, 2005.

Rode JA, Bartholow S, Ludders JW. Ventilation through an air sac cannula during tracheal obstruction in ducks. J. Assoc. Avian Vet 4:98–102, 1990.