Veterinary Procedures

Urine Collection Techniques

Urine can be removed from the bladder by one of four methods: (1) voided (the “free catch”), (2) manual compression of the urinary bladder (expressing the bladder), (3) catheterization, or (4) cystocentesis.


For routine urinalysis, collection of urine by voiding (micturition) is satisfactory. The major disadvantage is risk of contamination of the sample with cells, bacteria, and other debris located in the genital tract and the perineal hair coat. The first portion of the stream is discarded, as it is most likely to contain debris. Voided urine samples are not recommended when bacterial cystitis is suspected.

Manual Compression of the Bladder

Compressing the urinary bladder is occasionally used to collect urine samples from dogs and cats. Critical: Do not use excessive pressure; if moderate digital pressure does not induce micturition, discontinue the technique. Excessive pressure can culminate in forcing contaminated urine (bladder) into the kidneys, or, worse, in patients with a urethral obstruction the urinary bladder can rupture. The technique is most difficult to accomplish in male dogs and male cats.

Urinary Catheterization

Several types of urinary catheters are currently available for use in dogs and cats. The catheter types most often used today are made of rubber, polypropylene, and latex-free silicone. Stainless steel catheters are occasionally used but unless placed with care these can cause damage to the urethra and/or urinary bladder. Generally, urinary catheters serve one of four purposes:

  1. 1. To relieve urinary retention
  2. 2. To test for residual urine
  3. 3. To obtain urine directly from the bladder for diagnostic purposes
  4. 4. To perform bladder lavage and instillation of medication or contrast material

The size of catheters (diameter) usually is calibrated in the French scale; each French unit is equivalent to roughly 0.33 mm. The openings adjacent to the catheter tips are called “eyes.” Human urethral catheters are used routinely in male and female dogs; 4F to 10F catheters are satisfactory for most dogs (Table Recommended Urethral Catheter Sizes for Routine Use in Dogs and Cats). Polypropylene catheters should be individually packaged and sterilized by ethylene oxide gas.

TABLE Recommended Urethral Catheter Sizes for Routine Use in Dogs and Cats

Animal Urethral Catheter Type Size (French Units*)
Cat Flexible vinyl, red rubber, or Tom Cat catheter (polyethylene) 3.5
Male dog (<25 lb) Flexible vinyl, red rubber, or polyethylene 3.5 or 5
Male dog (>25 lb) Flexible vinyl, red rubber, or polyethylene 8
Male dog (>75 lb) Flexible vinyl, red rubber, or polyethylene 10 or 12
Female dog (<10 lb)) Flexible vinyl, red rubber, or polyethylene 5
Female dog (10-50 lb) Flexible vinyl, red rubber, or polyethylene 8
Female dog (>50 lb) Flexible vinyl, red rubber, or polyethylene 10, 12, or 14

*The diameter of urinary catheters is measured on the French (F) scale. One French unit equals roughly 0.33 mm.

Catheterization of the Male Dog

Patient Preparation

Equipment needed to catheterize a male dog includes a sterile catheter (4F to 10F, 18 inches long, with one end adapted to fit a syringe), sterile lubricating jelly, povidone-iodine soap or chlorhexidine, sterile rubber gloves or a sterile hemostat, a 20-mL sterile syringe, and an appropriate receptacle for the collection of urine.

Proper catheterization of the male dog requires two persons. Place the dog in lateral recumbency on either side. Pull the rear leg that is on top forward, and then flex it (). Alternatively, long-legged dogs can be catheterized easily in a standing position.

Before catheter placement, retract the sheath of the penis and cleanse the glans penis with a solution of povidone-iodine 1% or chlorhexidine. Lubricate the distal 2 to 3 cm of the appropriate-size catheter with sterile lubricating jelly. Never entirely remove the catheter from its container while it is being passed because the container enables one to hold the catheter without contaminating it.


The catheter may be passed with sterile gloved hands or by using a sterile hemostat to grasp the catheter and pass it into the urethra. Alternatively, cut a 2-inch “butterfly” section from the end of the thin plastic catheter container. This section can be used as a cover for the sterile catheter, and the clinician can use the cover to grasp and advance the catheter without using gloves.

If the catheter cannot be passed into the bladder, the tip of the catheter may be caught in a mucosal fold of the urethra or there may be a stricture or block in the urethra. In small-breed dogs, the size of the groove in the os penis may limit the size of the catheter that can be passed. One also may experience difficulty in passing the catheter through the urethra where the urethra curves around the ischial arch. Occasionally a catheter of small diameter may kink and bend on being passed into the urethra. When the catheter cannot be passed on the first try, reevaluate the size of the catheter and gently rotate the catheter while passing it a second time. Never force the catheter through the urethral orifice.

Special Considerations

Effective catheterization is indicated by the flow of urine at the end of the catheter, and a sterile 20-mL syringe is used to aspirate the urine from the bladder. Walk the dog immediately after catheterization to encourage urination.

Catheterization of the Female Dog

Patient Preparation

Equipment needed to catheterize a female dog includes flexible urethral catheters identical to those used in the male dog. The following materials also should be on hand: a small nasal speculum, a 20-mL sterile syringe, lidocaine 0.5%, sterile lubricating jelly, a focal source of light, appropriate receptacles for urine collection, and 5 mL of povidone-iodine or a dilute chlorhexidine solution.

Use strict asepsis. Cleanse the vulva with a solution of povidone-iodine or dilute chlorhexidine. Instillation of lidocaine 0.5% into the vaginal vault helps to relieve the discomfort of catheterization. The external urethral orifice is 3 to 5 cm cranial to the ventral commissure of the vulva. In many instances the female dog may be catheterized in the standing position by passing the female catheter into the vaginal vault, despite the fact that the urethral papilla is not visualized directly.


In the spayed female dog, in which blind catheterization may be difficult, the use of a sterilized otoscope speculum andlight source (), vaginal speculum, or anal speculum with a light source will help to visualize the urethral tubercle on the floor of the vagina. In difficult catheterizations it may be helpful to place the animal in dorsal recumbency (). Insertion of a speculum into the vagina almost always permits visualization of the urethral papilla and facilitates passage of the catheter. Take care to avoid attempts to pass the catheter into the fossa of the clitoris because this is a blind, possibly contaminated cul-de-sac.

Catheterization of the Male Cat

Patient Preparation

Before attempting urinary bladder catheterization of the male cat, administer a short-term anesthetic (e.g., ketamine, 25 mg/kg IM), but only after a careful assessment of the cats physical, acid-base, and electrolyte status (see treatment of hyperkalemia).

In some cases, drugs to treat hyperkalemia may be required before anesthetic induction. Once the patient’s electrolyte status has been evaluated and hyperkalemia, if present, addressed appropriately, anesthesia can be induced with a combination of propofol (4 to 7 mg/kg intravenously [IV]) and diazepam (0.1 mg/kg IV); then the patient is intubated and maintained on gas anesthesia.


Place the anesthetized patient in dorsal recumbency. Gently grasp the ventral aspect of the prepuce and move it caudally in such a manner that the penis is extruded. Withdraw the penis from the sheath and gently pull the penis backward. Keeping sterile catheters in a freezer will help them become more rigid to facilitate passage into the urethra. Pass a sterile, flexible plastic or polyethylene (PE 60 to 90) catheter or 3- to 5-inch, 3.5F urethral catheter into the urethral orifice and gently into the bladder, keeping the catheter parallel to the vertebral column of the cat.

Caution: Never force the catheter through the urethra. The presence of debris within the urethral lumen may require the injection of 3 to 5 mL of sterile saline to back-flush urinary “sand” or concretions so that the catheter can be passed. In some instances the presence of cystic and urethral calculi will prevent the passage of a catheter into the urethra. For this reason a lateral radiograph of the penis, with the patients hindlimbs pulled caudally, may help document the presence of a urethral stone.

Catheterization of the Female Cat

Patient Preparation

Urinary bladder catheterization of the female cat is not a simple procedure. When indicated, and after a preanesthetic examination has been performed, attempt the technique only in the anesthetized cat. Urinary bladder catheterization can be accomplished with the use of a rubber or plastic, side-hole (blunt-ended) urinary catheter. The same catheter type used in male cats is effective in female cats. Instilling lidocaine 0.5% has been recommended as a means of decreasing sensitivity to catheter insertion in sedated (not recommended) cats. Cleanse the vulva with an appropriate antiseptic.


Catheterization can be accomplished with the cat in dorsal or ventral recumbency.

Experience and size of the cat dictate which technique works best.

After cleansing of the perineum and vaginal vault, place the patient in sternal recumbency, and gently pass the catheter along the ventral floor of the vaginal vault. Conversely, if the patient is placed in dorsal recumbency, direct the catheter dorsally along the ventral vaginal floor. If a catheter cannot be placed blindly, a small otoscopic speculum can be placed into the vagina, and the catheter pushed into the urethral papilla once it is visualized directly.

Indwelling Urethral Catheter

Patient Preparation

For continuous urine drainage in the awake, ambulatory patient, use a closed collection system to help prevent urinary tract infection. A soft urethral or Foley catheter can be used, and polyvinyl chloride tubing should be connected to the catheter and to the collection bag outside the cage. The collection bag should be below the level of the animal’s urinary bladder. Place an Elizabethan collar on the animal to discourage chewing on the catheter and associated tubing.


The urinary bladder is catheterized as described previously. Despite the quality of care of the catheter, urinary tract infection still may develop in any patient fitted with an indwelling urinary catheter. Ideally, remove the catheter as soon as it is no longer necessary, or if there are clinical signs of a urinary tract infection or previously undiagnosed fever. A urinary catheter is generally changed after it has been in place for more than 48 hours.

Special Considerations

Observe the patient for development of fever, discomfort, pyuria, or other evidence of urinary tract infection. If infection is suspected, remove the catheter and submit urine for culture and sensitivity or determination of minimum inhibitory concentration (MIC). Previously, culture of the catheter tip was recommended to diagnose a catheter-induced infection. However, culture of the catheter tip is no longer recommended, as it may not accurately reflect the type of microorganisms in a urinary tract infection. The empiric use of antibiotics to help prevent catheter-induced infection is not recommended, as their use can allow colonization of resistant nosocomial bacteria in the patient’s urinary tract.


Patient Preparation

Cystocentesis is a common clinical technique used to obtain a sample of urine directly from the urinary bladder of dogs and cats when collecting a voided, or free-catch, aliquot is not preferred. The procedure is indicated when necessary to obtain bladder urine for culture purposes. Urine that is collected by free catch has passed through the urethra and may be contaminated with bacteria, thereby making interpretation of the culture results difficult. Cystocentesis also is performed as a convenience when it is desirable to obtain a small sample of urine but the patient is not ready or cooperative.

Cystocentesis involves insertion of a needle, with a 6- or 12-mL syringe attached, through the abdominal wall and bladder wall to obtain urine samples for urinalysis or bacterial culture. The technique prevents contamination of urine by urethra, genital tract, or skin and reduces the risk of obtaining a contaminated sample. Cystocentesis also may be needed to decompress a severely overdistended bladder temporarily in an animal with urethral obstruction. In these cases, cystocentesis should be performed only if urethral catheterization is impossible. Warning: Penetration of a distended (obstructed) urinary bladder with a needle could result in rupture of the bladder.


To perform cystocentesis, palpate the ventral abdomen just cranial to the junction of the bladder with the urethra, and trap the urinary bladder between the fingers and the palm of the hand. Use one hand to hold the bladder steady within the peritoneal cavity while the other guides the needle. Next, insert the needle through the ventral abdominal wall into the bladder at a 45-degree angle (). Although this procedure is relatively safe, the bladder must have a reasonable volume of urine, and the procedure should not be performed without first identifying and immobilizing the bladder. For the procedure to be performed safely and quickly, the patient must be cooperative. If collection of a urine sample by cystocentesis is absolutely necessary, sedation may be indicated to restrain the patient adequately for the procedure.

Special Considerations

Generally, cystocentesis is a safe procedure, assuming the patient is cooperative and the bladder can be identified and stabilized throughout the procedure. However, injury and adverse reactions can occur. In addition to laceration of the bladder with the inserted needle (patient moves abruptly), the needle can be passed completely through the bladder and into the colon, causing bacterial contamination of the bladder or peritoneal cavity. There is also risk of penetrating a major abdominal bloodvessel, resulting in significant hemorrhage.


Amantadine HCL (Symmetrel)

Antiviral (Influenza A); Nmda Antagonist

Highlights Of Prescribing Information

Antiviral drug with NMDA antagonist properties; may be useful in adjunctive therapy of chronic pain in small animals & treatment of equine influenza in horses

Very limited clinical experience; dogs may exhibit agitation & GI effects, especially early in therapy

Large interpatient variations of pharmacokinetics in horses limit its therapeutic usefulness

Overdoses are potentially very serious; fairly narrow therapeutic index in dogs & cats; may need to be compounded

Extra-label use prohibited (by FDA) in chickens, turkeys & ducks

What Is Amantadine HCL Used For?

While amantadine may have efficacy and clinical usefulness against some veterinary viral diseases, presently the greatest interest for its use in small animals is as a NMDA antagonist in the adjunctive treatment of chronic pain, particularly those tolerant to opioids.

Amantadine has also been investigated for treatment of equine-2 influenza virus in the horse. However, because of expense, interpatient variability in oral absorption and other pharmacokinetic parameters, and the potential for causing seizures after intravenous dosing, it is not commonly used for treatment.

In humans, amantadine is used for treatment and prophylaxis of influenza A, parkinsonian syndrome, and drug-induced extrapyramidal effects. As in veterinary medicine, amantadine’s effect on NMDA receptors in humans are of active interest, particularly its use as a co-analgesic with opiates and in the reduction of opiate tolerance development.


Like ketamine, dextromethorphan and memantine, amantadine antagonizes the N-methyl-D-aspartate (NMDA) receptor. Within the central nervous system, chronic pain can be maintained or exacerbated when glutamate or aspartate bind to this receptor. It is believed that this receptor is particularly important in allodynia (sensation of pain resulting from a normally non-noxious stimulus). Amantadine alone is not a particularly good analgesic, but in combination with other analgesics (e.g., opiates, NSAIDs), it is thought that it may help alleviate chronic pain.

Amantadine’s antiviral activity is primarily limited to strains of influenza A. While its complete mechanism of action is unknown, it does inhibit viral replication by interfering with influenza A virus M2 protein.

Amantadine’s antiparkinsonian activity is not well understood. The drug does appear to have potentiating effects on dopaminergic neurotransmission in the CNS and anticholinergic activity.


The pharmacokinetics of this drug have apparently not been described in dogs or cats. In horses, amantadine has a very wide interpatient variability of absorption after oral dosing; bioavailability ranges from 40-60%. The elimination half-life in horses is about 3.5 hours and the steady state volume of distribution is approximately 5 L/kg.

In humans, the drug is well absorbed after oral administration with peak plasma concentrations occurring about 3 hours after dosing. Volume of distribution is 3-8 L/kg. Amantadine is primarily eliminated via renal mechanisms. Oral clearance is approximately 0.28 L/hr/kg; half-life is around 17 hours.

Before you take Amantadine HCL

Contraindications / Precautions / Warnings

In humans, amantadine is contraindicated in patients with known hypersensitivity to it or rimantadine, and in patients with untreated angle-closure glaucoma. It should be used with caution in patients with liver disease, renal disease (dosage adjustment may be required), congestive heart failure, active psychoses, eczematoid dermatitis or seizure disorders. In veterinary patients with similar conditions, it is advised to use the drug with caution until more information on its safety becomes available.

In 2006, the FDA banned the use of amantadine and other influenza antivirals in chickens, turkeys and ducks.

Adverse Effects

There is very limited experience in domestic animals with amantadine and its adverse effect profile is not well described. It has been reported that dogs given amantadine occasionally develop agitation, loose stools, flatulence or diarrhea, particularly early in therapy. Experience in cats is limited; an adverse effect profile has yet to be fully elucidated, but the safety margin appears to be narrow.

Reproductive / Nursing Safety

In humans, the FDA categorizes amantadine as a category C drug for use during pregnancy (Animal studies have shown an adverse effect on the fetus, hut there are no adequate studies in humans; or there are no animal reproduction studies and no adequate studies in humans). High dosages in rats demonstrated some teratogenic effects.

Amantadine does enter maternal milk. The manufacturer does not recommend its use in women who are nursing. Veterinary significance is unclear.

Overdosage / Acute Toxicity

Toxic dose reported for cats is 30 mg/kg and behavioral effects may be noted at 15 mg/kg in dogs and cats.

In humans, overdoses as low as 2 grams have been associated with fatalities. Cardiac dysfunction (arrhythmias, hypertension, tachycardia), pulmonary edema, CNS toxicity (tremors, seizures, psychosis, agitation, coma), hyperthermia, renal dysfunction and respiratory distress syndrome have all been documented. There is no known specific antidote for amantadine overdose. Treatment should consist of gut emptying, if possible, intensive monitoring and supportive therapy. Forced urine acidifying diuresis may increase renal excretion of amantadine. Physostigmine has been suggested for cautious use in treating CNS effects.

How to use Amantadine HCL

Amantadine HCL dosage for dogs:

As adjunctive therapy for chronic pain:

a) 1.25-4 mg/kg PO ql2-24h. Usually use 3 mg/kg PO once daily as an adjunct with a NSAID May require 5-7 days to have a positive effect. ()

b) Approximate dose is 3-5 mg/kg PO once daily. ()

c) To decrease wind-up: 3-5 mg/kg PO once daily for one week ()

Amantadine HCL dosage for cats:

As adjunctive therapy for chronic pain:

a) 3 mg/kg PO once daily. May be useful addition to NSAIDs; not been evaluated for toxicity. May need to be compounded. ()

b) Approximate dose is 3-5 mg/kg PO once daily. ()

c) 3 mg/kg PO once daily. ()

Amantadine HCL dosage for horses:

For acute treatment of equine-2 influenza: a) 5 mg/kg IV q4h ()


■ Adverse effects (GI, agitation)

■ Efficacy

Client Information

■ When used in small animals, the drug must be given as prescribed to be effective and may take a week or so to show effect.

■ Gastrointestinal effects (loose stools, gas, diarrhea) or some agitation may occur, particularly early in treatment. Contact the veterinarian if these become serious or persist.

■ Overdoses with this medication can be serious; keep well out of reach of children and pets.

Chemistry / Synonyms

An adamantane-class antiviral agent with NMDA antagonist properties, amantadine HCL occurs as a white to practically white, bitter tasting, crystalline powder with a pKa of 9. Approximately 400 mg are soluble in 1 mL of water; 200 mg are soluble in 1 mL of alcohol.

Amantadine HCL may also be known as: adamantanamine HCL, Adekin, Amanta, Amantagamma, Amantan, Amantrel, Amixx, Antadine, Antiflu-DES, Atarin, Atenegine, Cerehramed, Endantadine, Infectoflu, Influ-A, Lysovir, Mantadine, Mantadix, Mantidan, Padiken, Symadine, Symmetrel, Viroifral and Virucid.

Storage / Stability

Tablets, capsules and the oral solution should be stored in tight containers at room temperature. Limited exposures to temperatures as low as 15°C and as high as 30°C are permitted. Avoid freezing the liquid.

Dosage Forms / Regulatory Status

Veterinary-Labeled Products: None

Human-Labeled Products:

Amantadine HCL Tablets & Capsules: 100 mg; Symmetrel (Endo); generic; (Rx)

Amantadine HCL Syrup: 10 mg/mL in 480 mL; Symmetrel (Endo); generic; (Rx)

In 2006, the FDA banned the extra-label use of amantadine and other influenza antivirals in chickens, turkeys and ducks.


HCM: Pathophysiology

Hypertrophy, Diastolic Dysfunction, and Congestive Heart Failure

Enlarged papillary muscles and a thick left ventricular myocardium with a normal to small left ventricular chamber characterize hypertrophic cardiomyopathy. hypertrophic cardiomyopathy may be mild, moderate, or severe. Severe concentric hypertrophy by itself increases chamber stiffness. In addition, blood flow and especially blood flow reserve to severely thickened myocardium is compromised, which causes myocardial ischemia, cell death, and replacement fibrosis. This has been documented in cats by showing that cardiac troponin I concentration, a protein released into the systemic circulation after cell necrosis, is increased in cats with severe hypertrophic cardiomyopathy. Increased concentrations of circulating neurohormones may also stimulate interstitial fibrosis. Fibrosis increases chamber stiffness (increased pressure for any given volume) further and is probably the primary reason for the marked diastolic dysfunction seen in this disease. The stiff ventricular chamber causes a greater increase in pressure for any given increase in volume when the ventricle fills in diastole. This produces congestive heart failure (i.e., pulmonary edema and pleural effusion).The myocardium from cats with hypertrophic cardiomyopathy also takes a longer time to relax in early diastole, although the clinical significance of this is unknown. When left ventricular hypertrophy is severe, it is common for the left ventricular wall thickness to be twice the normal thickness. Consequently, left ventricular wall thickness is commonly in the 7 to 10 mm range when hypertrophic cardiomyopathy is severe in cats. This severe concentric hypertrophy may encroach on the left ventricular cavity in diastole, decreasing its size, although left ventricular diastolic diameter is commonly within the normal range. The end-systolic volume and diameter are almost always reduced, often to zero (endsystolic cavity obliteration). Global myocardial contractility is normal in humans with hypertrophic cardiomyopathy, and die reduction in end-systolic volume is due to a decrease in afterload (wall stress) brought on by the increase in wall thickness.

Systolic Anterior Motion of the Mitral Valve A phenomenon called systolic anterior motion (SAM) of the mitral valve is common in cats with hypertrophic cardiomyopathy ().

Cats with hypertrophic cardiomyopathy and systolic anterior motion are commonly said to have the obstructive type of hypertrophic cardiomyopathy or hypertrophic obstructive cardiomyopathy (HOCM). In one survey of 46 cats, systolic anterior motion was present in 67%. systolic anterior motion of the mitral valve is the process of the septal (anterior] mitral valve leaflet or the chordal structures inserting on this leaflet being pulled into the left ventricular outflow tract during systole. Here it is caught in the blood flow and pushed toward (and often ultimately against) the IVS. The initial pulling of the mitral valve leaflet toward the left ventricular outflow tract in systole can clearly be seen on many echocardiograms from cats with hypertrophic cardiomyopathy. The grossly enlarged papillary muscles encroach on the left ventricular outflow tract (the region of the left ventricular between the anterior leaflet of the mitral valve and the IVS in diastole) and pull the mitral apparatus structures into the basilar region of the outflow tract. This situation has been reproduced experimentally in dogs by surgically displacing the papillary muscles cranially. systolic anterior motion of the mitral valve produces a dynamic subaor-tic stenosis that increases systolic intraventricular pressure in mid- to late systole. The dynamic subaortic stenosis increases the velocity of blood flow through the subaortic region and often produces turbulence. Simultaneously, when the septal leaflet is pulled toward the IVS, this produces a gap in the mitral valve, creating mitral regurgitation. These abnormalities are by far the most common cause of the heart murmur heard in cats with hypertrophic cardiomyopathy. The process of systolic anterior motion is dynamic — worsening when contractility increases and lessening when contractility decreases. This also makes the murmur dynamic — increasing in intensity with increasing excitement and softening when the cat becomes calmer.

Pleural Effusion Along with pulmonary edema, pleural effusion is common in cats with heart failure. It can be a modified transudate, pseudochylous or true chylous in nature. The most common cause of chylothorax in cats is heart failure. It is unknown exactly why pleural effusion develops in these cats. Two possibilities exist: The first is that left heart failure results in pulmonary hypertension severe enough to cause right heart failure. This does not appear to occur very frequently in cats because it is unusual to identify echocardiographic right heart enlargement, jugular and hepatic vein distension, or ascites in these cases. The second possibility is that feline visceral pleural veins drain into the pulmonary veins such that elevated pulmonary vein pressure (congestive left heart failure) causes the formation of pleural effusion. In the dog the visceral pleura is supplied by pulmonary arteries and drained by pulmonary veins. The dog, the cat, and the monkey have type II lungs.w One characteristic of type II lungs is that the visceral pleura is supplied not by bronchial arteries but by pulmonary arteries.

Presumably this means that the cat visceral pleura is also drained by pulmonary veins. This means that pulmonary venous hypertension secondary to left heart failure could cause pleural effusion in cats as it does in humans.

Pathology of hypertrophic cardiomyopathy

Gross Pathology Cats with severe hypertrophic cardiomyopathy have severe thickening of the left ventricular myocardium (the IVS and free wall), with the left ventricular wall commonly being 7 to 10 mm thick (). The hypertrophy may be symmetrical, involving the entire circumference of the LV, but it may also be asymmetrical. In some cats the IVS is significandy thicker than the free wall, whereas in others the free wall is thicker (asymmetrical hypertrophy). In those cats with primarily septa] hypertrophy, the hypertrophy may be confined to the basilar region of the septum, and in others may be apical. Isolated free wall hypertrophy most commonly occurs in the region between the papillary muscles. As in many pathologic specimens, hearts from cats with hypertrophic cardiomyopathy may undergo contraction (rigor) after death, resulting in a wall thickness that is closer to the end-systolic wall thickness in life rather than the end-diastolic thickness. Consequently, heart weight must be combined with subjective or objective evidence of left ventricular wall thickening to make the diagnosis of hypertrophic cardiomyopathy postmortem. To weigh a cat heart the pericardium should be removed and the aorta and pulmonary artery transected so that no more than 2 to 4 cm are left. Normal heart weight-to-body weight ratio has been reported to be 10.6 +/- 4 g/lb, with cats with hypertrophic cardiomyopathy having a ratio of 13.2 +/-3.1 g/1b. This results in a large overlap between the two groups. In the author’s experience, most normal-sized cats (6 to 12 lb) have a heart that weighs less than 20 g and most cats in this size range with hypertrophic cardiomyopathy have a heart that weighs more than 20 g. Cats with severe hypertrophic cardiomyopathy almost always have a heart that weighs more than 25 g, usually over 30 g, and can be as heavy as 38 g.

The left atrium is often enlarged in cats with severe hypertrophic cardiomyopathy, often markedly so. However, with early, severe disease, the author has identified normal left atrial size in some Maine coon cats. Occasionally a thrombus is present in the body of the left atrium or within the left auricle.

Cats with milder forms of the disease (mild to moderate hypertrophic cardiomyopathy) have lesser wall thickening and a more normal-sized left ventricular chamber. The left atrium may be normal in size or may be enlarged. Papillary muscle hypertrophy may be the predominant lesion.

Histopathology Histopathologically, a wide range of abnormalities exist. In some hearts, only myocyte hypertrophy is evident. On the other end of the spectrum, some cats have moderate to severe interstitial and replacement fibrosis and dystrophic mineralization (20% to 40% of cases). Intramural coronary arteriosclerosis is present in approximately 75% of cats with hypertrophic cardiomyopathy. Intramyocardial small artery disease is not specific for hypertrophic cardiomyopathy because it is also identified in cats and dogs with many cardiac diseases.

In humans, myocardial fiber disarray that involves at least 5% of the myocardium in the IVS is found in 90% of patients with familial hypertrophic cardiomyopathy. Other diseases that produce concentric hypertrophy can also cause myocardial fiber disarray, but this almost always involves less than 1% of the myocardium. In cats with hypertrophic cardiomyopathy, myocardial fiber disarray in the IVS of the same magnitude observed in humans is only identified in 30% to 60% of cases.’ However, myocardial fiber disarray is a consistent feature of hypertrophic cardiomyopathy in Maine coon cats. Sarcomeres also have disarray in human patients with hypertrophic cardiomyopathy. Interestingly, infecting isolated feline cardiocytes with an adenoviral vector containing a full-length mutated human β-myosin heavy chain gene causes sarcomere disruption.

Natural History and Prognosis

The prognosis, as with most cardiac diseases, is highly variable for hypertrophic cardiomyopathy. Some of it is determined by clinical presentation and echocardiographic severity of the disease. Adult cats that are asymptomatic and have mild to moderate disease and no to mild left atrial enlargement have a good short-term (and possibly a good long-term) prognosis. Some, however, may progress to more severe disease and some may die suddenly. Asymptomatic cats with severe wall thickening and mild to moderate left atrial enlargement have a guarded prognosis for developing heart failure in the future. They probably have some risk for developing thromboembolism and may be at risk for sudden death. Cats with no clinical signs but with severe wall thickening and moderate to severe left atrial enlargement are at risk for developing heart failure or often already have mild to moderate heart failure that has gone undetected. These cats are at risk for developing systemic thromboembolic disease and sudden death, although both of these risks appear to be relatively small. Cats presented in heart failure usually have a poor prognosis, but survival time is highly variable. Most die of intractable heart failure. Some develop thromboembolism, and some die suddenly. In one study a MST of 3 months was reported yet some cats (about 20% in one study) in this class stabilize and do well for prolonged periods. The author and colleagues have seen some cats with severe hypertrophic cardiomyopathy live as long as 2.5 years after the diagnosis of heart failure. Some of these cats may develop heart failure when they are stressed and become severely tachycardic and then stabilize after that time. Cats with severe hypertrophic cardiomyopathy and aortic thromboembolism in the aforementioned study had a very poor prognosis with a MST of 2 months.

Hypertrophic Cardiomyopathy: Clinical Manifestations

Differential Diagnoses

Hyperthyroidism and systemic hypertension need to be ruled out as either primary or complicating factors. Hyperthyroidism is usually easy to rule out. Devices to measure blood pressure in the cat, however, are not always readily available, and the technique requires some practice to acquire accurate values. In addition, systolic systemic arterial blood pressure may be increased in a normal cat that is stressed, so repeat measurements of increased blood pressure are preferred before a diagnosis of systemic hypertension is made If systemic arterial blood pressure cannot be measured, one should at least rule out the common causes of systemic hypertension in a cat with left ventricular concentric hypertrophy (i.e., hyperthyroidism, renal failure).

Rarely, infiltrative disease such as lymphoma will produce hypertrophy that is indistinguishable from hypertrophic cardiomyopathy on an echocar-diogram. One such case has been reported. Cats that are homozygous for the dystrophin deficiency seen in hypertrophic feline muscular dystrophy also have thickened but hypoechoic myocardium with hyperechoic foci in the left ventricular myocardium and papillary muscles. The myocardium contains foci of mineralization and no dystrophin.

Precipitating Factors

Certain factors may precipitate heart failure or sudden death in a cat with hypertrophic cardiomyopathy. Stress (cat fight), anesthesia (especially with ketamine), and surgery appear to be factors. The administration of a long-acting corticosteroid also appears to be a factor that either produces or worsens heart failure in cats, presumably through the mineralocorticoid effects of these drugs.

Therapy of Cats with No Clinical Signs

No evidence exists to show that any drug alters the natural history of hypertrophic cardiomyopathy in domestic cats until they are in heart failure. Diltiazem, atenolol, or enalapril are commonly administered to cats with mild to severe hypertrophic cardiomyopathy that are not in heart failure on an empiric basis. Whenever hypertrophic cardiomyopathy is diagnosed in a cat, the veterinarian should explain the situation to owners and try to let them make informed decisions based on their wishes and life styles. Because no intervention is known to change the course of the disease, treatment at this stage is not mandated.

Treatment Goals and General Therapy of Cats in Heart Failure

Cats that present in heart failure have clinical signs referable to pulmonary edema, pleural effusion, or both. Consequendy, therapy is generally aimed at decreasing left atrial and pulmonary venous pressures in these cats and physically removing the effusion. In some cats with severe heart failure, clinical evidence of hypoperfusion (low-output heart failure) may be apparent in addition to the signs of congestive heart failure. The signs may be manifested primarily as cold extremities.

Pulmonary edema is primarily treated with diuretics (almost exclusively with furosemide) acutely and chronically and an angiotensin-converting enzyme enzyme inhibitor chronically, although recent evidence suggests that angiotensin-converting enzyme inhibition may not be that helpful in prolonging survival in cats with hypertrophic cardiomyopathy. Diltiazem and beta-adrenergic blockers, usually atenolol, have been commonly used as adjunctive agents. Recent evidence suggests that diltiazem is not helpful in prolonging survival in cats with heart failure due to severe hypertrophic cardiomyopathy and that atenolol may actually shorten survival time. Plcurocentesis is most effective for treating cats with severe pleural effusion. Furosemide is helpful for preventing or slowing recurrent effusion.

Hypertrophic Cardiomyopathy: Acute Therapy

Hypertrophic Cardiomyopathy: Chronic Therapy

Refractory Heart Failure

Heart failure that is refractory to furosemide and an angiotensin-converting enzyme inhibitor portends a poor prognosis. Another diuretic may be added to the therapeutic regimen. A thiazide diuretic is generally the mast rewarding but is also more likely to cause complications, such as dehydration and electrolyte (sodium, potassium, chloride, magnesium) depletion. Spironolactone, in theory, may have some beneficial effects related to blocking aldos-terone’s actions; however, clinically it rarely results in noticeable improvement, and its efficacy is unproven. A low sodium diet may be helpful, if palatable. This diet can be a commercial one or one that is devised by a nutritional service. Home-cooked diets formulated by the owner are discouraged unless the owner is counseled. If severe systolic anterior motion is present and atenolol is not already part of the therapeutic regimen, it may be added at this stage.


Acepromazine Maleate (PromAce, Aceproject)

Phenothiazine Sedative / Tranquilizer

Highlights Of Prescribing Information

Negligible analgesic effects

Dosage may need to be reduced in debilitated or geriatric animals, those with hepatic or cardiac disease, or when combined with other agents

Inject IV slowly; do not inject into arteries

Certain dog breeds (e.g., giant breeds, sight hounds) may be overly sensitive to effects

May cause significant hypotension, cardiac rate abnormalities, hypo- or hyperthermia

May cause penis protrusion in large animals (esp. horses)

What Is Acepromazine Used For?

Acepromazine is approved for use in dogs, cats, and horses. Labeled indications for dogs and cats include: “… as an aid in controlling intractable animals… alleviate itching as a result of skin irritation; as an antiemetic to control vomiting associated with motion sickness” and as a preanesthetic agent. The use of acepromazine as a sedative/tranquilizer in the treatment of adverse behaviors in dogs or cats has largely been supplanted by newer, effective agents that have fewer adverse effects. Its use for sedation during travel is controversial and many no longer recommend drug therapy for this purpose.

In horses, acepromazine is labeled “… as an aid in controlling fractious animals,” and in conjunction with local anesthesia for various procedures and treatments. It is also commonly used in horses as a pre-anesthetic agent at very small doses to help control behavior.

Although not approved, it is used as a tranquilizer (see doses) in other species such as swine, cattle, rabbits, sheep and goats. Acepromazine has also been shown to reduce the incidence of halothane-induced malignant hyperthermia in susceptible pigs.

Before you take Acepromazine

Contraindications / Precautions / Warnings

Animals may require lower dosages of general anesthetics following acepromazine. Use cautiously and in smaller doses in animals with hepatic dysfunction, cardiac disease, or general debilitation. Because of its hypotensive effects, acepromazine is relatively contraindicated in patients with hypovolemia or shock. Phenothiazines are relatively contraindicated in patients with tetanus or strychnine intoxication due to effects on the extrapyramidal system.

Intravenous injections should be made slowly. Do not administer intraarterially in horses since it may cause severe CNS excitement/depression, seizures and death. Because of its effects on thermoregulation, use cautiously in very young or debilitated animals.

Acepromazine has no analgesic effects; treat animals with appropriate analgesics to control pain. The tranquilization effects of acepromazine can be overridden and it cannot always be counted upon when used as a restraining agent. Do not administer to racing animals within 4 days of a race.

In dogs, acepromazine’s effects may be individually variable and breed dependent. Dogs with MDR1 mutations (many Collies, Australian shepherds, etc.) may develop a more pronounced sedation that persists longer than normal. It may be prudent to reduce initial doses by 25% to determine the reaction of a patient identified or suspected of having this mutation.

Acepromazine should be used very cautiously as a restraining agent in aggressive dogs as it may make the animal more prone to startle and react to noises or other sensory inputs. In geriatric patients, very low doses have been associated with prolonged effects of the drug. Giant breeds and greyhounds may be extremely sensitive to the drug while terrier breeds are somewhat resistant to its effects. Atropine may be used with acepromazine to help negate its bradycardic effects.

In addition to the legal aspects (not approved) of using acepromazine in cattle, the drug may cause regurgitation of ruminal contents when inducing general anesthesia.

Adverse Effects

Acepromazine’s effect on blood pressure (hypotension) is well described and an important consideration in therapy. This effect is thought to be mediated by both central mechanisms and through the alpha-adrenergic actions of the drug. Cardiovascular collapse (secondary to bradycardia and hypotension) has been described in all major species. Dogs may be more sensitive to these effects than other animals.

In male large animals acepromazine may cause protrusion of the penis; in horses, this effect may last 2 hours. Stallions should be given acepromazine with caution as injury to the penis can occur with resultant swelling and permanent paralysis of the penis retractor muscle. Other clinical signs that have been reported in horses include excitement, restlessness, sweating, trembling, tachypnea, tachycardia and, rarely, seizures and recumbency.

Its effects of causing penis extension in horses, and prolapse of the membrana nictitans in horses and dogs, may make its use unsuitable for show animals. There are also ethical considerations regarding the use of tranquilizers prior to showing an animal or having the animal examined before sale.

Occasionally an animal may develop the contradictory clinical signs of aggressiveness and generalized CNS stimulation after receiving acepromazine. IM injections may cause transient pain at the injection site.

Overdosage / Acute Toxicity

The LD50 in mice is 61 mg/kg after IV dosage and 257 mg/kg after oral dose. Dogs receiving 20-40 mg/kg over 6 weeks apparently demonstrated no adverse effects. Dogs gradually receiving up to 220 mg/kg orally exhibited signs of pulmonary edema and hyperemia of internal organs, but no fatalities were noted.

There were 128 exposures to acepromazine maleate reported to the ASPCA Animal Poison Control Center (APCC; during 2005-2006. In these cases, 89 were dogs with 37 showing clinical signs and the remaining 39 reported cases were cats with 12 cats showing clinical signs. Common findings in dogs recorded in decreasing frequency included ataxia, lethargy, sedation, depression, and recumbency. Common findings in cats recorded in decreasing frequency included lethargy, hypothermia, ataxia, protrusion of the third eyelid, and anorexia.

Because of the apparent relatively low toxicity of acepromazine, most overdoses can be handled by monitoring the animal and treating clinical signs as they occur; massive oral overdoses should definitely be treated by emptying the gut if possible. Hypotension should not be treated with epinephrine; use either phenylephrine or norepinephrine (levarterenol). Seizures may be controlled with barbiturates or diazepam. Doxapram has been suggested as an antagonist to the CNS depressant effects of acepromazine.

How to use Acepromazine

Note: The manufacturer’s dose of 0.5-2.2 mg/kg for dogs and cats is considered by many clinicians to be 10 times greater than is necessary for most indications. Give IV doses slowly; allow at least 15 minutes for onset of action.

Acepromazine dosage for dogs:

a) Premedication: 0.03-0.05 mg/kg IM or 1-3 mg/kg PO at least one hour prior to surgery (not as reliable) ()

b) Restraint/sedation: 0.025-0.2 mg/kg IV; maximum of 3 mg or 0.1-0.25 mg/kg IM; Preanesthetic: 0.1-0.2 mg/kg IV or IM; maximum of 3 mg; 0.05-1 mg/kg IV, IM or SC ()

c) To reduce anxiety in the painful patient (not a substitute for analgesia): 0.05 mg/kg IM, IV or SC; do not exceed 1 mg total dose ()

d) 0.55-2.2 mg/kg PO or 0.55-1.1 mg/kg IV, IM or SC (Package Insert; PromAce — Fort Dodge)

e) As a premedicant with morphine: acepromazine 0.05 mg/kg IM; morphine 0.5 mg/kg IM ()

Acepromazine dosage for cats:

a) Restraint/sedation: 0.05-0.1 mg/kg IV, maximum of 1 mg ()

b) To reduce anxiety in the painful patient (not a substitute for analgesia): 0.05 mg/kg IM, IV or SC; do not exceed 1 mg total dose ()

c) 1.1-2.2 mg/kg PO, IV, IM or SC (Package Insert; PromAce — Fort Dodge)

d) 0.11 mg/kg with atropine (0.045-0.067 mg/kg) 15-20 minutes prior to ketamine (22 mg/kg IM). ()

Acepromazine dosage for ferrets:

a) As a tranquilizer: 0.25-0.75 mg/kg IM or SC; has been used safely in pregnant jills, use with caution in dehydrated animals. ()

b) 0.1-0.25 mg/kg IM or SC; may cause hypotension/hypothermia ()

Acepromazine dosage for rabbits, rodents, and small mammals:

a) Rabbits: As a tranquilizer: 1 mg/kg IM, effect should begin in 10 minutes and last for 1-2 hours ()

b) Rabbits: As a premed: 0.1-0.5 mg/kg SC; 0.25-2 mg/kg IV, IM, SC 15 minutes prior to induction. No analgesia; may cause hypotension/hypothermia. ()

c) Mice, Rats, Hamsters, Guinea pigs, Chinchillas: 0.5 mg/kg IM. Do not use in Gerbils. ()

Acepromazine dosage for cattle:

a) Sedation: 0.01-0.02 mg/kg IV or 0.03-0.1 mg/kg IM ()

b) 0.05 -0.1 mg/kg IV, IM or SC ()

c) Sedative one hour prior to local anesthesia: 0.1 mg/kg IM ()

Acepromazine dosage for horses:

(Note: ARCI UCGFS Class 3 Acepromazine)

a) For mild sedation: 0.01-0.05 mg/kg IV or IM. Onset of action is about 15 minutes for IV; 30 minutes for IM ()

b) 0.044-0.088 mg/kg (2-4 mg/100 lbs. body weight) IV, IM or SC (Package Insert; PromAce — Fort Dodge)

c) 0.02-0.05 mg/kg IM or IV as a preanesthetic ()

d) Neuroleptanalgesia: 0.02 mg/kg given with buprenorphine (0.004 mg/kg IV) or xylazine (0.6 mg/kg IV) ()

e) For adjunctive treatment of laminitis (developmental phase): 0.066-0.1 mg/kg 4-6 times per day ()

Acepromazine dosage for swine:

a) 0.1-0.2 mg/kg IV, IM, or SC ()

b) 0.03-0.1 mg/kg ()

c) For brief periods of immobilization: acepromazine 0.5 mg/ kg IM followed in 30 minutes by ketamine 15 mg/kg IM. Atropine (0.044 mg/kg IM) will reduce salivation and bronchial secretions. ()

Acepromazine dosage for sheep and goats:

a) 0.05-0.1 mg/kg IM ()


■ Cardiac rate/rhythm/blood pressure if indicated and possible to measure

■ Degree of tranquilization

■ Male horses should be checked to make sure penis retracts and is not injured

■ Body temperature (especially if ambient temperature is very hot or cold)

Client Information

■ May discolor the urine to a pink or red-brown color; this is not abnormal

■ Acepromazine is approved for use in dogs, cats, and horses not intended for food

Chemistry / Synonyms

Acepromazine maleate (formerly acetylpromazine) is a phenothiazine derivative that occurs as a yellow, odorless, bitter tasting powder. One gram is soluble in 27 mL of water, 13 mL of alcohol, and 3 mL of chloroform.

Acepromazine Maleate may also be known as: acetylpromazine maleate, “ACE”, ACP, Aceproject, Aceprotabs, PromAce, Plegicil, Notensil, and Atravet.

Storage / Stability/Compatibility

Store protected from light. Tablets should be stored in tight containers. Acepromazine injection should be kept from freezing.

Although controlled studies have not documented the compatibility of these combinations, acepromazine has been mixed with atropine, buprenorphine, chloral hydrate, ketamine, meperidine, oxymorphone, and xylazine. Both glycopyrrolate and diazepam have been reported to be physically incompatible with phenothiazines, however, glycopyrrolate has been demonstrated to be compatible with promazine HC1 for injection.

Dosage Forms / Regulatory Status

Veterinary-Labeled Products:

Acepromazine Maleate for Injection: 10 mg/mL for injection in 50 mL vials; Aceproject (Butler), PromAce (Fort Dodge); generic; (Rx). Approved forms available for use in dogs, cats and horses not intended for food.

Acepromazine Maleate Tablets: 5, 10 & 25 mg in bottles of 100 and 500 tablets; PromAce (Fort Dodge); Aceprotabs (Butler) generic; (Rx). Approved forms available for use in dogs, cats and horses not intended for food.

When used in an extra-label manner in food animals, it is recommended to use the withdrawal periods used in Canada: Meat: 7 days; Milk: 48 hours. Contact FARAD (see appendix) for further guidance.

The ARCI (Racing Commissioners International) has designated this drug as a class 3 substance. See the appendix for more information.

Human-Labeled Products: None


Postanesthetic Upper Respiratory Tract Obstruction

Upper respiratory tract () obstruction can occur in horses recovering from general anesthesia after various surgical procedures. Postanesthetic upper respiratory tract obstruction most often results from nasal edema and/or congestion and is usually mild. Other causes include arytenoid chondritis, dorsal displacement of the soft palate, and bilateral arytenoid cartilage paralysis. Bilateral arytenoid cartilage paralysis is relatively uncommon; however, it can result in severe upper respiratory tract obstruction with the horse becoming distressed, uncontrollable, and difficult to treat. The condition may rapidly become fatal, thus postanesthetic upper respiratory tract obstruction can be a serious complication after general anesthesia and surgery.

Etiology of Postanesthetic Upper Respiratory Tract Obstruction

Nasal Edema

Nasal edema and/or congestion is most often the result of venous congestion associated with a dependent head position during a prolonged anesthesia. Horses positioned in dorsal recumbency are thought to be more prone to nasal edema than horses in lateral recumbency. Nasal and pharyngeal edema may also result from trauma during endotracheal intubation that causes local inflammation and swelling.

Dorsal Displacement of the Soft Palate

Causes of dorsal displacement of the soft palate after ex-tubation are unknown. The condition is most likely a normal consequence of orotracheal intubation and of administration of sedative and anesthetic drugs that alter upper respiratory tract neuromuscular function. If dorsal displacement persists, it is most likely the result of an underlying upper respiratory tract problem or of inflammation in the pharynx secondary to intubation.

Arytenoid Chondritis

Arytenoid chondritis is an uncommon cause of postanesthetic upper respiratory tract obstruction but can be a longer-term consequence of traumatic intubation. Although this condition will not lead to obstruction in the same anesthetic period, it may at a later time if it is not recognized. Furthermore, the presence of an abnormal arytenoid will compromise the airway and can potentiate the possibility of an obstructive crisis.

Bilateral Laryngeal Paralysis

The etiology of postanesthetic bilateral laryngeal paralysis is unknown. Proposed etiologies include inflammation and edema of the larynx and neuromuscular failure. Physical trauma from endotracheal intubation or chemical irritation from residue after endotracheal tube cleaning may result in arytenoid chondritis, laryngeal dysfunction, and laryngeal inflammation and swelling. Laryngeal edema from venous congestion associated with a dependent head position during a prolonged anesthesia may cause swelling and failure of the arytenoid cartilages to adequately adduct. Causes of neuromuscular failure that lead to bilateral arytenoid cartilage paralysis include trauma to the cervical region or jugular vein; compression of the recurrent laryngeal nerve between the endotracheal tube or cuff and noncompliant neck structures; damage to the recurrent laryngeal nerve from intraoperative hypoxia, ischemia, or hypotension; and overextension of the neck when the horse is positioned in dorsal recumbency that causes damage to the recurrent laryngeal nerve as a result of compression of its blood supply.

α2-Adrenergic agonists have been shown to increase laryngeal asynchrony and increase upper airway resistance in horses. The muscle relaxant effects of xylazine are thought to decrease the tone of the supporting airway muscles, which in combination with low head carriage may cause an increase in airway resistance. The muscle relaxant effects of xylazine may have worn off at the time the horse has recovered from anesthesia; however, one study showed that upper airway resistance increased for 30 to 40 minutes after xylazine administration and then slowly returned to normal. Impaired laryngeal function associated with xylazine administration in combination with excitement associated with recovery from anesthesia and extubation may lead to dynamic collapse of the upper respiratory tract and result in the clinical signs described. Xylazine is a commonly used preanesthetic drug; therefore although it is unlikely to be the sole cause of the upper respiratory tract obstruction, it may be a contributing factor.

Underlying upper respiratory tract disease such as laryngeal hemiplegia may also predispose horses to severe postanesthetic obstruction. A few reports exist in the literature of severe postanesthetic upper respiratory tract obstruction in horses associated with laryngeal dysfunction. In two previous reports, bilateral arytenoid cartilage paralysis was associated with surgery in the head and neck region, and the horses recovered after establishment of a patent airway. These authors have recently seen several postanesthetic upper respiratory tract obstructions in horses that have undergone surgery for a variety of reasons including arthroscopy, tarsal arthrodesis, exploratory celiotomy, ovariohysterectomy, mastectomy, and prosthetic laryngoplasty/ventriculectomy. In addition to having undergone prosthetic laryngoplasty, some of these horses had a history of laryngeal hemiplegia before surgery. This fact suggests that preexisting disease may predispose to this condition. Postanesthetic upper respiratory tract obstruction in the horses at these authors’ hospital is often associated with excitement or exertion, including standing after anesthesia and vocalization. The cause of severe obstruction therefore could be laryngospasm or dynamic adduction of both paretic arytenoid cartilages into the airway during inspiration.

In the horses at these authors’ hospital, no association exists between difficult endotracheal intubation and upper respiratory tract obstruction. In horses that developed obstruction the duration of anesthesia was 90 to 240 minutes, and horses had mild-to-moderate hypotension, hypoventilation, and hypoxemia. These authors clean their endotracheal tubes with chlorhexidine gluconate between uses. If the tubes are not rinsed adequately, mucosal irritation from residual chlorhexidine gluconate could conceivably cause upper respiratory tract irritation and lead to obstruction. Most important, however, all these horses were positioned in dorsal recumbency for at least some of the time they were under anesthesia. The horses are positioned on a waterbed from the withers caudad. This position results in hyperextension of the neck and a dependent head position, both of which may predispose to postanesthetic bilateral arytenoid paralysis.

Negative-Pressure Pulmonary Edema

Pulmonary edema can result from upper respiratory tract obstruction and has been referred to as negative-pressure pulmonary edema because the pulmonary edema occurs secondary to strong inspiratory efforts against a closed airway. In humans vigorous inspiratory efforts against a closed glottis may create a negative pressure of as low as -300 mm Hg that, obeying Starling’s laws of fluid dynamics, fluid moves from the intravascular space into the interstitium and alveoli.

Clinical Signs

Although upper respiratory tract obstruction usually occurs immediately after extubation, severe obstruction associated with bilateral arytenoid paralysis may occur within 24 to 72 hours of recovery from anesthesia. The most obvious clinical sign is upper respiratory tract dyspnea. Horses with nasal edema have a loud inspiratory snoring noise, whereas horses with dorsal displacement of the soft palate have an inspiratory and expiratory snoring noise associated with fluttering of the soft palate. Horses with severe upper respiratory tract obstruction from bilateral laryngeal paralysis have a loud, high-pitched, inspiratory stri-dor associated with exaggerated inspiratory efforts.

Treatment of Postanesthetic Upper Respiratory Tract Obstruction

Nasal Edema

The most common type of upper respiratory tract obstruction is nasal edema, which often resolves rapidly without treatment. If obstruction is severe, it is critical to create a patent airway. The horse should be reintubated with a nasotracheal or orotracheal tube or 30-cm tubing placed in the nostrils to bypass the obstruction. Phenylephrine intranasal spray (5-10 mg in 10 ml water) or furosemide (1 mg/kg) may be used to reduce the nasal edema. Edema can be prevented by atraumatic intubation, reducing surgery time, and keeping the horse’s head elevated during anesthesia and surgery.

Dorsal Displacement of the Soft Palate

Dorsal displacement of the soft palate usually resolves spontaneously when the horse swallows, however, it may be corrected through induction of swallowing by gentle manipulation of the larynx or by insertion of a nasogastric tube into the pharynx.

Bilateral Laryngeal Paralysis

Severe obstruction often develops when the horse stands after being extubated. Emergency treatment is required because the horse will rapidly become severely hypoxic, develop cardiovascular collapse, and die. Horses are often difficult to treat because obstruction may not be noticed until the horse is severely hypoxic and uncontrollable. Treatment is then delayed until the horse collapses from hypoxia, however, emergency reintubation or tracheostomy is often too late.

Immediate treatment consists of rapid reintubation or tracheostomy. Horses may be reintubated with a nasotracheal tube (14-22 mm) or an orotracheal tube (20-26 mm). The clinician performs a tracheostomy by clipping, preparing, and blocking the ventral cervical region (if time permits), making a 8-cm vertical incision on midline at the junction of the upper and middle thirds of the neck, bluntly separating the sternothyrohyoideus muscles, and then making a transverse incision between the tracheal rings. These authors recommend having a kit available with a tracheostomy tube and drugs for reinduction of anesthesia (xylazine, 1.1 mg/kg; ketamine, 2.2 mg/kg; or a paralytic agent such as succinylcholine, 330 μg/kg IM). Horses should be treated with insufflation of oxygen immediately after establishment of an airway.

Prevention of upper respiratory tract obstruction after anesthesia requires treatment of hypotension, hypoxemia, and hypoventilation, avoidance hyperextension of the neck when horses are positioned in dorsal recumbency, and thorough rinsing of endotracheal tubes. These authors recover horses with the oral endotracheal tube in place, and following extubation closely monitor air movement.

If the horse has bilateral laryngeal paralysis, it may be necessary to establish a tracheostomy while the horse is treated aggressively with antiinflammatory treatment. Recovery should occur within days.

Negative-Pressure Pulmonary Edema

Previous reports have described successful treatment of negative-pressure pulmonary edema, however, treatment may fail if a delay occurs between obstruction and treatment or if an unknown underlying disease is present. Treatment of negative-pressure pulmonary edema consists of administration of oxygen through nasal insufflation (10-15 L/min for an adult horse), a diuretic (furosemide, lmg/kg IV, and mannitol, 0.5-1.0 g/kg IV), antiinflammatory agents (flunixin meglumine, 1.1 mg/kg; dexamethasone, 0.1-0.3 mg/kg; dimethyl sulfoxide [DMSO]; lg/kg), and the positive inotrope epinephrine (2-5 μg/kg). Fluid therapy with polyionic isotonic fluids and electrolytes should be administered, however, overhydration of horses with pulmonary edema must be avoided.


Management Of Dystocia

Materials required to correct a foaling problem may be as simple as an obstetrical sleeve, lubricant, and some baling twine. However it is common practice for a clinician to have on hand a pair of obstetrical chains (or straps) and handles or a Krey-Schotter hook, and a snare rod. Copious lubrication is often the key to success. A fetotome, wire, handles, guide, and a guarded scalpel are necessary to perform a fetotomy. Cleanliness is essential as is a large working area with good footing. The behavior of a foaling mare can be unpredictable and violent, thus safety for all personnel is an important consideration. Ideally the obstetrician should have access to a hospital facility where general anesthesia can be given and an overhead hoist system is available to lift the mare’s hindquarters.

The degree of restraint required for a safe examination and fetal extraction will vary with the individual mare. Although placement of a large-bore stomach tube or endotracheal tube into the mare’s trachea is reported to reduce straining, this procedure is of little benefit clinically. Application of a nose twitch or other methods of physical restraint offer limited help. Epidural anesthesia will reduce straining in the standing mare but the time needed to obtain an effective block precludes its routine use. Certainly the hindlimb ataxia that can be associated with an epidural is contraindicated if general anesthesia becomes necessary. Short-term xylazine-ketamine general anesthesia may not eliminate straining but will often permit positioning of the mare to facilitate manipulation of the fetus. Inhalation anesthesia will relax the mare and eliminate straining. Clinicians should be cautious about eliminating uterine contractions because they are beneficial to the delivery process after postural abnormalities of the fetus have been corrected.

Often there is insufficient space within the pelvic canal to permit correction of even simple fetal malpostures; thus repulsion of the fetus from the maternal pelvis back into the uterus is usually an integral part of dystocia correction. The degree of uterine contraction will influence the success of this procedure. Distention of the uterus with liquid obstetric lubricant often provides the extra space needed. If the mare is straining excessively, and/or the uterus is tightly contracted, administration of general anesthesia and elevation of the hindquarters is indicated. This method will reduce the amount of intraabdominal pressure on the uterus and permit the fetus to fall away from the pelvic canal. Elevation of the mare’s hindquarters allows the obstetrician to work at a more comfortable level and also eliminates the increased abdominal pressure that occurs if the mare is in lateral recumbency.

Because the value of a foaling mare may range from minimal to millions of dollars, it is impossible to be dogmatic about management of an obstetrical case. The economics of each case will play an important part in the decision process as the clinician contemplates the options — fetotomy, cesarean section, manipulation, and vaginal delivery. The breeding future of the mare must be considered because trauma to the genital tract will have an adverse effect on future fertility. Liberal application of lubricant is essential to protect the delicate membranes. Prolonged vaginal intervention is contraindicated in mares, because the cervix is easily traumatized. Slow traction while monitoring cervical stretching is recommended. If the mare is not under general anesthesia it is best to coordinate traction with the mare’s expulsive efforts.

Veterinary Medicine

Reptile Anesthesia

Restraint of reptiles may be performed without as much risk in the case of the debilitated animal in comparison to birds, for example. However, it is still worthwhile considering how to successfully and safely restrain the reptile patient in order for it to be anesthetised as well as some techniques that may make restraint less dangerous to animal and handler alike.

Initial Restraint Of The Reptile Patient

Points to consider include:

• Is the patient suffering from disease? Examples include patients with pneumonia, where mouth breathing and excessive oral mucus may be seen. Over-vigorous manual manipulation can exacerbate the problem.

• What is the species? Day geckos are extremely delicate and very prone to shedding their tails when handled. Similarly, species such as green iguanas are prone to conditions such as metabolic bone disease (MBD) whereby their skeleton becomes fragile and spontaneous fractures are common. Some reptiles are naturally aggressive, for example, snapping turtles, Tokay geckos and rock pythons. Other species are potentially dangerous, such as a venomous species of snake or lizard (e.g. rattlesnakes, cobras, Gila monsters and beaded lizards).

The need for restraint therefore needs to be considered carefully before physical attempts are made.

Restraint techniques and equipment

Order Sauria

This includes the members of the lizard family such as geckos, iguanas, chameleons and agamas.

Lizards come in many different shapes and sizes from the 1.2 m (4 foot) long adult green iguana, to the 10-12 cm (4-5 inch) long green anole. They have roughly all the same structural format with four limbs (although these may become vestigial in the case of the slow worm for example) and a tail. Their main danger areas therefore include their claws and teeth, and in some species, such as iguanas, their tails, which can lash out in a whip-like fashion.

Restraint is thus best performed by grasping the pectoral girdle with one hand from the ventral aspect, so controlling one forelimb with the thumb and the other between index and middle finger. The other hand is used to grasp the pelvic girdle from the dorsal aspect, controlling one limb with the thumb and the other again between index and middle finger. The lizard may then be held in a vertical manner with head uppermost to put the tail out of harm’s way. The handler should allow some flexibility in this method as the lizard may wriggle and overly rigid restraint could damage the spine.

The use of a thick towel to control the tail and claws is often very useful for aggressive lizards. Occasionally gauntlets are necessary for very large lizards, and for those that may have a poisonous bite (the Gila monster and the bearded lizard). It is important to assure that the lizard is not restrained with too much force, as those with skeletal problems such as metabolic bone disease may be seriously injured. In addition lizards, like other reptiles, do not have a diaphragm so over-zealous restraint will lead to the digestive system pushing on to the lungs and increasing inspiratory effort.

Geckos, as noted above, can be extremely fragile and the day geckos may be best anesthetised in a clear plastic container rather than physically restraining them. Other geckos have easily damaged skin so latex gloves and soft cloths should be used.

Small lizards may have their heads controlled between the index finger and thumb to prevent biting. Under no circumstances should lizards be restrained by their tail. Many will shed their tails if restrained in this way, but not all of them will regrow. Green iguanas, for example, will only regrow their tails as juveniles, i.e. <2.5-3 years of age. After this they will be left tail-less.

Vago-vagal reflex

There is a procedure which can be used to place members of the lizard and snake family into a trance-like state. It involves closing the eyelids of lizards (snakes eyelids are transparent and fused together over the eye as a ‘spectacle’) and placing firm but gentle digital pressure on to both eyeballs. Alternatively a ball of cotton wool may be placed over each eye and taped into place. This stimulates the parasympathetic autonomic nervous system which results in a reduction in heart rate, blood pressure and respiration rate. After 1-2 minutes of this, providing no loud noises are made, the lizard / snake may be placed on its side, front, back etc., so allowing radiography to be performed without further physical or chemical restraint. A loud noise or physical stimulation will immediately cause the lizard / snake to revert to its normal wakeful state.

Order Serpentes

This is the snake family. This order encompasses a wide range of sizes from the enormous anacondas and Burmese pythons, which may achieve lengths up to 9 m (30 feet) or more, down to the thread snake family, which may only be a few tens of centimetres long. However, they are all characterised by their elongated form with an absence of limbs. Their danger areas are their teeth (and in the case of the more poisonous species such as the viper family their fang teeth) and, in the case of the constrictor and python family, their ability to asphyxiate their prey by winding themselves around the victim’s chest / neck. Non-venomous snakes can be restrained by initially controlling the head. This is done by placing the thumb over the occiput and curling the fingers under the chin. Reptiles, like birds, have only the one occipital condyle so it is important to stabilise the neck occipital / atlantal joint. It is also important to support the rest of the snake’s body so that not all of the weight of the snake is suspended from the head. This is best achieved by allowing the smaller species to coil around the handler’s arm, so the snake is supporting itself. In larger species it is necessary to support the body length at regular intervals, so the help of several people may be necessary. Above all it is important not to grip the snake too hard as this will cause bruising and the release of myoglobin from muscle cells that will damage the kidney filtration membranes.

Poisonous species (such as the viper family, rattlesnakes etc.) or very aggressive species (such as anacondas, reticulated and rock pythons) may be restrained using snake hooks. These are 45-60cm (1.5-2 foot) long steel rods with a blunt-ended shepherd’s hook on the end and are used to loop under the body of a snake to move it at arm’s length into a container. The hook may also be used to trap the head flat to the floor before grasping it with the hand. Once the head is controlled safely the snake is rendered harmless unless it is a member of the spitting cobra family. These are unlikely to be encountered in general practice, but if they are, plastic goggles should be worn, as the poison is spat into the prey / assailant’s eyes, causing blindness.

Order Chelonia

This includes all land tortoises, terrapins and aquatic turtles, varying in size from the small Egyptian tortoises, weighing a few grams, to Galapagos species, weighing several hundred kilograms. The majority are harmless, although surprisingly strong. Exceptions include the snapping turtle and the alligator snapping turtle, both of which can give a serious bite, as can most of the soft-shelled terrapins. They also have mobile necks. Even red-eared terrapins may give a nasty nip!

For the mild-tempered Mediterranean species, the tortoise may be held with both hands, one on either side of the main part of the shell behind the front legs. For examination to keep the tortoise still he / she may be placed on to a cylinder / stack of tins which ensure that his / her legs are raised clear of the table, effectively balancing the tortoise on the centre of the underside of the shell (plastron).

For aggressive species it is essential to hold the shell on both sides behind and above the rear legs to avoid being bitten. For examination of the head region in these species it is necessary to chemically restrain them.

For the soft-shelled and aquatic species, soft cloths and latex gloves may be necessary in order not to mark the shell.

Order Crocodylia

This family is rarely seen in general practice and includes the crocodiles, both fresh and saltwater, the alligators, the fish-eating gharials and the caimans. Their dangers lie in their impressively arrayed jaws and their sheer size – an adult bull Nile crocodile may weigh many hundreds of kilograms and may live for up to 50 years or more. Readers are advised to consult standard texts for further information.

Aspects Of Chemical Restraint

Additional Supportive Therapy

Monitoring Anesthesia

Recovery And Analgesia

Reptiles often will recover rapidly from isoflurane anesthesia, but if other injectable drugs were used, recovery may be prolonged. It is thus essential to keep the reptile patient calm, stress free and at its optimum preferred body temperature. It may be necessary to keep the patient intubated with IPPV with oxygen if high doses of injectable agents have been used, until the reptile is once again breathing for itself. Care should be taken though, as high levels of oxygen actually suppress the stimulus for a reptile to breathe, as this is induced by a low level of PaO2 rather than high PaCO2 as in mammals. The use of doxapram at dose of 5 mg / kg IM or IV helps stimulate respiration.

Fluid therapy also helps to speed recovery, especially with agents such as ketamine which are cleared through the kidneys. Once recovery occurs, the patient should be encouraged to eat; if anorectic, the patient should be assist fed or stomach tubed.

The provision of analgesia, as is the case for other animal groups, is an important aspect of postoperative recovery (Table Commonly used analgesics in reptiles). Reptiles that have been given analgesia have been shown to have a quicker return to normality, i.e. eating, normal behaviour etc., than those who do not receive analgesia.

Table Commonly used analgesics in reptiles.

(SC = subcutaneously; IM = intramuscularly; IV = intravenously; PO = peros; q8h = every 8 hours; q24h = every 24 hours and so on)

Analgesic Dosage Dosage interval
Butorphanol 0.4 mg / kg SC, IM q6-8h
Buprenorphine 0.01 mg / kg SC, IM q8-12h
Meloxicam 0.1-0.2 mg / kg SC, IM, PO q24h
Carprofen 2-4 mg / kg SC, PO q24h (first day 4 mg / kg then decrease to 2 mg / kg)


Opioids have been shown to have some effect in reptiles: butorphanol at 0.4 mg / kg IM, IV or subcutaneously and buprenorphine at 0.02 mg / kg intramuscularly have been recommended.

Non-steroidal anti-inflammatory drugs (NSAIDs) also seem to be beneficial in reptiles. Carprofen at doses of 2-4 mg / kg intramuscularly initially, and then 1-2 mg / kg every 24-72 hours thereafter, and meloxicam at 0.1-0.2 mg / kg orally every 24 hours have both been recommended. It should be noted that all NSAIDs are potentially nephrotoxic and can have gastrointestinal ulcerative side effects, so fluid therapy and close monitoring are required.

Veterinary Medicine

Aspects Of Chemical Restraint

Chemical restraint is often necessary in reptile medicine to facilitate procedures from simply extracting the head of a leopard tortoise or box turtle, to enable a jugular blood sample to be performed, to coeliotomy procedures such as surgical correction of egg-binding.

Before any anesthetic / sedative is administered, an assessment of the reptile patient’s health is necessary. Is sedation / anesthesia necessary for the procedure required? Is the reptile suffering from respiratory disease or septicaemia, i.e. is the reptile’s health likely to be made worse by sedation / anesthesia?

Before any attempt to administer chemical restraint the reptilian respiratory system should be understood.

Overview of reptilian anatomy and physiology relevant to anesthesia

The reptilian patient has a number of variations from the basic mammalian anatomical and physiological systems. Starting rostrally:

(1) Reduced larynx: The reptile patient does have a glottis similar to the avian patient, which lies at the base of the tongue, more rostrally in snakes and lizards and more caudally in Chelonia. At rest the glottis is permanently closed, opening briefly during inspiration and expiration. In crocodiles the glottis is obscured by the basihyal valve which is a fold of the epiglottis that has to be deflected before they can be intubated.

(2) The trachea varies between orders: The Chelonia and Crocodylia have complete cartilaginous rings similar to the avian patient, with the Chelonia patient having a very short trachea, bifurcating into two bronchi in the neck in some species. Serpentes and Sauria have incomplete rings such as is found in the cat and dog, with Serpentes species having a very long trachea. Many Serpentes species have a tracheal lung – an outpouching from the trachea as a form of air sac.

(3) The lungs of Serpentes and saurian species are simple and elastic in nature. The left lung of most Serpentes species is absent or vestigial but may be present in the case of members of the Boid family (boa constrictors etc.). The right lung of Serpentes species ends in an air sac. Chelonia species have a more complicated lung structure, and the paired lungs sit dorsally inside the carapace of the shell. Crocodylia have lungs not dissimilar to mammalian ones and they are paired.

(4) No reptile has a diaphragm: Crocodylia species have a pseudodiaphragm, which changes position with the movements of the liver and gut, so pushing air in and out of the lungs.

(5) Most reptiles use intercostal muscles to move the ribcage in and out, as with birds: The exception being the Chelonia. These species require movement of their limbs and head into and out of the shell in order to bring air into and out of the lungs. This is important when they are anesthetised as such movements, and therefore breathing, cease.

(6) Some species can survive in oxygen-deprived atmospheres for prolonged periods: Chelonia species may survive for 24 hours or more and even green iguanas may survive for 4-5 hours, making inhalation induction of anesthesia almost impossible in these animals.

(7) Reptiles have a renal portal blood circulation system: This means that the blood from the caudal half of the body can pass through the kidney structure before passing into the caudal major veins and entering the heart. Therefore, if drugs that are excreted by the kidneys are injected into the caudal half of the body, then they may be excreted before they have a chance to work systemically (e.g. ketamine). In addition, if a drug is nephrotoxic (e.g. the aminoglycosides) then injection into the caudal half of the body may increase the risk of renal damage.

Pre-anesthetic preparation

Blood testing

It is useful to test biochemical and haemocytological parameters prior to administering chemical immobilising drugs. Blood samples may be taken from the jugular vein or dorsal tail vein in Chelonia, the ventral tail vein, palatine vein or by cardiac puncture in Serpentes and the ventral tail vein in Crocodylia and Sauria. Minimal testing advised is a haematocrit, blood calcium levels, blood total protein levels, aspartate transaminase (AST) levels for hepatic function and uric acid levels for renal function.


This is necessary prior to anesthesia in Serpentes (for a period of 2 days in small snakes up to 1-2 weeks for the larger pythons) to prevent regurgitation and pressure on the lungs / heart. Chelonia rarely regurgitate and do not need prolonged fasting. It is important not to feed live prey to insectivores (e.g. leopard geckos) within 24 hours of anesthesia as the prey may still be alive when the reptile is anesthetised!

Pre-anesthetic medications

Antimuscarinic drugs

Atropine (0.01-0.04mg / kg intramuscularly (IM)) or glycopyrrolate (0.01 mg / kg IM) can reduce oral secretions and prevent bradycardia. However, these problems are rarely of concern in reptiles.


Midazolam has been used in red-eared terrapins at 1.5 mg / kg as a premedicant and produced adequate sedation to allow minor procedures and induction of anesthesia.


Acepromazine (0.1-0.5 mg / kg intramuscularly (IM)) may be given 1 hour before induction of anesthesia to reduce the dose of induction agent required. Diazepam (0.22-0.62mg / kg IM in alligators) and midazolam (2mg / kg IM in turtles) are also useful.

Alpha-2 adrenoceptor agonists

Xylazine can be used 30 minutes prior to ketamine at 1 mg / kg in Crocodylia to reduce the dose of ketamine required. Medetomidine may be used at doses of 100-150 μ.g / kg, also reducing the required dose of ketamine in Chelonia, and it has the advantage of being reversible with atipamezole at 500-750 µg / kg.


Butorphanol (0.4mg / kg intramuscularly (IM)), can be administered 20 minutes before anesthesia, providing analgesia and reducing the dose of induction agent required. It may be combined with midazolam at 2mg / kg.

Induction of Anesthesia

Maintenance of anesthesia with injectable agents


This may be used on its own for anesthesia at doses of 55-88 mg / kg intramuscularly (IM). As the dose increases, so the recovery time also increases, in some instances to several days; doses above 110 mg / kg will cause respiratory arrest and bradycardia.

Ketamine may be combined with other injectable agents to provide surgical anesthesia.

Suitable agents include: midazolam at 2 mg / kg intramuscularly (IM) with 40 mg / kg ketamine in turtles; xylazine at 1 mg / kg IM, given 30min prior to 20mg / kg ketamine in large crocodiles and medetomidine at 0.1mg / kg IM with 50mg / kg ketamine in king snakes.

Ketamine at 5mg / kg has been combined with medetomidine at 0.1mg / kg intravenously (IV) to produce a short period of anesthesia in gopher tortoises although some hypoxia was observed and supplemental oxygen is advised.


Propofol may be used to give 20-30 min of anesthesia, which may allow minor procedures. It may be topped up at 1 mg / kg / min IV or intraosseously. Apnoea is extremely common and intubation and ventilation with 100% oxygen are advised.

Maintenance of anesthesia with inhalational agents

Maintaining Anesthesia

Veterinary Medicine

Induction of Anesthesia

Injectable agents

Advantages of injectable anesthetics include ease of administration, avoiding problems related to breath-holding and prolonged induction, low cost and good availability. Disadvantages include a recovery often dependent on organ metabolism, difficulty reversing medications in emergency situations, prolonged recovery periods and necrosis of muscle cells at injection sites. Also, due to the renal portal system, drugs injected into the caudal half of a reptile may either be excreted by the kidneys before they take effect, or increase renal damage if the case of nephrotoxic agents. Injectable medications should therefore be administered in the cranial half of the body.


The effects of ketamine are species and dose dependent. Recommended levels range from 22-44mg / kg intramuscularly (IM) for sedation and 55-88 mg / kg IM for surgical anesthesia. Lower doses are required if the drug is combined with a premedicant such as midazolam or medetomidine.

Anaesthesia is induced in 10-30 minutes but may take up to 4 days to wear off, particularly at low environmental temperatures. Therefore, ketamine is mainly used at lower doses to sedate the animal and allow intubation and maintenance of anesthesia using inhalational agents in species such as Chelonia, which may breath-hold.

Disadvantages of ketamine include pain on administration and renal excretion: due to the renal portal system administration of ketamine in the cranial half of the body is recommended.


Propofol produces rapid induction of and recovery from anesthesia, and is becoming the induction agent of choice. Its advantages include a short elimination half-life and minimal organ metabolism, making it safer in debilitated reptiles. Disadvantages include the need for intravenous administration, although use of the intraosseous route has been shown to be successful in green iguanas at a dose of 10 mg / kg. Propofol does produce transient apnoea and cardiac depression, often necessitating positive pressure ventilation.

Doses of 10-15 mg / kg of propofol administered via the dorsal coccygeal (tail) vein in Chelonia have successfully induced anesthesia in under 1 minute. Used alone this will provide a period of anesthesia of 20-30 minutes or will allow intubation and maintenance of anesthesia using inhalational agents.


This is a neuromuscular blocking agent and produces immobilisation without analgesia. It should only be used to aid the administration of another form of anesthetic, or for transportation and not as a sole method of anesthesia.

It can be used in large Chelonia at doses of 0.5-1 mg / kg intramuscularly (IM) and will allow intubation and conversion to inhalational anesthesia. Crocodilians can be immobilised with 3-5 mg / kg IM, with immobilisation occurring within 4 minutes and recovery in 7-9 hours. Respiration usually continues without assistance at these doses, but is important to have assisted ventilation facilities to hand as paralysis of the muscles of respiration can easily occur.

Inhalational agents

The usual inhalational agents may be used to induce anesthesia, either in an induction chamber or via face masks. Face masks may be either bought, or for snakes, syringe cases may be modified to form elongated masks for induction.

Veterinary Medicine

Maintaining Anesthesia

Inhalational anesthesia is becoming the main method of anesthetising reptiles for prolonged procedures and as described above offers many benefits. These are enhanced still further if the reptile is intubated allowing the anesthetic to be delivered in a controlled manner.


The glottis, which acts as the entrance to the trachea, is relatively rostral in many species. It is kept closed at rest, so the anesthetist must wait for inspiration to occur to allow intubation. Reptiles produce little or no saliva when not eating, so blockage of the endotracheal tube is uncommon.

In Serpentes, the glottis sits rostrally on the floor of the mouth caudal to the tongue sheath. Intubation may be performed consciously if necessary, as reptiles do not have a cough reflex. The mouth is opened with a wooden or plastic tongue depressor and the endotracheal tube inserted during inspiration. Alternatively, an induction agent may be given and then intubation performed.

In Chelonia, the glottis sits caudally at the base of the tongue. The trachea is very short and the endotracheal tube should only be inserted a few centimetres otherwise there is a risk one or other bronchus will be intubated, leading to only one lung receiving the anesthetic. As mentioned above, an induction agent such as ketamine or propofol is advised for Chelonia prior to intubation due to breath-holding and difficulty in extracting the head from the shell.

Sauria vary depending on the species, most having just a glottis guarding the entrance to the trachea. However, some species possess vocal folds, e.g. geckos. Some may be intubated consciously, but most are better induced with an injectable agent or following mask or chamber induction. Some species may be too small for intubation.

Crocodylia have a basihyal fold which acts like an epiglottis and needs to be depressed prior to intubation. These species, due to the risk associated with handling them, require some form of chemical restraint prior to intubation.

Intermittent positive pressure ventilation

If intubation is performed fully conscious, anesthesia may be induced, even in breath-holding species, by using positive pressure ventilation, in a matter of 5-10 min. This does have some advantages as avoiding injectable induction agents leads to rapid postoperative recovery.

Many, if not all species of reptile require positive pressure ventilation during the course of an anesthetic. Chelonia for example are frequently placed in dorsal recumbency during intracoelomic surgery (the absence of a diaphragm leads to one body cavity – the coelomic cavity). As they have no diaphragm and the lungs are situated dorsally, the weight of the digestive contents pressing on the lungs will reduce inspiration and lead to hypoxia. In addition, movement of the chelonian’s limbs induces most of the normal inspiratory effort: this should be abolished during anesthesia! Furthermore, if a neuromuscular blocking agent such as succinylcholine has been used, positive pressure ventilation may be needed due to the potential for respiratory muscle paralysis.

The aims of intermittent positive pressure ventilation (IPPV) are to just inflate the lungs for an adequately oxygenated state to be maintained and for the animal to remain anesthetised. Most reptiles require 4-8 breaths a minute, with inflation pressures of no more than 10-15 cmH2O (7-11 mmHg). A ventilator unit is useful, but with experience, manual ventilation of the patient with enough pressure to just inflate the lungs and no more, can be achieved. A rough guide is to inflate the first two fifths of the reptile’s body at each cycle.

Anaesthetic breathing systems

For species less than 5 kg a non-rebreathing system with oxygen flow at twice the minute volume (which approximates to 300-500 ml / kg / min for most species) is suggested. Ayres T-pieces, modified Bain circuits or Mapleson C circuits may all be used.