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.


Diseases of the Throat: Diagnosis

Diagnostic Imaging

Lateral and ventrodorsal radiographic views of both the skull and cervical areas are indicated. Radiopaque foreign bodies can be identified that may be missed on laryngoscopy and pharyngoscopy (e.g. sewing needle embedded in soft tissues). Radiographs are also useful in identifying bony changes associated with chronic inflammation or neoplasia, identifying clues of unreported trauma (e.g. subcutaneous emphysema), and occasionally soft tissue masses. Suggestion of a soft tissue mass is confirmed by direct visualization and histopathology. Thoracic radiographs are also indicated. Symptoms of lower respiratory disease may be masked when a patient has concurrent, and more severe, upper respiratory symptoms. Evaluation for aspiration pneumonia, metastases, or suggestion of a motility disorder (i.e. megaesophagus) is possible.

Ultrasonography and computed tomography (CT) are noninvasive modalities to evaluate the pharynx and larynx. Ultrasonography can identify soft tissue masses, help guide fine needle aspiration, and evaluate laryngeal function. The presence of air in these areas can limit the usefulness of this modality in establishing a definitive diagnosis. CT may be used to fully evaluate involvement of neoplasia or middle-ear disease if a nasopharyngeal polyp is suspected.

Videofluoroscopy is essential for any case of dysphagia. A barium swallow allows the act of swallowing to be recorded and studied for abnormalities. The patient should be recorded attempting to swallow barium to mimic liquids and then should be given a meal (canned food mixed with barium) to be recorded. Videofluoroscopy is superior to radiography because it allows all phases of deglutition to be evaluated instead of recording one moment (intermittent moments) of the event. Unfortunately videofluoroscopy is limited to referral centers only.

Pharyngoscopy and Laryngoscopy

Laryngoscopy and pharyngoscopy allow assessment of both structural abnormalities and function of the larynx. A flexible endoscope is used for these procedures because visualization of the nasopharynx requires retroflexion. Occasionally a foreign body will be found just caudal to the larynx and may be retrieved endoscopically. The patient is placed in sternal recumbency and anesthetized with either propofol or sodium thiopental. Once anesthetized, gauze is passed under the maxilla behind the canine teeth. The gauze is used to elevate the head, so external compression of the neck is avoided. Flexible endoscopy is ideal to evaluate the nasopharynx. If that is not possible, the caudal pharynx can be evaluated using a dental mirror and a snook hook. This will be sufficient in evaluating most nasopharyngeal polyps, masses, or caudal foreign bodies. It will not allow diagnosis of more rostral diseases such as nasopharyngeal stenosis. Laryngeal function is usually evaluated first by assessing the motion of the arytenoid cartilages. The traditional approach involves titrating anesthesia that allows both visualization of the arytenoid cartilages and deep spontaneous breaths to occur. In a normal animal the arytenoid cartilages will abduct symmetrically with each inspiration and close on expiration. The frustration with this technique is multiple. Maintaining the correct level of anesthesia is difficult (i.e. the animal is too awake to allow adequate visualization of the arytenoid cartilages or anesthetized so that the patient will not spontaneously breathe); shallow breathing can limit adequate assessment; and concerns about the effect of anesthesia on laryngeal function are legitimate concerns when performing the traditional laryngeal examination. The recently introduced technique attempts to eliminate the effects of anesthesia from the examination. Patients are premedicated with acepromazine maleate and butorphanol tartrate and induced with propofol. Doxapram hydrochloride (2.2 mg/kg intravenously) is used to increase laryngeal motion and minimize or eliminate the effects of anesthesia,


Hematology and biochemical profiles should be performed on patients with pharyngeal and laryngeal dysfunction, but they will rarely confirm the definitive diagnosis. Occasionally virus isolation (feline calicivirus (FCV)) and PCR (feline herpes-1 virus (FHV-1), Chlantydia spp. and Mycoplasma spp.) are indicated in the diagnostic workup. Culture and sensitivity of tissue or secretions can provide valuable information during the diagnostic workup. Cytology and histopathology are also essential for critically evaluating infiltrative disease or mass lesions.


Alfentanil HCL (Alfenta)

Opiate Anesthetic Adjunct

Highlights Of Prescribing Information

Injectable, potent opiate that may be useful for adjunctive anesthesia, particularly in cats

Marginal veterinary experience & little published data available to draw conclusions on appropriate usage in veterinary species

Dose-related respiratory & CNS depression are the most likely adverse effects seen

Dose may need adjustment in geriatric patients & those with liver disease

Class-ll controlled substance; relatively expensive

What Is Alfentanil HCL Used For?

An opioid analgesic, alfentanil may be useful for anesthesia, analgesia, or sedation similar to fentanyl; fentanyl is generally preferred because of the additional experience with its use in veterinary patients and cost. Alfentanil may be particularly useful in cats as adjunctive therapy during anesthesia to reduce other anesthetic (i.e., propofol or isoflurane) concentrations.


Alfentanil is a potent mu opioid with the expected sedative, analgesic, and anesthetic properties. When comparing analgesic potencies after IM injection, 0.4-0.8 mg of alfentanil is equivalent to 0.1-0.2 mg of fentanyl and approximately 10 mg of morphine.


The pharmacokinetics of alfentanil have been studied in the dog. The drug’s steady state volume of distribution is about 0.56 L/kg, clearance is approximately 30 mL/kg/minute, and the terminal half-life is approximately 20 minutes.

In humans, onset of anesthetic action occurs within 2 minutes after intravenous dosing, and within 5 minutes of intramuscular injection. Peak effects occur approximately 15 minutes after IM injection. The drug has a volume of distribution of 0.4-1 L/kg. About 90% of the drug is bound to plasma proteins. Alfentanil is primarily metabolized in the liver to inactive metabolites that are excreted by the kidneys into the urine; only about 1% of the drug is excreted unchanged into the urine. Total body clearance in humans ranges from 1.6-17.6 mL/minute/kg. Clearance is decreased by about 50% in patients with alcoholic cirrhosis or in those that are obese. Clearance is reduced by approximately 30% in geriatric patients. Elimination half-life in humans is about 100 minutes.

Before you take Alfentanil HCL

Contraindications / Precautions / Warnings

Alfentanil is contraindicated in patients hypersensitive to opioids. Because of the drug’s potency and potential for significant adverse effects, it should only be used in situations where patient vital signs can be continuously monitored. Initial dosage reduction may be required in geriatric or debilitated patients, particularly those with diminished cardiopulmonary function.

Adverse Effects

Adverse effects are generally dose related and consistent with other opiate agonists. Respiratory depression, and CNS depression are most likely to be encountered. In humans, bradycardia that is usually responsive to anticholinergic agents can occur. Dose-related skeletal muscle rigidity is not uncommon and neuromuscular blockers are routinely used. Alfentanil has rarely been associated with asystole, hypercarbia and hypersensitivity reactions.

Respiratory or CNS depression maybe exacerbated if alfentanil is given with other drugs that can cause those effects.

Reproductive / Nursing Safety

In humans, the FDA categorizes alfentanil 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). If alfentanil is administered systemically to the mother close to giving birth, offspring may show behavioral alterations (hypotonia, depression) associated with opioids. Although high dosages given for 10-30 days to laboratory animals have been associated with embryotoxicity, it is unclear if this is a result of direct effects of the drug or as a result of maternal toxicity secondary to reduced food and water intake.

The effects of alfentanil on lactation or its safety for nursing offspring is not well defined, but it is unlikely to cause significant effects when used during anesthetic procedures in the mother.

Overdosage / Acute Toxicity

Intravenous, severe overdosages may cause circulatory collapse, pulmonary edema, seizures, cardiac arrest and death. Less severe overdoses may cause CNS and respiratory depression, coma, hypotension, muscle flaccidity and miosis. Treatment is a combination of supportive therapy, as necessary, and the administration of an opiate antagonist such as naloxone. Although alfentanil has a relatively rapid half-life, multiple doses of naloxone may be necessary. Because of the drug’s potency, the use of a tuberculin syringe to measure dosages less than 1 mL with a dosage calculation and measurement double-check system, are recommended.

How to use Alfentanil HCL

(Note: in very obese patients, figure dosages based upon lean body weight.)

Alfentanil HCL dosage for dogs:

As a premed:

a) 5 mcg/kg alfentanil with 0.3-0.6 mg of atropine IV 30 seconds before injecting propofol can reduce the dose of propofol needed to induce anesthesia to 2 mg/kg, but apnea may still occur. ()

As an analgesic supplement to anesthesia:

a) 2-5 mcg/kg IV q20 minutes. ()

b) For intra-operative analgesia in patients with intracranial disease: 0.2 mcg/kg/minute ()


■ Anesthetic and/or analgesic efficacy

■ Cardiac and respiratory rate

■ Pulse oximetry or other methods to measure blood oxygenation when used for anesthesia

Client Information

■ Alfentanil is a potent opiate that should only be used by professionals in a setting where adequate patient monitoring is available

Chemistry / Synonyms

A phenylpiperidine opioid anesthetic-analgesic related to fentanyl, alfentanil HCL occurs as a white to almost white powder. It is freely soluble in alcohol, water, chloroform or methanol. The commercially available injection has a pH of 4-6 and contains sodium chloride for isotonicity. Alfentanil is more lipid soluble than morphine, but less so than fentanyl.

Alfentanil may also be known as: alfentanyl, Alfenta, Fanaxal, Fentalim, Limifen, or Rapifen.

Storage / Stability/Compatibility

Alfentanil injection should be stored protected from light at room temperature. In concentrations of up to 80 mcg/mL, alfentanil injection has been shown to be compatible with Normal Saline, D5 in Normal Saline, D5W, and Lactated Ringers.

Dosage Forms / Regulatory Status

Veterinary-Labeled Products: None

The ARCI (Racing Commissioners International) has designated this drug as a class 1 substance. See the appendix for more information.

Human-Labeled Products:

Alfentanil HCL for injection: 500 mcg/mL in 2 mL, 5 mL, 10 mL, and 20 mL amps; preservative free; Alfenta (Akorn); Alfentanil HCL (Abbott); (Rx, C-II).

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.

Veterinary Medicine

Portosystemic Shunts

1. What is a portosystemic shunt?

A portosystemic shunt is an abnormal vessel that connects the portal vein to a systemic vein. The most common locations for portosystemic shunts are a patent ductus venosus or a connection between the portal vein and caudal vena cava or azygous vein. Single extraheptic shunts are most common in small-breed dogs and cats, whereas single intrahepatic shunts are most common in large-breed dogs.

2. What is the difference between congenital and acquired portosystemic shunts?

Most acquired shunts are multiple and extrahepatic. Acquired shunts develop because of sustained portal hypertension from chronic liver disease and cirrhosis. Congenital portosystemic shunts are usually single and may be intra- or extrahepatic. The most common intrahepatic portosystemic shunt is a patent ductus venosus.

3. Are certain breeds associated with portosystemic shunts?

Congenital portosystemic shunts may occur in any breed of dog but are common in miniature schnauzers, miniature poodles, Yorkshire terriers, dachshunds, Doberman pinschers, golden retrievers, Labrador retrievers, and Irish setters. There are affected lines in miniature schnauzers, Irish wolfhounds, Old English sheepdogs, and Cairn terriers. Mixed breed cats are more commonly affected than purebred cats, but Himalayans and Persians seem to overrepresented as purebreds. Acquired portosystemic shunts are secondary to chronic hepatic disease and so may occur in any breed.

4. Where are most portosystemic shunts located?

Single extrahepatic shunts most commonly connect the portal vein (or the left gastric or splenic vein) with the caudal vena cava cranial to the phrenicoabdominal vein. Single intrahepatic shunts can be a communication of the portal vein to the caudal vena cava which is a failure of the ductus venosus to close. Shunts in the right medial or lateral liver lobes occur with an unknown pathogenesis.

5. Why do patients with portosystemic shunts have decreased liver function?

Portal venous blood is important because it brings hepatotropic growth factors and insulin to the liver. If insulin bypasses the liver in a shunt, significant quantities are utilized by other organs and the liver receives less benefit. Portal venous blood flow is important for normal liver development as well as glycogen storage, hypertrophy, hyperplasia, and regeneration. Congenital portosystemic shunts are often associated with hepatic atrophy, hypoplasia, and dysfunction.

6. What are the most common clinical signs of portosystemic shunts?

Failure to thrive and failure to gain weight are appropriately common. Most clinical signs are referable to hepatic encephalopathy, which is defined as clinical signs of neurologic dysfunction secondary to hepatic disease. Signs include ataxia, stupor, lethargy, unusual behavior, disorientation, blindness, and seizures. Some animals display anorexia, vomiting, and diarrhea. Polyuria and polydipsia may be present. Some animals have ammonium biurate urolithiasis, which may result in pollakiuria, hematuria, stranguria, or obstruction. Increased production of saliva (ptyalism) and abdominal distention due to ascites occur in some animals. Ptyalism is more common in cats.

7. What causes hepatic encephalopathy associated with portosystemic shunts?

Products of bacterial metabolism in the intestine, such as ammonia, short-chain fatty acids (SCFAs), mercaptans, gamma-aminobutyric acid (GABA), and endogenous benzodiazepines have been suggested as mediators of hepatic encephalopathy. In addition, the ratio of aromatic amino acids to branched-chain amino acids is often increased in patients with portosystemic shunts. The aromatic amino acids may act as false neurotransmitters. Phenylalanine and tyrosine may act as weak neurotransmitters in the presynaptic neurons of the CNS. Tryptophan causes increased production of serotonin, which is a potent inhibitory neurotransmitter. The GABA receptor has binding sites for barbiturates, benzodiazepines, and substances with similar chemical structure to benzodiazepines. These agents may be responsible for depression of the CNS in hepatic encephalopathy.

8. What factors may precipitate an hepatic encephalopathy crisis?

A protein rich meal, gastrointestinal bleeding associated with parastites, ulcers or drug therapy; administration of methionine- containing urinary acidifiers; or lipotropic agents may precipitate a crisis. Blood transfusions with stored blood may also contribute to a crisis as the ammonia levels can be high in the stored blood.

9. How is hepatic encephalopathy treated?

The animal should be evaluated for hypoglycemia immediately and treated appropriately if it is present. Appropriate fluid therapy based on acid-base and electrolyte status (see chapter 81) should be initiated to correct abnormalities. LRS should be avoided. Hypoglycemia, alkalosis, hypokalemia, and gastrointestinal bleeding should be identified and corrected. Ammonia concentration and production should be decreased by administering lactulose and neomycin (10-20 mg/kg orally every 6 hr) if a swallow response is present. Oral metronidazole may be used at a dose of 10 mg/kg every 8 hr in place of neomycin. If the animal is comatose, 20-30 ml/kg of lactulose diluted 1:2 with water or a 1:10 dilution of povidone-iodine solution may be given as an enema. Seizures may be treated initially with elimination of ammonia by enemas as listed above. Oral loading doses of potassium bromide may be useful. If seizures cannot be controlled, IV propofol as a constant rate infusion may be necessary, but respiratory support may be needed. Some animals with hepatic encephalopathy have difficulty in metabolizing benzodiazepines such as diazepam, which should be avoided. If these drugs do not control seizures, intravenous phenobarbital may be titrated slowly to effect. Patients often have decreased clearance of barbiturates.

10. What routine blood work and urinalysis abnormalities suggest portosystemic shunts?

Microcytosis is a consistent abnormality of complete blood cell count in animals with portosystemic shunts. Some animals manifest acid-base, electrolyte, and glucose disturbances (hypoglycemia). Because of vomiting and dehydration, prerenal azotemia may be present. There is no consistent finding with regard to alanine aminotransferase (ALT), aspartate aminotransferase (AST), and serum alkaline phosphatase (ALP); activities of these enzymes may be elevated, decreased, or normal in patients with portosystemic shunts. Hypoalbuminemia is common, as are coagulopathies. Some animals have isothenuric urine due to medullary wash-out; ammonium biurate crystals may be identified on microscopic examination of urine sediment.

11. What are the best ways to diagnose a portosystemic shunt?

Elevated serum pre- and postprandial bile acids in a young animal with signs of hepatic encephalopathy and stunted growth are consistent with but not diagnostic for portosystemic shunts. A nuclear medicine scan using transcolonic sodium pertechnetate Tc99m demonstrates radioactivity in the heart before the liver in an animal with portosystemic shunt. Nuclear medicine is rapid, noninvasive, and safe to the animal. The disadvantages are that the animal is radioactive for 24 hours, studies can be performed only by specially trained personnel, exact location of the shunt cannot be determined, and cases of hepatic microvascular dysplasia, which have shunting within the liver (as in Cairn terriers), may give false-negative results. When nuclear medicine facilities are unavailable, positive contrast portography may demonstrate the anomalous vessel. Portography, however, is technically demanding and invasive. Furthermore, a second surgical procedure is required to repair the shunt because of an otherwise dangerously long period of anesthesia. The major advantage of positive contrast portography is that it definitively locates the shunt.

12. What is the best way to manage a patient with portosystemic shunt?

Although medical management may be beneficial, surgical ligation of the shunt is optimal. In one study, animals that receive total ligation, even if it had to be done in two or more surgeries, showed more clinical improvement than patients with incomplete shunt ligation. In general, cats do not do as well with medical therapy.

13. Describe the preoperative management of a patient with portosystemic shunt.

In animals displaying hepatic encephalopathy, it is important to correct acid-base and electrolyte disturbances before surgery. Measures to control hepatic encephalopathy also should be performed before surgery, including a low protein diet, oral lactulose, and neomycin or metronidazole. A moderately protein-restricted diet with the bulk of calories coming from carbohydrates and fat is optimal. Vegetable and dairy proteins are better tolerated than meat and egg proteins. With each patient, the protein level should be increased to the maximum tolerated. Psyllium at 1-3 teaspoons per day has been advocated to help tolerance of proteins. Some have recommended supplementation with vitamins A, B, C, E, and K. Medical stabilization for 1-2 weeks before surgery is recommended for all patients with portosystemic shunts. A preoperative coagulation screen should be performed, and crossmatched fresh whole blood should be available. Fresh frozen plasma transfusions may be necessary for hypoalbuminemic patients. Most surgeons administer a broad-spectrum antibiotic (e.g., first-generation cephalosporin) intravenously before and during surgery.

14. What considerations must be given to drug therapy and anesthetic use in patients with portosystemic shunts?

Because liver function decreases in patients with portosystemic shunts, drugs that are potentially hepatotoxic should be avoided. In addition, hepatic clearance of drugs and anesthetic agents may be delayed.

15. What parameters should be monitored postoperatively in patients with portosystemic shunts?

After surgery, many patients with portosystemic shunts are hypoglycemic, hypothermic, and hypoalbuminemic. A postoperative database should include body weight, temperature, packed cell volume, total solids, and glucose. Additional useful information is provided by electrolytes and albumin. Maintaining hydration status and perfusion with a balanced electrolyte solution is important. Mucous membrane color, capillary refill time, pulse rate and quality, and temperature should be assessed, and the patient should be monitored for seizures. In addition, serial measurement of abdominal circumference is helpful because a number of patients develop portal hypertension and ascites postoperatively.

16. What are common postsurgical complications?

Sepsis, seizures, and portal hypertension are the most critical complications that may develop postoperatively, although pancreatitis and intussusceptions have been reported. Gastrointestinal hemorrhage also may result, which can precipitate a hepatic encephalopathy crisis. Animals with seizures should be treated with appropriate measures to normalize acid-base and electrolyte balance. Sepsis should be treated aggressively.

17. What are common signs of postoperative portal hypertension?

Portal hypertension most commonly results in abdominal distention secondary to ascites. In some cases, portal hypertension is subclinical and ascites resolves in several days. Some patients develop abdominal distention, pain, and hypovolemia; others have abdominal distention with severe pain, hypovolemia, cardiovascular collapse, hemorrhagic diarrhea, and septic or endotoxic shock.

18. How should postoperative portal hypertension be treated?

If the animal develops abdominal distention with no clinical signs of pain or discomfort, continued medical therapy is indicated. Most animals with pain and abdominal distention stabilize with colloid fluid therapy. Patients with severe pain, abdominal distention, bloody diarrhea, and cardiovascular shock should be treated for shock with fluids, stabilized as much as possible, and taken for exploratory surgery to remove the ligature or thrombus that has probably developed in a partially attenuated portosystemic shunt.

19. Why may a patient with portosystemic shunt become septic postoperatively?

A patient with portosystemic shunt may develop septic peritonitis postoperatively because of bacteremia in the portal vein. The monocyte-phagocyte system in the liver may not be fully functional. Sepsis may develop as a result of inadequate filtering of portal blood by the liver before the blood reaches the systemic circulation.

20. What is hepatic microvascular dysplasia?

Hepatic microvascular dysplasia is a congenital disorder with histologic vascular abnormalities that resemble those seen in portosystemic shunts.

21. Are there breed predispositions for hepatic microvascular dysplasia?

Cairn and Yorkshire terriers are most commonly affected with hepatic microvascular dysplasia. However, many other breeds, including dachshund, poodle, Shih Tzu, Lhasa Apso, cocker spaniel, and West Highland White terrier may be affected.

22. What are the clinical signs of hepatic microvascular dysplasia?

Clinical signs are not consistently seen, but in severe cases they are quite similar to those seen with portosystemic shunts. Hyperammonemia and ammonium biurate cystalluria rarely develop. A dog may have hepatic microvascular dysplasia with elevated bile acids but be sick for another cause.

23. When should hepatic microvascular dysplasia be considered as a differential diagnosis?

Hepatic microvascular dysplasia should be considered in a patient with clinical signs consistent with a portosystemic shunt, increased bile acid concentration, and consistent liver biopsy results. Scintigraphy is consistently normal.

24. What is the treatment for hepatic microvascular dysplasia?

Treatment should not be done if the patient is subclinical. If signs of hepatic encephalopathy are present, treatment is indicated as for patients with portosystemic shunts. It is unknown at this time whether subclinical patients will develop signs of disease.

Veterinary Medicine

Avian Anesthesia: Induction of anesthesia

Injectable agents

The advantages of injectable anesthetics include ease of administration, rapid induction, low cost and availability. Disadvantages include the fact that recovery is often dependent on organ metabolism, potentially difficult reversal of medications in emergency situations, prolonged and sometimes traumatic recovery periods, muscle necrosis at injection sites and lack of adequate muscle relaxation with some medications.


This must be used intravenously, and hence a jugular, medial metatarsal or brachial vein catheter needs to be placed. It produces profound apnoea and is rarely used as an induction / anesthetic agent in birds.

Ketamine and ketamine combinations

When used alone this produces inadequate anesthesia and recoveries are often traumatic with the patient flapping wildly (doses of 20-50mg / kg are quoted). In addition, due to the high doses required, the duration of anesthesia may be prolonged, leading to poor recovery rates, potential hypothermia and hypoglycaemia.

Combining ketamine with other injectable anesthetic agents allows reduction of the dose of ketamine and can therefore allow faster recovery times. Diazepam at a dose of 1-1.5 mg / kg allows the dose of ketamine to be reduced to 20-40mg / kg or midazolam at a dose of 0.5-1.5mg / kg allows the ketamine dose to be reduced to 10mg / kg. These two tranquillisers help with muscle relaxation and sedation, and reduce flapping on recovery as well as shortening recovery time.

Ketamine may also be combined with xylazine at 1-2.2 mg / kg, which can reduce the dose of ketamine to 5-10mg / kg. Coles (1997) has recommended a dosage of 20 mg / kg ketamine and 4 mg / kg xylazine, intramuscularly, and finds that this gives 10-20 min of anesthesia with full recovery in 1-2 hours. It should however be avoided for pigeons and doves and for all wading birds.

Medetomidine (60-85 μg / kg) can reduce the ketamine dose to 5mg / kg. However slightly higher doses of 10mg / kg ketamine and 100-150 μg / kg medetomidine may be required for full surgical anesthesia in some species such as waterfowl and owls.

Again the use of these a2 agonist drugs reduces the ketamine dosage appreciably and so improves recovery rates, whilst enhancing levels of sedation and analgesia. Xylazine has the side effect of inducing respiratory and cardiac suppression and so has become less popular and should be used with caution in debilitated patients. It may be reversed with yohimbine at 0.1mg / kg intravenously, but this may not fully reverse its effects. Medetomidine may be reversed fully with atipamezole (administer the same volume as medetomidine given) and does not seem to produce such profound side effects making this a very useful injectable anesthetic. It can still have cardiopulmonary depressive effects and may compromise blood flow to the kidneys, risking renal damage. Sun conures have been noted to be particularly intolerant of ketamine-oc2 agonist combinations.

The above combined medications are generally given intramuscularly and induction will take on average 5-10 min. Complete recovery may take 2-4 hours or more unless reversible agents are used.


Midazolam (Hypnovel®) may be used on its own as a sedative in many species, and of course as an anticonvulsant. Doses for sedation in geese and swans have been quoted as 2mg / kg intramuscularly.

Extending and maintaining anesthesia

It is always advisable if using injectable anesthetics that the avian patient is intubated and oxygen supplied, or at least that it is available on standby. This also allows for anesthesia to be extended from the 15-20 minutes provided by many injectable agents, by introducing low levels of isoflurane (0.5-1%) if the surgical procedure is likely to be prolonged.

Alternatively, anesthesia may be induced directly using an inhalational agent via a face mask. It may be necessary with some species with long bills to adapt face masks from juice or water bottles, although the majority of birds of prey and parrots have short enough beaks to fit inside standard face masks.

Inhalation agents

Nitrous oxide

This has been used in avian anesthesia. It has good analgesic properties, but accumulates in large hollow viscous organs. There is some thought that it may therefore accumulate in the air sacs and may prolong anesthetic recovery times. Recent evidence disputes this as air sacs have a means of venting gases. Some species of bird have subcutaneous air pockets, such as many diving birds (e.g. gannets and pelicans), and these can rupture following gas accumulation: its use in these species should therefore be avoided.

Nitrous oxide cannot be utilised on its own for anesthesia, and halothane or isoflurane is required to allow a surgical plane to be reached. It should not exceed 50% of fresh gas flow, and should not be used for avian anesthetics if respiratory disease is suspected. As with mammalian anesthesia, the nitrous oxide supply must be discontinued some 5-10 min prior to the end of anesthesia to minimise diffusion hypoxia.


Halothane is now rarely used for birds and is not licensed for use in the UK. One of the reasons is that halothane is partly metabolised by the liver and, as many sick avian patients have some degree of hepatic function impairment, this can place the patient at some risk. Recovery is often extended and cardiac arrest often occurs at the same time as respiratory arrest giving little response time in an emergency. Halothane also depresses the responsiveness of the bird’s intrapulmonary chemoreceptors (IPCs) to carbon dioxide. This is important as the intrapulmonary chemoreceptors only respond to increasing carbon dioxide levels in the anesthetised bird and not to hypoxia. Hence birds anesthetised with halothane are less able to adjust ventilation in response to changes in carbon dioxide levels.


This is the anesthetic of choice for the avian patient. Induction may be achieved by face mask on 4-5% concentration, being turned down to 1.25-2% for maintenance, preferably delivered via an endotracheal tube. Maintaining the patient using a face mask requires an extra 25-30% increase in gas concentration.

The advantages of this agent include its low blood solubility, which allows rapid changes in anesthetic depth to occur. At sedative or light anesthesia levels the adverse cardiopulmonary effects are minimal. It does not require liver metabolism for recovery to occur. Hence it is a useful anesthetic for sick avian cases. Cardiovascular arrest tends not occur at the same time as respiratory arrest, as happens with halothane, so giving some time for resuscitation.


Sevoflurane produces a quicker recovery time than isoflurane, but otherwise seems to have the same safety margins and anesthetic effects as isoflurane, although it is not licensed for use in birds in the UK. It often requires higher induction percentages than isoflurane (5-6% compared with 3-4%). This is due to its much lower blood solubility (blood:gas coefficient 0.68), which leads to rapid recovery rates once supply of the anesthetic is discontinued. Maintenance levels average at 3%. It is minimally metabolised in the body (<1%) and, like isoflurane, seems not to produce cardiac dysrhythmias. Its high cost currently restricts its use, however its quicker revival rate may make it the gaseous anesthetic of choice for avian patients in the future.

Veterinary Medicine

Induction And Maintenance Of Anesthesia

Injectable agents

The advantages of the injectable anesthetics are that they are often easy to administer, they frequently involve minimal stress and they prevent the problems encountered with breath-holding when using gaseous induction techniques. Disadvantages include the problem of reversal for some agents, the often varying responses depending on the individual animal and the frequent respiratory depression, hypoxia and hypotensive effects they produce.


Propofol has some use in small mammals. It may be used in most species of small mammal at a dose of 10 mg / kg after the use of a pre-anesthetic medication such as acepromazine, but requires intravenous or intraosseous access. Its effects in rabbits are broadly similar to those seen in cats and dogs although breath-holding is very common and may cause problems where intubation is not possible.



Ketamine may be used alone for chemical restraint in ferrets at doses of 10-20mg / kg but, as with cats and dogs, the muscle relaxation is poor and salivation occurs. More often ketamine is combined with other drugs such as the alpha-2 agonists, xylazine and medetomidine. In ferrets 10-30 mg / kg ketamine may be used with 1-2 mg / kg xylazine, preferably giving the xylazine 5-10 min before the ketamine.

Ketamine has been used with medetomidine and butorphanol in ferrets in the same manner as so-called ‘triple anesthesia’ in cats with dosages of 5 mg / kg (ketamine), 0.08 mg / kg (medetomidine) and 0.4 mg / kg (butorphanol). Reversal is achieved with atipamezole at 1 mg / kg.


In rabbits ketamine has a very rapid renal excretion and so is short acting. It may be used at a dose of 20-35 mg / kg in conjunction with medetomidine at 0.3-0.5 mg / kg or with xylazine at 5 mg / kg using lower doses for debilitated animals. The advantages are a quick and stress-free anesthetic, but the combination will cause ‘blueing’ of the membranes and make detection of hypoxia difficult. True cyanosis also occurs. Respiratory depression during longer procedures may become a problem and intubation is often advised. If used, medetomidine may be reversed using atipamezole at 1 mg / kg.

It should be noted that multiple anesthetics using a combination of ketamine and xylazine have resulted in myocardial necrosis in rabbits.

A triple combination can also be used in rabbits with 0.2 mg / kg medetomidine, 10 mg / kg ketamine and 0.5 mg / kg butorphanol given subcutaneously which will provide approximately 20min anesthesia. Alternatively, the intravenous route may be used when a lower dose of the three drugs is required: 0.05 mg / kg medetomidine, 5 mg / kg ketamine and 0.5 mg / kg butorphanol. This gives a more rapid induction, but shorter duration of anesthesia, although it may be used as an induction combination prior to intubation and maintenance on gaseous anesthesia. Again reversal with atipamezole will hasten recovery.

Muridae / Cricetidae

Ketamine can be used at 90 mg / kg in combination with xylazine at 5 mg / kg intramuscularly or intraperitoneally in rats, with mice and hamsters requiring 100-150 mg / kg of ketamine and 5 mg / kg xylazine. These combinations provide 30min or so of anesthesia. In gerbils the dose of xylazine may be reduced to 2-3 mg / kg as they appear more sensitive to the hypotensive effects of the alpha-2 agonists drugs, with ketamine doses reduced to 50 mg / kg.

Ketamine may also be used in combination with medetomidine at doses of 0.5 mg / kg in these species. The advantages of the alpha-2 agonists are that they produce good analgesia (which ketamine does not) and that they may be quickly reversed with atipamezole at 1 mg / kg. Their disadvantages include their severe hypotensive effects, and that once administered any injectable anesthetic is always more difficult to control than a gaseous one. They also increase diuresis and may exacerbate renal and, of course, cardiovascular dysfunction.


Ketamine at 20-40 mg / kg may be used in conjunction with xylazine at 3-5 mg / kg in guinea pigs and chinchillas to produce a light plane of anesthesia. Ketamine at 40mg / kg may also be used with medetomidine at 0.5mg / kg for guinea pigs, or ketamine at 30 mg / kg with medetomidine at 0.3 mg / kg for chinchillas. Reversal with 1 mg / kg atipamezole may be performed. Both of these may be improved after an acepromazine pre-anesthetic medication of 0.25 mg / kg.

Alternatively for chinchillas a ketamine (40 mg / kg) and acepromazine (0.5 mg / kg) combination can be used. Induction with these drugs takes 5-10 min and typically lasts for 45-60 min, but recovery may take 2-5 hours. Reducing the dose of acepromazine / ketamine and using a reversible alpha-2 agonist in addition may be beneficial, but should be weighed against the greater hypotensive effects of the alpha-2 agonist drugs.

This author prefers a combination of acepromazine, medetomidine and ketamine for minor procedures such as radiography, dental work and straightforward extractions in chinchillas. Acepromazine is given subcutaneously (0.1-0.2 mg / kg) 10-15 min before administering a combination of 1-5 mg / kg ketamine and 0.01-0.05 mg / kg medetomidine, using the lower end of the range for very debilitated patients.

Fentanyl / fluanisone (Hypnorm®)

This drug combination is a neuroleptanalgesic licensed for use in rats, mice, rabbits and guinea pigs in the UK. Fentanyl is an opioid derivative and fluanisone is a neuroleptic.


Fentanyl / fluanisone may be used as sedation only on its own at a dose of 0.5 ml / kg intramuscularly. This produces sedation and immobilisation for 30-60 min according to the data sheets, but its analgesic effect due to the opioid derivative fentanyl will persist for some time after. It may be reversed with 0.5mg / kg butorphanol intravenously or 0.05mg / kg buprenorphine, both of which will counteract the fentanyl and its analgesia and substitute their own pain relief.

Alternatively, to provide greater anesthetic depth, fentanyl / fluanisone may be combined with diazepam (0.3 ml Hypnorm® to 2mg / kg diazepam) and administered intraperitoneally or intravenously (but drawn up in separate syringes as they do not mix), or with midazolam (0.3 ml Hypnorm® to 2mg / kg midazolam) administered intramuscularly or intraperitoneally in the same syringe. Hypnorm® also may be given intramuscularly followed 15min later by midazolam intravenously into the lateral ear vein. These two combinations provide good analgesia and muscle relaxation with a duration of anesthesia of 20-40 min. Again fentanyl may be reversed with buprenorphine or butorphanol given intravenously, or in emergencies naloxone at 0.1 mg / kg intramuscularly or intravenously may be used, but this provides no substitute analgesia.

Fentanyl / fluanisone combinations are well tolerated in most rabbits, but they can produce respiratory depression and hypoxia, which can lead to cardiac arrhythmias and even arrest.


Fentanyl / fluanisone may be used as sedation only on its own at a dose of 0.01ml / 30g body weight in mice and 0.4 ml / kg in rats. Again this produces sedation and immobilisation for 30-60 min and may be reversed with buprenorphine or butorphanol as above.

Alternatively it may be combined with diazepam (mice 0.01ml / 30g Hypnorm® with 5mg / kg diazepam intraperitoneally; rats 0.3 ml / kg Hypnorm® with 2.5 mg / kg diazepam intraperitoneally) where the diazepam and Hypnorm® are drawn up in separate syringes as they do not mix, or with midazolam. Midazolam is miscible with Hypnorm® and for rodents the recommendation is that each drug is mixed with an equal volume of sterile water first and then mixed together. Of this stock solution, mice receive 10ml / kg and rats 2.7ml / kg as a single intraperitoneal injection. These two combinations provide anesthesia for a period of 20-40 min.


Hypnorm® may be used for sedation only on its own at a dose of 1 ml / kg intramuscularly. This may be problematic in guinea pigs as large volumes are required and Hypnorm® is an irritant and may cause lameness when the whole dose is placed in one spot: multiple sites are therefore preferred. Alternatively it may be combined as above with diazepam (1 ml / kg Hypnorm® and 2.5 mg / kg diazepam each drawn up in separate syringes) and administered intraperitoneally, or with midazolam by making the stock solution as described for Muridae, and then administering 8 ml / kg of this solution intraperitoneally. Hypnorm® may be reversed with the partial opioid agonists buprenorphine and butorphanol or with the full antagonist naloxone.

Volatile agents

Volatile agents have the advantage over injectable agents in that it is easier to alter the depth of the anesthetic quickly. The recovery times are often much shorter than with injectable anesthetics and frequently their side effects are less, particularly with isoflurane and sevoflurane. However, their disadvantages include the fact that there is a drying effect on the airways of the patient when using inhalational anesthetic agents that can cause dehydration during long procedures. They also create problems if used as an induction agent, as many species will breath-hold during this procedure.


This has been used in all small mammals, however its margin of safety is less than that of isoflurane. Induction concentrations should not exceed 3% and anesthesia can be maintained with 1.5%. Disadvantages include possible cardiac arrhythmias, particularly in lagomorphs who are some of the main culprits of breath-holding. These may lead to apnoea and cardiac arrest. Use of halothane is best after pre-anesthetic medication with acepromazine rather than Hypnorm® as the latter agent requires extensive hepatic metabolism as does halothane.


This is now becoming the most widespread volatile agent used for induction and maintenance of general anesthesia in small mammals as well as in dogs and cats. A pre-anesthetic medication incorporating an analgesic is usually administered as isoflurane has no post-anesthetic analgesic qualities and it is irritant to the mucous membranes of many animals. Inspired concentrations required for induction of anesthesia vary from 2.5-4%. Breath-holding still occurs, but the practice of supplying 100% oxygen to the patient for 2min prior to anesthetic administration helps minimise hypoxia. After this pre-oxygenation, gradually introduce the isoflurane, first 0.5% for 2min, then assuming regular breathing, increase to 1% for 2min and so on until anesthetic levels are reached allowing a smooth induction. Surgical anesthesia can usually be maintained at 1.5-2.5% assuming adequate analgesia; the MAC for this agent is 2.05% in rabbits. One of the main advantages of isoflurane is its improved safety profile in debilitated patient as <0.3% of the gas is metabolised hepatically, the rest merely being exhaled for recovery to occur. Recovery following use of this agent is rapid.


This is not currently licensed for use in small mammals in the UK unlike isoflurane, but its wide safety margins and the fact that it appears less noxious to small mammals and so induces less breath-holding when used as an induction agent, have made it an ideal anesthetic choice. Induction with sevoflurane still causes some breath-holding in rabbits and guinea pigs if used at 8%, therefore this author prefers to use an induction level of 4%. Inspired concentrations somewhere around 2-3% in 100% oxygen are required to maintain anesthesia: sevoflurane has an MAC of 3.7% in rabbits.

Aspects of maintaining gaseous anesthesia

As with all gaseous anesthetics, placing an endotracheal tube after induction of anesthesia is recommended whenever possible. This is relatively straightforward in rabbits using a number 1 Wisconsin flat bladed paediatric laryngoscope and a 2-3 mm tube. The aid of a rabbit mouth gag helps visibility, however the procedure becomes a specialised one for rodents such as rats and mice where rigid guide tubes / wires and smaller scopes are used to guide the tube into the larynx. In these species therefore, face masks are more commonly used. In rabbits and ferrets the use of lidocaine spray on the larynx helps to reduce laryngospasm and aids intubation. Ferret intubation is straightforward and resembles the procedure for a cat.

Intermittent positive pressure ventilation

This may be necessary in some individuals who breath-hold during induction. If intubation is not possible then three options are available.

(1) Ensure a tight-fitting face mask and have an Ayres-T piece / Mapleson C / modified Bain circuit with half litre bag attached which can be used to attempt ventilation.

(2) Place a nasopharyngeal tube through the medial meatus of the nose, into the pharyngeal area and insufflate oxygen (41 / min is required to combat the resistance of the small diameter tubing).

(3) Perform an emergency tracheotomy with a 25 / 27 gauge needle attached to the oxygen outlet.

Anaesthetic breathing systems

Most of the small mammals described here are <2kg in weight. For this reason an Ayres T-piece, a modified Bain or Mapleson C circuit are the best anesthetic systems to use to minimise dead space. For larger rabbits, an Ayres T-piece is usually sufficient.

Veterinary Drugs

Induction of Anesthesia

General anesthesia may be induced by the use of inhalational agents or, more commonly, by the use of injectable drugs. The latter are usually administered by the intravenous route, but some will also work when given intramuscularly, e.g. ketamine. Other routes may occasionally be used (e.g. rectal, intraperitoneal, transmucosal) but, in common with all other extravenous routes, they prevent titration of the induction agent to a suitable endpoint. That is to say, with intravenous induction techniques, the drug can be given slowly to effect and administration stopped when the patient is at a suitable depth of anesthesia for endotracheal intubation: drugs given by other routes are administered based solely on body weight, implying that some animals will receive a relative overdose while others will be underdosed, as not all patients of a particular weight will require the same dose of anesthetic.

Induction agents are often chosen on the basis of anticipated recovery time, effects on the cardiovascular system and so on, but in many cases several agents may be suitable for a particular case, and personal preference and expense play a role.


A number of derivatives of barbituric acid have previously been used for induction and/or maintenance of general anesthesia (methohexital, pentobarbital) in veterinary patients. However, the only barbiturate currently in use as an anesthetic induction agent is thiopental, although newer induction agents have largely superseded it.






Doses of injectable anesthetic agents

Depending on the agents chosen, premedication may dramatically alter the dose of injectable anesthetic agent required (see Table Suggested drug doses for patients following ‘standard premedication’ with acepromazine ± opioid). For example, acepromazine will reduce the dose of thiopental for induction of anesthesia by about 30-50%, while medetomidine will decrease it by anything up to 90%. In addition, the state of the patient’s health will also have a major bearing on the quantity of induction agent required. It has been said that: ‘The dose of induction agent required is as much as the patient needs, and no more’, i.e. anesthetic agents should be given to effect. Thus, it is not possible to be completely prescriptive about induction doses, and the information provided in Table Suggested drug doses for patients following ‘standard premedication’ with acepromazine ± opioid should be considered only as a guideline.

Table Suggested drug doses for patients following ‘standard premedication’ with acepromazine ± opioid. Much lower doses will generally be required following medetomidine.

Drug Dogs Cats Comments
Thiopental 7-10 mg/kg intravenous 7-10 mg/kg intravenous
Alfaxalone 2mg/kg intravenous 2-5 mg/kg intravenous
Propofol 3-4 mg/kg intravenous 4-6 mg/kg intravenous
Etomidate 0.5-2 mg/kg intravenous 0.5-2 mg/kg intravenous No veterinary licence
Ketamine 0.2 mg/kg diazepam + 5 mg/kg ketamine intravenous 0.2 mg/kg midazolam + 5-10 mg/kg Diazepam-ketamine combination useful for examination of laryngeal function. The two drugs can be administered in the same syringe, but should be mixed just before administration 

Midazolam-ketamine can be mixed in the same syringe, and provide deep sedation in cats

Medetomidine-ketamine can be mixed in the same syringe, with potential for reversal with atipamezole (ketamine dose is so low in this combination that once medetomidine is reversed, ketamine exerts little effect). This combination is usually used without prior premedication

ketamine intramuscular; 80 μg/ kg medetomidine + 2.5-7.5 mg/kg ketamine intramuscular