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


Bronchoalveolar Lavage

Today the use of fiberoptic bronchoscopy is a common and standard diagnostic procedure, which allows direct observation of the upper and lower conducting airways. During passage of the endoscope through the nasopharynx, trachea, and large bronchi, the quantity of mucous secretions can be assessed readily in addition to the degree of mucosal edema and bronchospasm. In addition to examination of the airway lumen, one of the greatest advantages and rewards from bronchoscopy is the ability to sample the large and small airways and the alveoli. The specimens collected are then analyzed for their cellular and noncellular constituents.

In recent years, bronchoalveolar lavage (bronchoalveolar lavage) using either an endoscope or specialized tubing has gained some popularity over more traditional sampling methods such as tracheal aspiration for most cases in which a diffuse inflammatory disorder is suspected. For many years, it has been assumed that sampling the lower trachea provides a representative impression of the alveoli and small airways because airway free cells from the peripheral lung eventually were swept toward the trachea for clearance.

However, a large clinical case survey of young athletic horses presented with poor performance attributable to the lower respiratory system has shown that the cytologic and bacteriologic results are correlated poorly between samples obtained from the tracheal aspirate versus those from bronchoalveolar lavage. The study demonstrated that tracheal aspirate and bronchoalveolar lavage cytologic cell differential counts differed greatly within the same horse, which suggests that samples from the tracheal puddle may not reflect accurately the population of cells and secretions present within the small airways and alveoli. This is relevant insofar as exercise intolerance, airway injury resulting from inflammation, and airway hyperreactivity are associated with disease in the small airways, reflected best by bronchoalveolar lavage cytology. In addition, a higher rate of positive bacterial cultures was obtained from tracheal aspirate samples versus bronchoalveolar lavage samples performed on the same occasion. Thus the lower trachea apparently harbors a normal bacterial flora that may not be present within the small airways and alveoli. For these reasons, bronchoalveolar lavage is becoming a more popular tool to assess distal (small) airway inflammation rather than the tracheal aspirate method of sampling.

To validate the relevance of bronchoalveolar lavage differential cell counts as a complementary diagnostic tool in the assessment of the respiratory system, other quantitative measurements are necessary beyond the routine clinical examination. In the last two decades, the syndrome of heaves has been studied extensively, and several research laboratories throughout the world have clearly demonstrated a high correlation between the bronchoalveolar lavage cell differential and results of pulmonary function testing and histamine bronchoprovocation in heaves-affected horses. In recent years, similarly characterized lung function in young athletic horses with noninfectious inflammatory airway disease (IAD) has paralleled these findings with respect to the diagnostic usefulness of bronchoalveolar lavage.

The purpose of this chapter is to discuss the use of the bronchoalveolar lavage technique as a tool to identify and characterize pulmonary inflammation in horses that suffer from diffuse lung pathology such as inflammatory airway disease in the young athletic horse and the heaves syndrome in mature horses. In addition viral and bacterial pulmonary conditions are discussed briefly with respect to their diagnosis by bronchoalveolar lavage.

Indications For Bronchoalveolar Lavage

Bronchoalveolar Lavage Procedure

bronchoalveolar lavage can be performed on most conscious horses with mild sedation (xylazine 0.3-0.5 mg/kg IV or romifidine 0.03-0.05 mg/kg IV) and airway desensitization by a local anesthetic (lidocaine solution 0.4% w/v, without epinephrine). The procedure can be conducted using either a bronchoscope 1.8 to 2 m in length or a specialized bronchoalveolar lavage tube (Bivona Medical Technologies, Gary, Ind.). Once the bronchoscope or bronchoalveolar lavage tube is in the trachea, reaching the tracheal bifurcation (carina) usually induces coughing. Infusing 60 to 100 ml of prewarmed lidocaine solution (0.4%, without epinephrine) is therefore beneficial at this point to desensitize cough receptors located at the carina. After this infusion step the endoscope or bronchoalveolar lavage tube is gently but securely wedged, as detected by resistance to further advancement. Prewarmed sterile saline (200-300 ml) is infused rapidly into the lung and is subsequently aspirated.

The total amount of saline should be divided into two separate boluses for infusion, with attempts to retrieve as much fluid as possible between each bolus. In general, retrieval of 40% to 60% of the total amount of infusate indicates a satisfactory bronchoalveolar lavage. In horses with advanced disease, lower volumes are recovered and a tendency exists for less foam (surfactant) to be present. The bronchoalveolar lavage fluid samples are then pooled and kept on ice if processing is not possible within 1 hour after collection. Gross examination of the fluid should be performed to detect any flocculent debris or discoloration. One or two serum or ethylenediaminetetraacetic acid (EDTA) tubes of bronchoalveolar lavage fluid are centrifuged (1500 X g for 10 min) and air-dried smears are made from the sample pellet after removal of the supernatant. In preparation of the smears, the slides must be air dried rapidly using a small bench-top fan to preserve good cellular morphology. Smears thus prepared can be kept at room temperature for up to 8 to 10 months with little cellular alterations. The air-dried smears can be stained with Diff-Quik, Wright-Giemsa, May Gruenwald, Leishman’s, or Gram’s stain for cellular and noncellular constituent interpretation. The cellular profile and morphology may serve as a guide to the nature of airway injury, inflammation, and the pulmonary immunologic response to infections or foreign antigens.

Differential Cell Counts And Their Interpretation


bronchoalveolar lavage is undoubtedly becoming a powerful ancillary diagnostic tool to assist in the diagnosis of clinical and sub-clinical lower airway respiratory conditions such as non-infectious inflammatory airway disease in the young athletic horse and recurrent airway obstruction, or heaves, in older horses. Using recognized, standardized procedures, the bronchoalveolar lavage differential cell count is fairly consistent for normal horses and any alteration in the cytologic profiles from normal values identifies a wide range of nonseptic inflammatory processes. Although at present, clinicians are recommending specific treatment according to cytologic findings of the bronchoalveolar lavage cell differential, a more in-depth knowledge of the various disorders in the future may allow equine practitioners to provide more accurate prognostic information to members of the horse industry with respect to respiratory diseases in athletic horses. More so, the majority of young and mature athletic horses with an excess amount of white mucopus within the airways and markedly elevated neutrophil percentage on the cell differential do not represent a septic process. Rather, these cases demonstrate nonseptic inflammatory airway disease.


Tracheal Aspirates: Technique

Several methods for obtaining TAs have been developed, each having advantages and disadvantages. The most important consideration when choosing a technique is whether microbiologic culture of the tracheobronchial secretions is indicated. In general, aspirates obtained endoscopically are unsuitable for this use because they invariably become contaminated by upper airway flora. However, a guarded catheter passed through the endoscope may be used to obtain samples suitable for microbiologic culture. Alternatively, the transtracheal (percutaneous) aspiration technique may be employed.

The choice of technique can affect significantly the numbers and types of cells obtained. Thus standardization of procedures with regard to type of technique, time of sampling, volume and type of fluid instilled, sample handling, and processing is recommended for meaningful interpretation and comparison of results.

Transtracheal Aspirates

The rationale for use of transtracheal aspiration is based on the assumption that the bacterial population derived from the upper airway of normal horses is negligible beyond the proximal trachea. Therefore organisms cultivated from a tracheal aspirates represent bacteria found in the distal trachea and lower airways. These bacteria may be present transiently, or they may be part of a pathologic process. The distinction between these phenomena is important.

Samples obtained by transtracheal aspiration are suitable for cytologic and Gram’s stain evaluation and bacteriologic or fungal cultivation. However, this technique is invasive, and possible complications have tended to preclude its widespread application. These include subcutaneous abscessation at the puncture site, tracheal laceration and hemorrhage, chondritis, and pneumomediastinum. In addition, the catheter may break off in the tracheal lumen, although in most cases it is coughed up rapidly and swallowed. Good technique prevents most untoward complications.

A variety of needle-catheter combinations may be used, but maintaining asepsis is critical. The components may be purchased either individually or prepackaged and include an introducer catheter-over-needle, flushing catheter, and a stylet (Catheter TW 1228 and 1628, Mila International, Phoenix, Ariz.). A convenient combination of catheters comprises a 12-gauge needle, 3-inch over-the-needle cannula, and number 5 French canine urinary catheter with the tip cut off obliquely.

Sedation generally is indicated when performing a tracheal aspiration, with xylazine (Rompun) used commonly. An area measuring approximately 6 by 6 cm over the middle third of the cervical trachea should be clipped and prepared for aseptic surgery. A bleb of local anaesthetic (approximately 1 ml) is injected subcutaneously over the midline and a stab incision is made through the skin and subcutaneous tissue with a number 15-scalpel blade. The trachea is stabilized with one hand and the cannula is introduced into the tracheal lumen between two cartilage rings. The stylet is removed, and the urinary catheter is passed down into the tracheal lumen to the level of the thoracic inlet, where the washing and aspiration is performed. In most cases 10 to 15 ml of sterile isotonic saline is adequate to obtain a satisfactory sample. However, repeated infusions may be required. Once an adequate sample has been collected, the catheter should be withdrawn, maintaining the cannula in situ during retraction to minimize contamination of peritracheal tissues.

Tracheal Aspirates: Endoscopic Technique Using Unguarded Catheters

An increasingly popular and well-tolerated alternative for collection of TAs is via a fiberoptic endoscope. However, samples collected using unguarded catheters are contaminated with nasopharyngeal bacteria and are unsuitable for microbial cultivation. Endoscopy allows visualization of the respiratory tract at the time of sampling, where evaluation of the mucosa of the trachea (degree of hyperemia) and its luminal contents (quantity and quality of mucus, mucopurulent secretions, and blood) may assist in interpretation of cytologic results. Furthermore, if the length of the endoscope permits, the large bronchi may be examined, and purulent debris draining from a specific bronchus suggestive of pulmonary abscess occasionally may be recognized.

A small polyethylene catheter is passed through the biopsy channel of the endoscope and 10 to 15 ml of sterile, isotonic saline instilled. Most horses have a ventrally depressed area in the trachea, anterior to the carina and level with the thoracic inlet. Fluid accumulates at this site and forms a puddle from where it can be aspirated. The principal use of samples collected using this technique is for cytologic examination.

Tracheal Aspirates: Endoscopic Technique Using Guarded Catheters

Recently, guarded systems have been evaluated for collection of uncontaminated samples from the lower airways via endoscopy. In adult horses, the advantages of collection using guarded catheters include noninvasiveness, speed with which samples can be obtained, visual inspection of the airways, and guidance of the catheter. These advantages generally outweigh those of the transtracheal method, which include reduced chance of bacterial or cellular contamination from the upper respiratory tract.

Several multilumen, telescoping, plugged catheters have been assessed. One is the endoscopic microbiology aspiration catheter (Catheter EMAC800, Mila International, Phoenix, Ariz.). This catheter contains a glycol plug in the outer catheter, to maintain sterility as the catheter is being advanced through the endoscope and trachea, and an inner catheter for retrieval of the sterile specimen. Another system involves a 5 French inner catheter within, an 8 French guiding catheter (Catheter V-EBAL-8.0-190, Cook Veterinary Products, Bloomington, Ind.). Before each sample collection the endoscope and its biopsy channel must be disinfected. Glutaraldehyde (Cidex) is an appropriate disinfectant.

Some controversy remains regarding the adequacy of samples collected through guarded catheters for microbiologic cultivation. Technical prowess definitely influences the quality of sample obtained. Factors that help prevent contamination include rapid collection of the sample, limited volume of infusate (10-15 ml), and advancement of only the inner catheter into the tracheal puddle rather than the catheter as a whole. In addition, if the horse has coughed frequently during the procedure, an increased risk of contamination is likely, and these samples are rarely appropriate for bacteriologic cultivation.


Laser Surgery of the Upper Respiratory Tract

Lasers have become a common instrument for surgical and nonsurgical therapy in equine medicine. The many different tissue interactions that can be produced, the precision of its use, and the ability to apply laser energy to less accessible areas are the great advantages of the laser compared with other forms of therapy.

Laser is an acronym for Zight amplification of stimulated emission of radiation. The light emitted by lasers works according to the basic properties of light and electromagnetic radiation, but it is very different from the light produced by more common light sources such as incandescent bulbs, fluorescent lamps, or sunlight. The similarity between laser light and common white light is that all light consists of particles (photons) that travel through space in unique waveforms. White light consists of a mixture of many different wavelengths. Each color of visible light has its own characteristic wavelength. Visible light has an electromagnetic spectrum of wavelengths that range from approximately 400 nm to 700 nm.

Laser light can be within the visible spectrum of light, but it differs significantly from white light because of its monochromacity, collimation, and coherence. Laser light consists of a single wavelength or an extremely narrow range of wavelengths, and is therefore considered “monochromatic.” Also light emitted from bulbs or headlights diverge rapidly, but laser light has a very narrow cone of divergence. Finally, light waves can travel through space without any fixed relationship to each other, meaning they are incoherent. If all waves are lined up together so their peaks and valleys match, they are in phase, or coherent. Laser light is coherent, and white light is not.

The components to create laser light are an active medium, a power source, an optical resonator, and an output coupler (partially transmitting mirror). The active medium is the material that determines the wavelength of the laser. The medium can be a gas, a liquid, a solid material, or a junction between two plates of semiconductor materials. The power source is the pump that stimulates the emission of radiation and the type of energy used as a power source is determined by the lasing medium. The optical resonator can be thought of as mirrors on either side of the medium that reflects the light back into the medium for “amplification.” The output coupler allows a portion of the laser light contained between the two mirrors to leave the laser resonator in the form of a beam.

Lasers are characterized in two main ways. They can be delineated by the medium (diode, CO2, neodymium: yttrium-aluminum-garnet [Nd:YAG]) or the power output (pulsed vs continuous). A general classification system also exists for laser power and safety (classes I-IV). Classes I and II are low-risk lasers with a power of less than 1.0 mW. “Cold” or therapeutic lasers are class III lasers. All surgical lasers are class IV (>0.5 watts). Although the power is measured in watts, the power density is termed “irradiance” and is the amount of power per unit of surface area. Irradiance is equal to the laser power output/laser beam size (W/cm2). Therefore a larger beam size of a given power will have a smaller irradiance. The number of joules depicts the total energy, which is equal to the laser output (watts) multiplied by the exposure time (seconds). The “energy fluence” is equal to joules/laser beam size, and measures the total amount of energy directed to the tissue during a treatment. An understanding of this fact is important because the effectiveness of a particular laser is determined not only by its wavelength but also by how it is used.

Laser light interacts with tissue in several ways. It can be absorbed, transmitted, reflected, or scattered. The percentage of each interaction is dependent on the characteristics of the tissue and the laser light. The amount of absorption is dependent on the wavelength of the light relative to the chromophore content of the tissue (hemoglobin, keratin, protein, water, melanin). Each chromophore has its own absorption spectrum for different wavelengths of light. If the light is absorbed it is transformed into heat energy. Heating tissue to 60° C will lead to coagulation of proteins, and heating tissue to higher than 100° C will result in vaporization. Thus lasers will yield different biologic effects dependent on the energy absorption coefficient. Although vaporization and coagulation can be seen at the time of surgery, a zone of thermal injury exists beyond what can be seen at surgery. If a large amount of energy is expended that is not strictly focused on the area of interest, excessive swelling and trauma to the tissues may occur postoperatively.

Therapeutic Lasers

Lasers have become a common tool to speed healing in many different types of injuries. The lasers used for this purpose differ greatly from surgical lasers. Therapeutic lasers are considered “cold” or low-power lasers and fall into classes II and III. They may induce some heat but no greater than that which would be felt from a 60-W bulb held close to the skin. The benefits of these lasers are the analgesic effects caused by alterations in nerve conduction and wound healing caused by stimulation of changes in intracellular calcium that ultimately results in increased protein synthesis and collagen production. The most common lasers employed are the gallium arsenide (GaAs) and helium neon (HeNe) lasers at a distance of 1 to 2 mm from the surface of the target tissue for a total energy density of 5 J/cm2.

Surgical Lasers

Although surgical lasers have existed since 1960, it was not until lasers could be applied through small flexible fibers that these tools had an enormous impact on equine surgery. These fibers can be passed down the biopsy channel of a videoendoscope and employed under videoendoscopic control. This development revolutionized the treatment of upper respiratory conditions by providing the surgeon an opportunity to approach lesions within the nasal cavity, larynx, and pharynx without making a surgical skin incision. This new procedure also provided a technique for cutting a fairly reactive and very wellvascularized tissue that is precise and provides significant hemostasis.

The two most common lasers used for upper respiratory surgery are the diode and Nd:YAG lasers. They have wavelengths of 980 nm and 1064 nm, respectively, and can pass down a small flexible optical quartz fiber without significant disruption of wavelength. The diode laser has two main advantages compared with the Nd:YAG. The diode laser is a much smaller unit (less than 15 lb) and is significantly less expensive than the Nd:YAG. The major disadvantage of the diode laser is its power limitation of 25 W, whereas the Nd:YAG can exceed 50 W. Other lasers such as the CO2 cannot pass down a small fiber effectively because of their much larger wavelength and therefore cannot be used with a standard videoendoscope. Although the CO2 laser wavelength is strongly absorbed by water and therefore is an excellent precise cutter, it has only poor-to-fair coagulating capability. The diode or Nd:YAG wavelengths are diffusely absorbed by all protein molecules and therefore have greater coagulation capabilities, although they do not cut as well as the CO2 laser.

The laser can be used in contact or noncontact mode. Most surgeries can be performed with a bare fiber (no special tip) in contact mode. This method provides very accurate, controlled cutting and hemostasis of the small vessels in the respiratory mucosa, and provides the surgeon some tactile sense of the procedure. A lower power setting of 14 to 18 W is sufficient in most cases. This also means that a small very portable diode laser can be employed. If the laser is used correctly, little lateral thermal damage should occur. The surgeon resects the tissue by dragging the fiber across the tissue as he or she would lightly drag a scalpel blade. The types of surgeries commonly done in this fashion include axial division of epiglottic entrapment, resection of subepiglottic or pharyngeal cysts, vocal cord resections, resection of granulation tissue, and treatment of guttural pouch tympanites.

With noncontact laser surgery, the fiber is held 3 to 5 mm away from the target tissue. A higher power setting of 40 to 60 W is commonly required to work effectively, which requires an Nd:YAG laser. Noncontact surgery is used mostly for ablation of cystlike structures such as ethmoid hematomas or pharyngeal cysts and to vaporize membranous structures.

General Use

A great advantage of the use of lasers in respiratory surgery is that many of the surgeries can be done on the standing, sedated animal on an outpatient basis. This fact also equates to a shorter, easier postoperative management because no skin incision is present. Procedures can be performed with the animal standing in the stocks with just intravenous sedation such as xylazine (0.44 mg/kg). Repeated half doses or a longer-acting agent may be required depending on the procedure and the experience of the surgeon. With the horse sedated, a twitch is normally not required to pass the endoscope. The horse’s head can be suspended from cross ties for support, but an individual must always be positioned at the horse’s head for safety and to alter the head position as needed. Topical anesthetic is applied to the area of interest through polyethylene tubing that is advanced down the biopsy channel of the endoscope. The horse often swallows while the anesthetic is applied, and application should be intermittently suspended to make certain the anesthetic is applied appropriately in between swallows. The anesthetic is usually effective for approximately 2 hours, so the animal is not allowed to eat for 1 to 2 hours after surgery.

Laser safety should always be considered. Although the laser is used within the respiratory cavity, surgical personnel should still wear laser safety glasses as a precaution against any misfiring of the laser. The laser should always be kept in the standby mode when not being used. If a procedure is performed with the horse under general anesthesia near an endotracheal tube, the oxygen concentration should be decreased with helium to dramatically reduce the risk of spontaneous ignition. Smoke evacuation is usually not necessary in contact laser surgery in the standing horse but may be required in noncontact work or when the horse is under general anesthesia.

Antiinflammatory medication is the cornerstone of postoperative management in the upper respiratory tract. Phenylbutazone (4.4 mg/kg) and dexamethasone (0.044 mg/kg) are given in the immediate postoperative period. Both medications are recommended for several days at a decreased dose depending on the type of surgery and anticipated degree of inflammation. Local antiinflammatory medication can also be administered through a 10-Fr catheter that is advanced through the nasal passage into the nasopharynx. Ten milliliters of a mixture of dimethyl sulfoxide, glycerine, and dexamethasone solution are administered slowly through the catheter while watching the horse swallow. This mixture is administered twice daily for as long as 7 days.

Antimicrobials are not commonly given unless the surgeon is working on areas of thickened scar tissue where the vascularity may be compromised, or extensive use of the laser is required. Although vaporization of all tissue with the laser results in a sterile incision, the adjacent tissues of the throat and mouth can easily contaminate the open wound bed at the conclusion of surgery. Surgical inexperience can lead to greater thermal injury than visually appreciated and increased susceptibility to infection even in healthy tissues, particularly when the laser is used on subepiglottic tissues.

Laser Surgery of the Upper Respiratory Tract: Common Procedures


Although the laser has become an invaluable tool for many upper respiratory surgeries, its improper use can create significant trauma and irreparable damage. Great care should be taken to use only as much energy as necessary to complete the task and minimize extraneous firing. When used appropriately, the laser greatly diminishes the need for more extensive surgery and speeds the recovery of the patient.


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.


Axial Deviation of the Aryepiglottic Folds

Axial deviation of the aryepiglottic folds () has been recognized as a cause of dynamic upper respiratory obstruction in horses since the first use of high-speed treadmill exercise testing to evaluate poor performance. The membranous portions of the aryepiglottic folds, which extend from the abaxial margin of the epiglottis to the corniculate processes at the lateral aspect of the arytenoid cartilages, collapse axially to occlude the glottis during inspiration (). Horses with axial deviation of the aryepiglottic folds have poor performance and are often reported to “finish poorly” or “stop” near the end of a race. During inspiration at exercise, some affected horses make an abnormal noise that may sound similar to the “roar” associated with laryngeal hemiplegia. The cause is unknown, although immaturity may be a factor in younger horses and should be suspected if concurrent dynamic upper respiratory abnormalities are present.

Clinical Signs And Diagnosis

Affected horses are typically presented with a chief complaint from the owner of poor performance. Horses with axial deviation of the aryepiglottic folds may or may not make an abnormal upper respiratory noise during exercise. No breed or gender predisposition exists, and the condition has been diagnosed in Thoroughbreds, Standardbreds, and racing Arabians. The condition has been reported in racehorses from 2 to 8 years of age, but the percentage of 2- and 3-year-old horses that were diagnosed with axial deviation of the aryepiglottic folds in one hospital population was significantly greater than in the overall hospital population evaluated for poor performance.

Physical examination and endoscopic examination of the resting horse typically do not yield any abnormalities related to the condition. At endoscopic examination at rest, the membranous portion of the aryepiglottic folds of affected horses has no visible structural or functional abnormalities. Nasal occlusion during endoscopic examination, which mimics airway pressures generated during exercise, does not induce axial deviation of the aryepiglottic folds in horses that subsequently demonstrate the condition during treadmill exercise. Endoscopic examination during high-speed treadmill exercise is required to diagnose axial deviation of the aryepiglottic folds. axial deviation of the aryepiglottic folds most often occurs as a distinct clinical problem but also can occur with other upper airway abnormalities. Horses may be unilaterally or bilaterally affected. No association has been identified between the development of axial deviation of the aryepiglottic folds and subsequent dorsal displacement of the soft palate or other causes of dynamic upper respiratory abnormalities.

Severity of axial deviation of the aryepiglottic folds is evaluated based on the extent to which the membranous portion of the aryepiglottic folds collapse across adjacent structures of the larynx. With mild collapse, the fold remains abaxial to the vocal fold. Moderate cases have collapse of the fold beyond the vocal fold but less than halfway between the vocal fold and the midline. In severe collapse, the fold reaches or crosses the midline of the glottis. Mild collapse results in less than or equal to 20% obstruction of the glottis and may not be of clinical significance in some cases. Horses with moderate collapse have 21% to 40% obstruction of the glottis and those with severe collapse have been reported to have 41% to 63% obstruction.

Axial Deviation of the Aryepiglottic Folds: Treatment

Horses with moderate and severe cases of axial deviation of the aryepiglottic folds and those with clinically significant mild axial deviation of the aryepiglottic folds are candidates for surgical treatment. Transendoscopic laser excision of the aryepiglottic folds (TLEAF) to remove a 2-cm isosceles right triangle of tissue from each collapsing aryepiglottic folds with use of a neodymium/yttrium-aluminum-garnet or diode laser in contact fashion is recommended. This approach is easier to perform in a sedated, standing horse with topical anesthesia, but it may be performed successfully with the horse anesthetized in lateral recumbency. The procedure may also be performed with the horse under general anesthesia through a laryngotomy with conventional instruments. The disadvantage for the clinician of performing the procedure this way is the inability to see the exact tissue being resected relative to its normal position to the larynx. If surgical resection is performed through the laser with general anesthesia, the horse is nasotracheally intubated and heliox (70% helium, 30% oxygen) should be mixed with 100% oxygen to achieve a fraction of oxygen in inspired air equal to 0.4 to prevent ignition while the laser is activated.

For surgery with TLEAF in the standing animal, horses are sedated with xylazine hydrochloride (0.4 mg/kg IV). Additional doses of xylazine hydrochloride (0.2 mg/kg IV) may be required. A videoendoscope is inserted into the nasal passage ipsilateral to the target aryepiglottic fold and held in place by an assistant. Topical anesthesia is achieved with an aerosolized solution that contains benzocaine hydrochloride (14%), butyl aminobenzoate (2%), and tetracaine hydrochloride (2%; Cetacaine) administered through polyethylene tubing (PE-240; Becton Dickinson, Sparks, Md.) passed through the biopsy channel of the videoendoscope.

Bronchoesophagoscopic forceps (Richard Wolf Medical Instrument, Vernon Hills, 111.), 60 cm in length and bent manually to conform to the curve of the equine nasal passage and pharynx, are used to provide traction on the aryepiglottic folds during excision. These forceps are passed into the nasal passage contralateral to the target aryepiglottic fold and are manipulated by a second assistant. The free margin of the membranous portion of the aryepiglottic fold is grasped halfway between the arytenoid and epiglottic attachments and elevated caudodorsally (). The laser is set to 18 W of power and excision of the tissue is performed in contact fashion. Beginning rostrally and immediately adjacent to the epiglottic attachment, the clinician makes a horizontal incision in the mucosa by sweeping the fiber side to side and gradually cutting tissue in a rostral to caudal direction. The grasping forceps are then rotated to apply traction to the aryepiglottic fold in a rostromedial direction. A vertical incision is then made from dorsal to ventral to cut the tissue adjacent to its attachments on the corniculate process of the arytenoid cartilage. The vertical incision is extended ventrally to intersect the horizontal incision and the tissue is removed with the grasping forceps. For bilateral excision, the videoendoscope and forceps are positioned in reverse for excision of the contralateral aryepiglottic fold.

To excite the aryepiglottic fold with the horse under general anesthesia, the horse’s mouth is held open with a mouth speculum and the soft palate is manually displaced dorsally. Active suction is used to evacuate smoke from the pharynx. The videoendoscope, grasping forceps, and suction tubing are all positioned in the oral cavity to perform the same surgical procedure. Surgical excision has been performed through a laryngotomy; however, this approach does not afford the same visual perspective of the surgical field as does the videoendoscopic approach.

Broad-spectrum antimicrobial therapy is given preoperatively and continued for 7 days postoperatively because of the open mucosal wound created in the larynx by excision of the aryepiglottic fold. Antiinflammatory therapy is recommended and should consist of tapering courses of phenylbutazone (2 mg/kg orally twice daily for 3-4 days, then once daily for 3-4 days), prednisolone (0.8 mg/kg orally once daily for 7 days, then 0.8 mg orally every other day for 3 treatments then 0.4 mg/kg orally every other day for 3 treatments), in addition to a topical pharyngeal spray (37 ml nitrofurazone solution [0.2%], 12 ml dimethyl sulfoxide [DMSO; 90%], 50 ml glycerine, and 0.2 ml prednisolone acetate [5%]; 10 ml twice daily for 7 days). The pharyngeal spray is administered though a 10-Fr male dog urinary catheter (Monoject, division of Sherwood Medical, St Louis, Mo.) that is placed up the ventral meatus of the nasal passage to a point level with the medial canthus of the eye. The pharyngeal spray is given slowly. If the horse swallows during administration, the catheter is correctly placed in the pharynx.

Postoperative management instructions for horses that have TLEAF should include at least two weeks of daily hand-walking or turnout in a small paddock. Additional rest may be indicated if other surgical procedures are performed for concurrent airway problems. Follow-up endoscopy is recommended before returning the horse to training. Postoperatively, the edge of the tissue will look slightly more fibrous and concave but not dramatically different than the preoperative appearance.

Some horses, especially younger animals and those with multiple upper respiratory abnormalities, may benefit from conservative management with prolonged rest. Additionally, these horses may benefit from longer periods of time between races when returned to training.

Prognosis of Axial Deviation of the Aryepiglottic Folds

In a retrospective study of racehorses with an exclusive diagnosis of axial deviation of the aryepiglottic folds as the cause of their poor performance, 75% of horses that had surgical excision of the aryepiglottic folds and 50% of the horses managed with rest had improved performance. Improvement of the upper respiratory noise is more likely to occur with surgical treatment. No complications have been recognized after surgical excision, and no adverse effects on deglutition or laryngeal or pharyngeal function have been reported.


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.


Stallion Behavior Problems

This post briefly outlines several of the most common behavior problems of breeding stallions. These problems include self-mutilation, inadequate libido, rowdy breeding behavior, specific erection dysfunction, mounting and thrusting difficulties, frenzied hyperactive behavior, and specific ejaculation dysfunction. Also briefly outlined is the common problem of residual stallionlike behavior in geldings.

Inadequate Libido

Specific stallion libido problems include slow starting novices, slow or sour experienced stallions, and specific aversions or preferences. Although certain genetic lines tend to be shy or quiet breeders, the majority of inadequate libido in stallions is man-made in the sense that it is the result of domestic rearing, training, or breeding conditions. Stallions that have been disciplined for showing sexual interest in mares during their performance career, discouraged from showing spontaneous erection and masturbation, or mishandled during breeding under halter are at risk of libido problems. When exposed to a mare for teasing, stallions such as these may simply stand quietly, may appear anxious and confused, or may savage the mare.

Most stallions with such experience-related libido problems respond well to behavior therapy alone or in combination with anxiolytic medication. These stallions typically respond best to continued exposure to mares, initially with minimal human presence, and then with gradual introduction of quiet, respectful, patient, positive reinforcement-based handling. These stallions appear to respond favorably to reassurance for even small increments of improvement. Tolerance of minor misbehavior rather than punishment is often the most effective strategy with low-libido stallions. The anxiolytic diazepam (0.05 mg/kg through slow IV 5-7 min before breeding) is useful in about half of such cases as an adjunct to behavior modification.

Some libido problems are hormone-related, with androgens on the low side of the normal range. These stallions will likely improve with management aimed at increasing exposure to mares and reduced exposure to other stallions. This will typically increase androgen levels, general confidence, as well as sexual interest and arousal. Gonadotropin-releasing hormone (GnRH; 50 μg SQ 2 hr and again 1 hr before breeding) can be useful to boost libido in stallions, particularly in those with low normal levels. In rare cases when more rapid improvement is required to rescue a breeding career, treatment with testosterone can effectively jump-start a slow novice without apparent significant adverse effects on spermatogenesis. Current recommendations are 0.1 to 0.2 mg/kg aqueous testosterone SQ every other day for as long as 2 weeks, with frequent assay of circulating testosterone not to exceed 4 ng/ml.

Specific Erection Dysfunction

Libido-independent erection dysfunction is rare in stallions. The majority of erection dysfunction that does occur is related to traumatic damage of the corpora cavernosa that results in insufficient or asymmetric tumescence (lateral or ventral deviations) that impairs insertion. In some instances, penile injury appears to impair sensory and or proprioceptive feedback from the penis, delaying ejaculation, coupling, or organized thrusting. Common causes include stallion ring injuries, drug-related paralyzed penis and paraphimosis, kick injuries, and self-serve breeding dummy accidents.

An interesting and often confusing type of erection dysfunction involves the folding back of the penis within the prepuce. The behavioral hallmark of this situation is a stallion that appears aroused and ready to mount, without a visible erection. The stallion may also appear uncomfortable or intermittently distracted, pinning the ears, kicking toward the groin, and/or stepping gingerly on the hind legs. Close observation reveals a rounded, full-appearing prepuce, with the skin stretched taut. Resolution usually requires removal of the stallion from the mare until the penis detumesces. Once the penis is fully withdrawn, application of a lubricating ointment to the prepuce facilitates subsequent normal protrusion. This situation tends to repeat occasionally over time, particularly in stallions with profuse smegma production or with dryness of the penis and sheath from frequent cleansing.

Mounting And Thrusting Difficulties

A significant percentage of breeding dysfunction appears to involve neurologic or musculoskeletal problems that affect the stallion’s ability to mount and thrust. Many such stallions can continue breeding with therapy aimed to reduce discomfort and accommodate disabilities during breeding, including adjustments to the breeding schedule aimed at reducing the total amount of work. This author has found that long-term treatment with oral phenylbutazone (2-3 mg/kg orally twice daily) often works well to keep such stallions comfortable for breeding. Certain debilitated stallions can benefit from semen collection while standing on the ground.

Specific Ejaculation Dysfunction

Although any libido, erection, or mounting and thrusting problem can result in failure to ejaculate, stallions also exist in which the dysfunction seems to be specific to ejaculation. Specific ejaculation problems can include apparent

failure of the neural ejaculatory apparatus, physical or psychologic pain associated with ejaculation, and genital tract pathology. Goals of therapy are to address as many contributing conditions as possible, as well as to optimize handling and breeding conditions and maximize musculoskeletal fitness and libido to enhance the stallion’s ability to overcome ejaculatory difficulty. Imipramine hydrochloride (0.5-1.0 mg/kg orally 2 hr before breeding) can effectively reduce the ejaculatory threshold.

Rowdy Breeding Behavior

Rowdy, misbehaved breeding stallions in most cases represent a human-animal interaction problem. Most problems can be overcome with judicious, skillful, respectful re-training. Even strong, vigorous, and misbehaved stallions can be brought under control by using consistent positive and negative reinforcement, with very little or no severe punishment. Re-training can be done in a safe and systematic manner without abuse or commotion, usually within a few brief sessions. Some of the most challenging, rowdy stallions may benefit from vigorous exercise under saddle or ground work immediately before breeding. This practice not only fatigues the stallion but also establishes a pattern of the stallion taking direction from a handler. For similar reasons, this author recommends an intensive schedule for breeding shed retraining, with as many as several breedings per day. With fatigue and reduced urgency to breed, many stallions seem more able to abide direction and learn a routine. With rapid repetition, stallions seem to more readily understand the routine. Tranquilization is generally not recommended. Levels of sedation that improve controllability without compromising musculoskeletal stability or ejaculatory function are difficult to achieve. Tranquilizing agents commonly used in stallions, such as xylazine or detomidine, can both facilitate and inhibit erection and ejaculation depending on dose.

Frenzied Behavior

Distinct from simple rowdiness, some stallions are hyperactive or even frenzied. This is typically greater during the breeding season. Some will spend nearly their entire time budget frantically “climbing the walls,” or running a fence line. In general frenzied breeding stallions can benefit from more roughage and less grain in the diet, organized physical work and pasture exercise, and consistent housing in a quiet area. Careful observation (particularly video surveillance) can be useful to identify environmental conditions and events that set off episodes or tend to quiet a stallion. In extreme cases, pasturing directly with mares can effectively quiet or sensibly occupy a frenzied stallion. L-Tryptophan supplementation (1-2 g twice daily in feed) can have a calming effect on such stallions. Tranquilization for this purpose is not recommended in breeding stallions because of risk of paralyzed penis and paraphimosis.


Although not unique to stallions, self-mutilation is a severe and relatively uncommon fertility limiting and/or life-threatening problem. This behavior typically takes the form of self-biting of the flank, chest, or limbs, with violent spinning, kicking, and vocalization. Self-mutilation in horses appears to occur in two distinct forms. One appears to be a severe reaction to irritation or pain, and would be similar in males or females. The self-biting is typically targeted toward the site of discomfort. Another form occurs in males and is reminiscent of stallion intermale aggression. The behavior is targeted at the typically intermale sites of aggression — the groin, flank, knees, chest, and hocks. The sequence of the behavior follows closely to that of two males fighting, with sniffing and nipping of the groin, vocalization, stamping with a fore leg, kicking out with a hind leg, and then taking occasional larger bites from anywhere on the opponent’s body.

Episodes often appear to be stimulated by sight, sound, or smell (feces or oily residues) of another stallion. For some stallions, episodes are set off by sniffing their own excrement or oily residues on stall walls or doorways. Current recommendations to control episodes are as follows: (1) physically protect the stallion from injury by padding walls or limbs, blanketing, and muzzling as effective; (2) aggressively evaluate the housing and social environment to identify exacerbating and ameliorating conditions that may be manipulated for greatest relief; (3) reduce concentrates and increase grass and hay in the diet to increase feeding time and eliminate highly palatable meals (feeding tends to distract and occupy the stallion; concentrate meals tend to increase stereotypic behavior); (4) apply odor-masking agents (Vicks or Acclimate) around the nares; and (5) provide as much organized exercise as possible, also to distract the stallion.

Residual Stallionlike Behavior In Geldings

Castration, regardless of age or previous sexual experience, does not always eliminate stallionlike behavior in horses. If given the opportunity, as many as half of geldings will show stallionlike behavior to mares, many will herd mares, and even mount and appear to breed. Similarly, although castration does tend to “mellow” most horses, it does not eliminate general misbehavior. Traditional behavior modification is usually much more effective in the control of sexual and aggressive behavior in a gelding under saddle or in-hand than it is with an intact stallion. Also, treatment aimed at quieting sexual and aggressive behavior, such as progesterone (e.g., altrenogest, 50-75 mg orally daily), is typically more effective in geldings than in intact stallions. Elimination of stallionlike herding and teasing at pasture is difficult. Separation from mares is recommended.


Interpretation of Peritoneal Fluid Changes in Peripartum Mares

Abdominal discomfort in the peripartum mare poses a diagnostic dilemma for the equine clinician because of the difficulty in differentiating between normal uterine contractions and other sources of abdominal pain. When a periparturient mare displays abdominal discomfort she may be experiencing a reproductive problem including uterine torsion or rupture, vaginal tear involving the peritoneal cavity, hematoma of the uterine wall, or a uterine artery rupture. Possible lesions in other abdominal organs include rupture of the urinary bladder or cecum, large colon impaction or torsion, or vascular compromise of a segment of bowel as a result of mesenteric rents or trauma. Several of these conditions can cause the affected mare to rapidly become depressed and febrile, with accompanying signs of shock and toxemia. A prompt and accurate diagnosis followed by aggressive medical and/or surgical intervention can often prevent an otherwise fatal outcome.

Recently, transabdominal ultrasonography has become an integral part of the diagnostic evaluation of the equine abdomen. The quantity and cellularity of fluid accumulated in the ventral abdomen is readily seen with a 3.5-MHz probe. However, detection of abnormalities in peritoneal fluid is still extremely useful when managing equine colic. The significance of these abnormalities in peripartum mares that are experiencing abdominal discomfort has only recently been widely appreciated. Intuitively, because peritoneal fluid composition reflects the pathophysiologic state of the visceral and parietal mesothelial surfaces, one would anticipate that the mechanics of the foaling process (and certainly obstetrical manipulations) would be likely to incite some changes in peritoneal fluid composition.

Obtaining A Peritoneal Fluid Sample In A Periparturient Mare

The mare is sedated with xylazine hydrochloride (0.3 mg/kg of body weight, IV) and butorphanol tartrate (0.01 mg/kg, IV) if needed. The most dependent portion of the ventral abdomen is clipped, shaved, and aseptically prepared. The abdominocentesis is performed as far cranial as possible, and approximately half an inch to the right of midline, to avoid penetrating the spleen or gravid uterus. An 18-gauge, 1.5-inch needle is introduced through the skin and slowly advanced into the abdominal cavity. If no fluid is obtained, the needle should be repositioned. Sometimes rotation of the needle or injection of a small volume of air with a sterile syringe is necessary to facilitate drainage of fluid. A minimum of 1.0 ml of peritoneal fluid should be collected in a tube that contains sodium ethylenediaminetetraacetic acid (EDTA) as an anticoagulant. In clinically ill mares, it maybe difficult to obtain a sample, especially if the mare is dehydrated or has a large ventral plaque of edema that extends cranially from the mammary glands. If difficulty is experienced in obtaining a sample, a local anesthetic block can be made over the site and a stab incision made through the skin, subcutaneous tissue, and muscular fascia with use of a number 15-scalpel blade. A blunt teat cannula or sterile female catheter can then be carefully advanced into the peritoneal cavity. The incision is allowed to heal by second intention.

Potential risks of an 18-gauge needle used to perform abdominocentesis include inadvertent laceration of the spleen or intestinal puncture (enterocentesis). Contamination of the sample with blood is more likely to be caused by penetration of superficial blood vessels or vessels in the abdominal musculature. This does not affect the usefulness of the sample because as much as 17% blood contamination does not alter the interpretation of the nucleated cell count or the total protein concentration of peritoneal fluid samples. Inadvertent enterocentesis may cause a transient increase in the nucleated cell count. Studies have shown that intestinal puncture with a needle rarely causes clinical signs of disease. Likewise, repeated abdominocentesis at 24- to 48-hour intervals has been shown to not alter the peritoneal fluid composition. Thus monitoring progressive changes in the peritoneal fluid may alert the clinician to the presence of a deteriorating condition in the abdominal cavity.

Peritoneal Fluid Analysis

The peritoneal fluid is visually inspected for color and clarity (turbidity). Specific gravity and total protein concentration (TPr) estimations can be made with a hand-held refractometer. Total white blood cell (WBC) counts can be determined manually with a hemocytometer, or measured by using an automated analysis system. Differential white blood cell counts can be made by viewing 100 cells on a smear that is stained with Wright’s stain. A direct smear can be made if the WBC count exceeds 10,000 cells/|xl. When the count is low the sample should be centrifuged to concentrate the cells. The white blood cell population of the peritoneal fluid consists of a mixture of nondegenerate neutrophils and large mononuclear cells. The latter are a combination of mesothelial cells that desquamate from the peritoneal surface, as well as blood-borne monocytes and macrophages that have migrated into the peritoneal cavity.

Normal values for an adult horse may vary between laboratories. A sample is classified as a transudate if the total protein concentration is less than 2.5 g/dl and the nucleated cell count is less than 5000 cells/μl. Modified transudates are characterized by an increase in TPr concentration or WBC count. If the total protein concentration exceeds 3.0 g/dl and the nucleated cell count exceeds 10,000 cells/μl then the sample is classified as an exudate. The normal differential WBC count is approximately 60% neutrophils and 40% mononuclear cells. As much as 70% neutrophils (%N) is considered normal in equine peritoneal fluid. In acute inflammatory processes (i.e., peritonitis) the %N in peritoneal fluid may increase to 85% to 100%.

Assessment of cell morphology through cytologic examination is an extremely important part of any peritoneal fluid analysis. The morphologic characteristics of the cell types can be used to differentiate between septic and nonseptic inflammation. Nondegenerate neutrophils predominate in transudates and mild exudates. Degenerate neutrophils are characterized by nuclear pyknosis, karyorrhexis, karyolysis, and cytoplasmic vacuolization. A large number of degenerate neutrophils indicates bacterial toxin-induced cell disruption, and they predominate in septic effusions, and a guarded-to-grave prognosis is warranted. Detection of phagocytosed bacteria confirms the presence of a septic process.

Effect Of Obstetric Conditions On Peritoneal Fluid


Determination of Fetal Gender

Fetal gender determination has been incorporated into the management programs of many breeding farms. Depending on the sire or the dam, the fetal gender may affect the value of the fetus and therefore influence the value of the pregnant mare. This knowledge could change various management decisions such as appraisals, foaling location, sales’ reserves, insurance coverage, collateral limits for loans, mating lists for the next year, and buy/sell decisions.

Table Development of Ultrasound Findings that Indicate Fetal Gender shows what to expect at the different stages of fetal development. The basis for sex determination when the fetus is between 55 and 90 days’ gestation is the location of the genital tubercle — a bilobulated hyperechoic structure 2 to 3 mm in length. This structure resembles a brightly colored “equals” symbol (), and is the precursor for the penis in the male and the clitoris in the female. The tubercle develops between the hind legs on the ventral midline in both sexes and at approximately day 53 or 54 of gestation appears to begin migrating toward the umbilical cord in the male and toward the anus in the female. Location of the tubercle between 55 and 90 days’ gestation enables the practitioner to determine the gender.

Table Development of Ultrasound Findings that Indicate Fetal Gender

Day(s) Ultrasound Findings
55-60 Fetus is very small; genital tubercle is difficult to see; tubercle may or may not be fully migrated.
60-70 Ideal time for examination — fetal tubercle is distinct and fully migrated; fetus is accessible for viewing.
70-80 Fetus becomes more difficult to reach.
80-90 Most difficult time to view fetus — tubercle is less distinct; genitalia development is just beginning; fetus is frequently out of reach.
90-100 Fetus is generally accessible, but genitalia are not very well-developed.
100-110 Genitalia are becoming more evident.
110-120 Ideal time — genitalia is well-developed.
120-140 Genitalia is well-developed, but posterior of fetus may be difficult to access at times.
140-150* At times the fetus has anterior presentation with posterior out of the examiner’s reach.
150+ Usually the fetus has anterior presentation, and the posterior of the fetus is out of the examiner’s reach.

*Mares of 130 to 150 days’ gestation that are classified as being “out of reach” should be viewed again for possible position changes.

When the fetus has reached 55 to 80 days’ gestation, a veterinarian should be able to make a gender diagnosis 95% of the time with one examination. The accuracy of fetal sexing should reach 99%, and the time required to make the determination should range from a few seconds to 5 minutes depending on the experience of the clinician. At 80 to 90 days the fetus is temporarily difficult to reach due to the positioning of the uterus in the posterior abdomen. At approximately 80 days the fluid of the pregnancy pulls the uterus over the rim of the pelvis. The fetus is small, falls to the ventral portion of the uterus, and is difficult to reach. As the fetus grows, the uterus actually elevates more in the abdominal cavity and the fetus becomes easier to reach and view (). After 90 days’ gestation, the tubercle becomes less distinct and more difficult to see. Therefore, the clinician must rely on developing external genitalia — in the female, the mammary gland, teats, and clitoris (), and in the male, the penis and prepuce () — to make the gender diagnosis. Consistent differentiation between male and female gonads at differing stages of gestation is difficult (). Consequently, gonads are used only for the reinforcement of a diagnosis, not for the diagnosis itself. At 90 to 150 days of gestation, a veterinarian should be able to formulate a highly accurate gender diagnosis 85% to 90% of the time. The diagnosis should require a few seconds to 10 minutes to perform, again, depending on the experience of the practitioner.

An attempt to sex the fetus should not last for more than 10 minutes per session on any one mare, because depending on the type of restraint used she may become fractious. No person or mare should experience injury during this elective procedure. If the mare becomes fractious, the veterinarian should stop and attempt the procedure on another day.


A high-quality ultrasound machine with a 5-MHz linear array rectal transducer (ALOKA 500 SSD [Aloka Co, Ltd., Wallingford, Conn.] or equivalent) is essential. If the overall gain, near gain, or far gain are set too high, the contrast between the fetus and background is less and the tubercle is more difficult to see. For optimal close eye level viewing, an ultrasound stand on wheels, subdued lighting for good screen visibility, and a hat are recommended. Video recordings or printers are helpful to verify the diagnosis and for record keeping.

Rectal palpation essentials include lubricant and sleeves and adequate restraint that might include a twitch, stocks, or tranquilizer. Depending on the situation, use of tranquilizers is acceptable, but this may cause the uterus to relax and drop away from the examiner and become more difficult to reach. This author uses xylazine (200 mg IV) mixed with butorphanol tartrate (10 mg). Propantheline bromide (30 mg IV) may be used to prevent rectal straining. Fly spray (if needed) will help to keep mare movement to a minimum.


The procedure involves a thorough evacuation of the feces from the rectum to allow easy manipulation of the transducer. The clinician can determine the position of the fetus by scanning the entire fetus. The skull is a good anterior marker, the heart a good ventral marker, ribs coming off the vertebrae and the base of the tail are good dorsal markers, and the tail is a good posterior marker.

Once the position of the fetus has been determined, the veterinarian should proceed with the transducer to the posterior part of the fetus until the image of the latter has gone completely off the transducer. The veterinarian should gradually ease back onto the fetus with the transducer and cut a plane perpendicular to the axis of the spine of the fetus. A cross-section of the tailhead should be picked up on the dorsal aspect of the fetus. This cross-section will appear as a hyperechoic round mass with very little muscle tissue around it. On the ventral aspect of the fetus, two tibias, which appear as hyperechoic round structures with no muscle mass, should be seen to form a triangle with the tailhead. If the fetus is female, a hyperechoic tubercle will appear within the tibia-tailhead triangle with the tubercle slightly toward the tailhead. The female tubercle is difficult to consistently identify other than within the tibia-tailhead triangle If the fetus is a male, nothing will be seen within the tibia-tailhead triangle. If this is the case, the veterinarian should move the transducer gradually along the anterior aspect of the fetus, keeping the same perpendicular plane (plane II) to the axis of the spine of the fetus. A hyperechoic structure resembling an “equals” symbol should be seen between the two tibias, which when the transducer is moved anteriorly become the stifles or femurs (). It is important to remember that tibias have no muscle mass around them but femurs do have muscle mass around them. When the transducer is moved further anteriorly, the large round abdomen will be seen. The male genital tubercle can often be seen on the outside ventral wall of the abdomen just posterior to the urachus, which is seen as a dark hole 4 to 5 mm in diameter. When the transducer is moved back and forth over the posterior area in this plane a tubercle is usually seen.

The male tubercle can also be readily seen from a frontal plane, This plane exhibits the front legs, ventral abdomen, and hind legs with the tubercle appearing slightly anterior to a line drawn between the hind legs (usually femurs or stifles).

After 90 days of gestation, the tubercle is less distinct. The veterinarian should proceed to the posterior aspect of the fetus and find the point at which the posterior muscles of the buttocks come together and form a definite cleavage on the ventral midline. Next, the cleavage should be followed posteriorly to the tailhead. If the fetus is a female, a clitoris will appear as a small round structure in the cleavage shortly before the tailhead is reached. If the small round structure is too close to the tailhead, it could be the anus. If the midline or cleavage line is followed anteriorly, the mammary gland in the female or the prepuce and penis in the male are encountered. A mammary gland appears as a triangular, slightly denser tissue than surrounding muscle tissue. Each half of the gland may have two bright hyperechogenic teats and slightly dense areas; a translucent division between halves of the mammary gland may also be present. If the prepuce is viewed from a cross-section across the posterior ventral abdomen, it will appear as a cone-shaped structure off the ventral abdominal wall just posterior to the urachus. The prepuce may appear as a cone shaped structure with a hyperechoic area within the cone. The hyperechoic area is the penis itself. Sometimes the shaft of the penis can be seen with a hyperechoic distal segment .

When the fetus gets older, the genitalia become more developed and more easily differentiated from surrounding tissue. When the fetus develops, however, it becomes more difficult to access the posterior area. After day 150 of gestation, the fetus begins to have an anterior presentation that puts the posterior area out of reach. Also, because of its size the fetus is less apt to rotate the posterior part to a more accessible position. A fetal sex determination has been made at 184 days on trans-rectal exam, but this is very unusual and possibly not a good sign if the fetus is in a posterior presentation this late in gestation.

This procedure is for gender identification only and not for gender control. It would be unusual to have a mare successfully pregnant at 60 days, terminate the pregnancy, and be successfully re-mated that season.

Mastery of these sex determination techniques by veterinarians provides a worthwhile service to clients but requires many hours of actual sonographic visualization of the equine fetus at between 55 and 150 days of gestation. Only when the sonographic cross-sectional anatomy of the fetus is learned will a consistent, accurate diagnosis of the fetus’ sex be made.