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 Ear: General Principles Of Management

The therapeutic plan for otitis externa requires identification of the primary disease process and perpetuating factors. Ideally management is aimed at thoroughly cleaning and drying the ear canal, removing or managing the primary factors, controlling perpetuating factors, administering appropriate topical or systemic therapy (or both), and evaluating response to therapy.

Ear Cleaning

Ear cleaning serves several functions: (1) it removes material that supports or perpetuates infection; (2) it removes bacterial toxins, white blood cells (WBCs), and free fatty acids that stimulate inflammation; (3) it allows complete evaluation of the external ear canal and tympanum; (4) it allows topical therapy to contact all portions of the ear canal; and (5) it removes material that may inactivate topical medications. Significandy painful ears may benefit from initial anti-inflammatory therapy to decrease pain and swelling of the ear canal prior to cleaning. Severe cases of otitis externa often require general anesthesia to facilitate complete cleaning and evaluation of the external and middle ear.

Many different solutions are available for removing cerumen, exudate, and debris from the ear canal (Table Otic Cleaning Solutions). If the tympanic membrane cannot be visualized, only physiologic saline solution or water should be used, because many topical cleaning agents are ototoxic or incite inflammation of the middle ear. An operating otoscope, ear loops, and alligator forceps facilitate manual removal of large amounts of cerumen or debris. Debris is carefully removed under direct visualization, and care is taken deeper in the ear canal (close to the tympanic membrane). Aggressive hair removal is not advised, because inflammation and damage to the epithelium can result in secondary bacterial colonization and infection. Flushing may be performed after large accumulations of cerumen and debris are mechanically removed from the ear canal.

Otic Cleaning Solutions

Trade Name Acetic Acid Boric Acid Salicylic Acid Isopropyl Alcohol Propylene Clycol Dss Other
Ace-Otic Cleanser 2%   0.1%       Lactic acid 2.7%
Adams Pan-Otic         X X Parachlorometaxylenol, tris EDTA, methylparaben, diazolidinyl urea, popylparaben, octoxynol
Alocetic Ear Rinse X     X     Nonoxynol-12, methylparaben, alovera gel
Cerulytic Ear Ceruminolytic         X   Benzyl alcohol, butylated hydroxytoluene
Cerumene             25% Isopropyl myristate
DermaPet Ear/Skin Cleanser for Pets X X          
Docusate Solution         X X  
Earmed Boracetic Flush X X         Aloe
Earmed Cleansing Solution & Wash         X   50A 40B alcohol, cocamidopropyl phosphatidyl and PE dimonium chloride
Earoxide Ear Cleanser             Carbamide peroxide 6.5%
Epi-Otic Ear Cleanser     X   X X Lactic acid, chitosanide
Fresh-Ear X X X X X   Lidocaine hydrochloride, glycerin, sodium docusate, lanolin oil
OtiCalm     X       Benzoic acid, malic acid, oil of eucalyptus
Otic Clear X X X X X   Glycerin, lidocaine hydrochloride
Oticlean-A Ear Cleaning Lotion X X X 35% X   Lanolin oil, glycerin
Oti-Clens     X   X   Malic acid, benzoic acid
Otipan Cleansing Solution         X   Hydroxypropyl cellulose, octoxynol
Otocetic Solution 2% 2%          
Wax-O-Sol 25%             Hexamethyltetracosane

Flushing and evacuation of solution is done under direct visualization through an operating otoscope. A bulb syringe and red rubber catheter system may be used to both flush and evacuate solutions and accumulations from the ear canal. The operator, avoiding drastic pressure changes within the external ear canal that could damage the tympanum, should carefully control suction and manual evacuation of the contents of the bulb syringe from the ear canal. Other alternatives include tomcat catheters (3.5 F) or flexible, intravenous catheters (14 gauge, Teflon); stiff, narrow catheters should be used cautiously and under direct visualization deep in the external ear canal. Other reservoir systems for delivery or evacuation of solutions include a 12 mL syringe or suction tubing attached to in-house vacuum systems. In-house vacuum systems should be used cautiously and under direct visualization. Care should be taken to avoid trauma to the tympanic membrane until its integrity can be assessed. Initial flushes should be done with physiologic saline solution or water until the integrity of the tympanic membrane is established.

Other solutions may aid in the removal of wax in the ear canal. Ceruminolytics are emulsifiers and surfactants that break down ceruminocellular aggregates by causing lysis of squamous cells. A ceruminolytic agent in an alkaline pH may more effectively lyse squamous cells via cell surface protein disruption. Oil-based products soften and loosen debris to aid in their removal but do not cause cell lysis. Water-based ceruminolytics are easier to remove and dry more quickly than oil-based solutions, which are occlusive if they remain in the ear canal. Water-based products include dioctyl sodium sulfosuccinate, calcium sulfosuccinate, and carbamate peroxide, which has a foaming action with the release of urea and oxygen. Oil-based products include squalene, triethanolamine polypeptide, hexamethyltetracosane, oleate condensate, propylene glycol, glycerin, and mineral oil. In a recent study only the combination of squalene and isopropyl myristate in a liquid petrolatum base had no adverse effects on hearing, the vestibular system, and histopathologic examination. Other agents tested contained glycerin, dioctyl sodium sulfosuccinate (2% or 6.5%), parachlorometaxylenol, carbamide peroxide (6%), propylene glycol, triethanolamine polypeptide oleate condensate (10%), and chlorobutanol (0.5%).

Alcohol-based drying agents added to ceruminolytics include boric acid, benzoic acid, and salicylic acid, which decrease the pH of the ear canal, cause keratolysis, and have a mild antimicrobial effect. Drying the ear canal is important to combat increased humidity, which potentiates infection.

If the tympanum is intact, the ear canal is filled with a ceruminolytic agent for at least 2 minutes and the pinna is cleaned at the same time. The solution is flushed twice with warm water, and the canal inspected. The procedure is repeated until cleaning is complete. Other solutions commonly advocated for ear flushing include dilute chlorhexidine solution (0.05%), dilute povidone-iodine, and acetic acid (2.5%). The first two agents are potentially ototoxic or induce inflammation and should not be used if the tympanum is ruptured. A combination of propylene glycol malic, benzoic, or salicylic acid; 2% acetic acid; or dilute povidone-iodine have been suggested for use in dogs with a ruptured tympanum.

Owners may clean the ears at home with mild preparations of ceruminolytics and drying agents if mild otitis is present without severe accumulation of cerumen or exudate. Aqueous solutions are usually recommended because they are less occlusive and easier to clean from the ear, dog, and home environment.

The ear should be filled with the solution, then massaged for 40 to 60 seconds. The pet should be allowed to shake its head to remove the majority of the solution, and the excess should be wiped from the ear canal and pinna with a tissue. Daily flushing is usually recommended, followed by every other day, weekly, then as needed, depending on the solution. Ear swabs are not recommended for home use, because cerumen and debris may be forced into the horizontal ear canal and impact against the tympanic membrane

Topical Therapy

Erythematous ceruminous otitis externa is diagnosed 2.7 times more often than acute suppurative otitis according to one report. Yeast ± cocci were identified in those cases, with cocci or rods identified in suppurative otitis. Topical therapy should be based on the cytologic examination to diminish the incidence of inappropriate treatment (Table Topical Medications Used in the Treatment of Ear Disease). Many preparations combine anti-inflammatories and antimicrobials in an attempt to decrease the inflammation and combat bacterial or yeast overgrowth. All topical medications should be considered supportive, and specific treatment should be aimed at controlling the primary disease process.

Topical Medications Used in the Treatment of Ear Disease

Generic Name Trade Name Dose Frequency Description
Fluocinolone 0.01% DMSO 60% Synotic 4-6 drops; total dose<17mL q12h initially. q48-72h maintenance Potent corticosteroid anti-inflammatory
Hydrocortisone 1.0% HB101,

Burrows H,

2-12 drops, depending on ear size q12h initially. q24-48h maintenance Mild corticosteroid anti-inflammatory
Hydrocortisone 1.0%, lactic acid Epiotic HC 5-10 drops q12h for 5 days Mild corticosteroid anti-inflammatory, drying agent
Hydrocortisone 0.5%, sulfur 2%. acetic acid 2.5% Clear X Ear Treatment 2-12 drops, depending on ear size q12-24h initially. q24-48h maintenance Mild corticosteroid anti-inflammatory, astringent, germicidal
DSS 6.5%. urea (carbamide peroxide 6%) Clear X Ear Cleansing Solution 1-2 mL per ear Once per week to as necessary Ceruminolytic, lubricating agent
Chlorhexidine 2% Nolvasan Dilute 1:40 in water As necessary Antibacterial & antifungal activity
Chlorhexidine 1.5% Nolvasan Dilute 2% in

propylene glycol

q12h Antibacterial & antifungal activity
Povidone-iodine 10% Betadine solution Dilute 1:10-1:50 in water As necessary Antibacterial activity
Polyhydroxidine iodine 0.5% Xenodyne Dilute 1:1-1:5 in water As necessary, q12h, once weekly Antibacterial activity
Acetic acid 5% White vinegar Dilute 1:1-1:3 in water As necessary; q12-24h for Pseudomonas Antibacterial activity, lowers ear canal pH
Neomycin 0.25%, triamcinolone 0.1%, thiabendazole 4% Tresaderm 2-12 drops depending on ear size q12h up to 7 days Antibacterial & antifungal activity, parasiticide (mites), moderate corticosteroid anti-inflammatory
Neomycin 0.25%, triamcinolone 0.1%, nystatin 100,000 U/mL Panalog 2-12 drops depending on ear size q12h to once weekly Antibacterial & antifungal activity, moderate corticosteroid anti-inflammatory
Chloramphenicol 0.42%. prednisone 0.17%, tetracaine 2%, squalene Liquachlor, Chlora-Otic 2-12 drops depending on ear size q12h up to 7 days Antibacterial activity, mild corticosteroid anti-inflammatory
Neomycin 1.75 & polymyxin B 5000 lU/mL, penicillin C procaine 10,000 lU/mL Forte Topical 2-12 drops depending on ear size q12h Antibacterial activity
Centamicin 0.3%, betamethasone valerate 0.1% Centocin Otic Solution, Betagen Otic Solution 2-12 drops depending on ear size q12h for 7 to 14 days Antibacterial activity, potent corticosteroid anti-inflammatory
Centamicin 0.3%, betamethasone 0.1%, clotrimazole 0.1% Otomax, Obibiotic Ointment 2-12 drops depending on ear size q12h for 7 days Antibacterial & antifungal activity, potent corticosteroid anti-inflammatory
Centamicin 0.3%, betamethasone valerate 0.1%, acetic acid 2.5% Centaved Otic Solution 2-12 drops, depending on ear size q12h for 7 to 14 days Antibacterial activity, potent corticosteroid anti-inflammatory
Polymixin B 10,000 lU/mL, hydrocortisone 0.5% Otobiotic 2-12 drops, depending on ear size q12h Antibacterial activity, mild corticosteroid anti-inflammatory
Enrofloxacin 0.5%, silver sulfadiazine 1% Baytril Otic 2-12 drops, depending on ear size q12h for up to 14 days Antibacterial activity
Carbaryl 0.5%, neomycin 0.5%, tetracaine Mitox Liquid 2-12 drops, depending on ear size   Antibacterial activity, parasiticide (mites)
Pyrethrins 0.06%, piperonyl butoxide 0.6% Ear Mite and Tick Control 5 drops q12h Parasiticide (mites)
Pyrethrins 0.05%, squalene 25% Cerumite 2-12 drops, depending on ear size q24h for 7 to 10 days Parasiticide (mites), ceruminolytic
Isopropyl alcohol 90%, boric acid 2% Panodry Fill ear canal As necessary Drying agent
Acetic acid 2%, aluminum acetate Otic Domeboro Fill ear canal q12-48h Drying agent, antibacterial activity, lowers ear canal pH
Silver sulfadiazine Silvadene Dilute 1:1 with water, 1 g powder in 100 mL water q12h for 14 days Antibacterial & antifungal activity
Tris EDTA±

gentamicin 0.03%

  2-12 drops, depending on ear size q12h for 14 days 1 L distilled water, 1.2g Tris EDTA, 1 mL glacial acetic acid; antibacterial activity
Silver nitrate   Use sparingly As necessary Cauterization of

ulcerative otitis externa

Miconazole 1%; ± topical glucocorticoid (7.5 mL of dexamethasone phosphate (4 mg/mL] to10mLof1% miconazole) Conofite 2-12 drops, depending on ear size q12-24h Antifungal activity
Ivermectin 0.01% Acarexx 0.5 mL per ear Once Parasiticide (mites)
Pyrethrins 0.15%, piperonyl butoxide 1.5% Many 2-12 drops, depending on ear size Twice at 7-day interval Parasiticide (mites)
Pyrethrins 0.05%, piperonyl butoxide 0.5%, squalene 25% Cerumite 2-12 drops, depending on ear size q24h for 7 days Parasiticide (mites), ceruminolytic
Pyrethrins 0.04%, piperonyl butoxide 0.49%, DSS 1.952%, benzocaine 1.952% Aurimite 10 drops q12h  
Rotenone 0.12%, cube resins 0.16% Many 2-12 drops, depending on ear size Every other day Parasiticide (mites)

Topical glucocorticoids benefit most cases of otitis externa by decreasing pruritus, exudation, swelling, and proliferative changes of the ear canal. The most potent glucocorticoids available in topical preparations are betamethasone valerate and fluocinolone acetonide. Less potent corticosteroids include triamcinolone acetonide and dexamethasone; the least potent is hydrocortisone. Most dogs benefit from short-term therapy with topical corticosteroids at the initiation of therapy, with concurrent therapy aimed at the primary and other perpetuating factors. Long-term therapy with topical corticosteroids can be deleterious because of systemic absorption of drug. Increased serum liver enzymes and depressed adrenal responsiveness may occur; with prolonged use iatrogenic hyperadreno-corticism is possible. Glucocorticoids alone may be of benefit for short-term therapy in cases of allergic or erythematous ceruminous otitis.

Antimicrobials are important for controlling secondary bacterial or yeast overgrowth or infection. Antimicrobials are indicated in any case with cytologic evidence of bacterial overgrowth or infection, with attention paid to the morphology and gram-staining characteristics of the bacteria. Otic preparations commonly contain aminoglycoside antibiotics. Neomycin is effective against typical otitis bacteria such as Staphylococcus intermedium. Gentamicin and polymyxin B are also appropriate initial topical treatments for gram-negative bacterial otitis externa.The significant risk of bone marrow toxicity in people limits the use of chloramphenicol for treating otitis in dogs and cats despite its antibacterial spectrum and availability.

Due to the frequency of resistant gram-negative bacteria such as Pseudomonas, other topical preparations have been developed. Enrofloxacin, ophthalmic tobramycin, and topical application of injectable ticarcillin have been used to treat otitis in dogs.< Their use should be limited to cases of resistant bacteria, and culture and susceptibility testing should be performed prior to application. Other topical agents may be used to supplement treatment of resistant Pseudomonas, such as silver sulfadiazine solution and tris EDTA. Tris EDTA can render Pseudomonas susceptible to enrofloxacin or cephalosporins by enhancing membrane permeability and altering ribosome stability. Frequent ear cleaning may also assist in the treatment of resistant bacterial otitis; ceruminolytics have antimicrobial properties, and their use in clinical cases has been evaluated. Acetic acid in combination with boric acid is effective against both Pseudomonas and Staphylococcus, depending on concentration and duration of exposure. Ear cleaning removes proinflammatory products, cells, and substances that diminish the effectiveness of topical antibiotics.

Many topical preparations control yeast organisms, which may complicate erythematous ceruminous otitis and suppurative otitis. Common active ingredients include miconazole, clotrimazole, nystatin, and thiabendazole. Preparations containing climbazole, econazole, and ketoconazole have also been evaluated. Eighty percent of yeast were susceptible to miconazole and econazole, intermediately resistant to ketoconazole, and 90% were resistant to nystatin and amphotericin B in one in vitro study. Topical ear cleaning agents have some efficacy against Malassezia organisms. Other preparations (e.g. chlorhexidine, povidone-iodine, acetic acid) are also effective in the treatment of secondary yeast overgrowth.

Response to topical therapy should be gauged by re-evaluation of physical, cytologic, and otoscopic examinations every 10 to 14 days after the initiation of therapy. Any changes in the results of these examinations should be recorded. Most cases of otitis can be managed topically; failure to respond to therapy should prompt re-evaluation of the diagnosis and treatment.

Systemic Therapy

Systemic glucocorticoid administration may be beneficial in cases of severe, acute inflammation of the ear canal, chronic proliferative changes of the ear canal, and allergic otitis. Anti-inflammatory doses should be limited to 7 to 10 days. Cases of significant thickening or proliferative changes in the external ear canal benefit from systemic antimicrobial therapy. Systemic therapy should be considered if concurrent dermatologic changes of the surrounding skin, pinna, or other regions of the body are present. Long-term administration of appropriate antimicrobials based on culture and susceptibility is required in all cases of otitis media. Systemic therapy for yeast is rarely recommended in animals with otitis alone. One study evaluated oral itraconazole therapy, and in ear samples evaluated on cytology and culture, no change in cytology score was found.



Therapy For Specific Diseases Of The External Ear Canal


Thorough cleaning of the external ear canal, treatment of all household pets, and whole-body therapy should be considered in the treatment regimen for ear mites. Pets with no clinical signs may be asymptomatic carriers and a reservoir for reinfestation. Otic parasiticides such as pyrethrins, rotenone, amitraz, and carbaryl must be administered every 24 hours throughout the 20-day mite life cycle because they do not kill mite eggs. Thiabendazole eliminates all mite stages, but it must be applied every 12 hours for 14 days. Ivermectin (0.3 to 0.5 mg/kg) may be applied topically once weekly for 5 weeks. Otic administration of medication does not affect mites on adjacent or distant skin locations, and systemic or other total-body parasiticide may be indicated. Alternatively, ivermectin administered subcutaneously (0.2 to 0.3 mg/kg) 2 to 3 times at 10- to 14-day intervals or orally (0.3 mg/kg) every week for four treatments eliminates otic mites and those found elsewhere on the body. Other topicals proven safe and effective for ear mite treatment include selamectin (6 mg/kg) applied to the skin between the shoulder blades and fipronil spray. Selamectin administered once in cats and two times, 30 days apart in dogs gave results similar to topical pyrethrin therapy.

Idiopathic Inflammatory or Hyperplastic Otitis in Cocker Spaniels

Treatment is aimed at decreasing the secondary ear canal changes associated with this condition. Anti-inflammatory doses of corticosteroids administered orally may be useful. Topical corticosteroid preparations in combination with antimicrobials decrease the soft tissue mass affecting the ear canal but may not be as effective as oral administration. Maintenance therapy may be required both topically and orally; however, low doses of corticosteroids should be used. Re-evaluation should include attention to the potential side effects of corticosteroid therapy. Intermittent treatment of secondary bacterial or yeast overgrowth and infection may be required. Surgery is often indicated due to the severe secondary changes within the ear canal.

Excessive Moisture (Swimmer’s Ear)

Other primary disease conditions such as allergic otitis should be ruled out in any dog with erythematous ceruminous otitis. Dogs with frequent exposure to water, however, may require ear cleaning and drying agents to diminish the humidity of the ear canal. Many cleaning and drying agents also posses antimicrobial effects. Products that combine a drying agent and corticosteroid decrease the ear canal humidity and inflammation associated with allergic otitis complicated by swimming. Care should be taken to control primary disease (i.e. allergic otitis), however, and intermittendy manage the predisposing factor (i.e. excessive moisture) as necessary. The dog’s ears should be cleaned and dried the day of water exposure and for 2 to 5 days after. For continued frequent exposure, maintenance cleaning may be required every other day to twice weekly.

Chronic Bacterial Otitis

Resistant bacteria play an important role in the development of chronic otitis externa. Any dog not responding to initial therapy should be re-evaluated for primary and perpetuating conditions such as allergic disease, foreign body, neoplasia, otitis media, and secondary anatomic changes of the ear canal. Primary disease processes identified in one study included hypothyroidism, atopy, food allergy, and immune-mediated disease. Infection with Pseudomonas species frequently occurs with repeated treatment of otitis extema, and acquired resistance is common. Culture and susceptibility testing is imperative to guide therapy. Oral antimicrobials combined with topical therapy are used in severe cases with secondary changes of the ear canal. Identification of otitis media is vital to remove the middle ear as a source of otitis extema. Otitis media requires long-term treatment.

Ear cleaning prior to the application of topical medication may increase the efficacy of the agent by decreasing exudate in the ear canal that inactivates antimicrobial drugs such as polymyxin. In cases that fail to respond to first-line drug treatments such as polymyxin or gentamicin, other topical antimicrobial agents should be tried. Ophthalmic tobramycin and injectable amikacin have been described for use as topical antimicrobials in ear disease. The integrity of the tympanic membrane should be known prior to use; the clinician should avoid these medications if the tympanic membrane cannot be proven intact. Enrofloxacin or ticarcillin injectable preparations diluted in saline or water may be applied topically for resistant Pseudomonas. Parenteral ticarcillin was used in cases with a ruptured tympanic membrane until healing was observed, at which time topical therapy was instituted; clinical response occurred in 11 of 12 cases. Enrofloxacin and silver sulfadiazine combination is also available in an otic preparation (Baytril Otic, Bayer Shawne Mission, KS).

Other topical therapy may assist in eliminating resistant Pseudomonas from the ear canal.

Decreasing the pH of the ear canal with 2% acetic acid is lethal to Pseudomonas; diluted vinegar in water (1:1 to 1:3) may be used to flush the ear canal. Acetic acid combined with boric acid is lethal to Pseudomonas and Staphylococcus, depending on the concentration of each agent. Increasing the concentration of acetic acid may broaden its spectrum of activity but causes irritation of the external and middle ear. Silver sulfadiazine in a 1% solution exceeds the minimum inhibitory concentration of Pseudomonas and may be instilled into the ear canal. One gram of silver sulfadiazine powder mixed in 100 mL of water may be used for topical therapy and is also effective against Proteus species, enterococci, and Staphylococcus intermedium. Dilute acetic acid (2%) and silver sulfadiazine (1%) have not caused adverse effects in cases with a ruptured tympanic membrane.I Tris EDTA may be applied after thorough ear cleaning to increase the susceptibility of Pseudomonas to antimicrobial agents. It must be mixed, pH adjusted, and autoclaved prior to use or is available in an otic preparation (TrizEDTA, DermaPet ®, Potomac, MD), which is used to clean the ears prior to instillation of topical antibiotic. Topical antiseptics such as chlorhexidine and povidone-iodine solutions may be helpful, but ototoxicity is an issue, particularly in cases in which the tympanum is ruptured or cannot be evaluated.

Re-evaluation of the pet is important for monitoring response to therapy. Evaluation of the ear canal for progressive secondary changes and cytologic examination will allow alterations in therapy as needed. Significant narrowing of the ear canal is an indication for surgical intervention. Yeast overgrowth may occur with aggressive medical management of bacterial otitis and should be identified to maintain proper medical management.

Refractory or Recurrent Yeast Infection

Malassezia infection is a common perpetuating factor with erythematous ceruminous otitis and alterations in the otic microenvironment. Primary causes of the otitis should be identified and treated. Cytologic examination, not culture, should be relied upon for the diagnosis of yeast infection. If a case becomes refractory to therapy, reassessment of the primary condition and perpetuating factors should be done. Miconazole, clotrimazole, cuprimyxin, nystatin, and amphotericin B have all been described for treating Malassezia otitis. Climbazole had better in vitro activity against isolates of Malassezia pachydermatis in one study. Yeast were more susceptible to azole antifungals than polyene antifungals; however, oral ketoconazole, itraconazole, or fluconazole have been recommended for refractory cases. Long-term therapy may require topical antibacterial and antifungal combinations.

Ear cleaning may aid in the elimination of yeast organisms by removing cerumen, debris, or exudate and altering the microenvironment of the ear canal. Cleaning with antimicrobial agents such as chlorhexidine, povidone-iodine, and acetic acid may be beneficial; but as always the integrity of the tympanum should be established prior to use. Ear cleaning solutions may also have some efficacy against yeast organisms both in vitro and in clinical cases of otitis.


Chronic otitis externa may be the result of otic neoplasia, or otitis may be a predisposing factor in the development of neoplasia. Cocker spaniels are over-represented for benign and malignant neoplasia and otitis extema. Tumors of the skin and adenexal structures of the ear predominate. Benign tumors in dogs include sebaceous gland adenoma, basal cell tumor, polyp, ceruminous gland adenoma, and papilloma. Cats are more frequendy diagnosed with malignant neoplasms, but benign conditions include inflammatory polyps, ceruminous gland adenomas, ceruminous gland cysts, and basal cell tumors. Malignant neoplasms in both species include ceruminous gland adenocarcinoma, undifferentiated carcinoma, and squamous cell carcinoma. Ceruminous gland adenocarcinomas are the most frequendy diagnosed tumors of the ear canal in dogs and cats; however, one report stated that squamous cell carcinoma occurs with equal incidence in the cat.

The biologic behavior of otic tumors cannot be judged by their gross appearance; however, benign masses are usually nodular and pedunculated. Ulceration can be secondary to otitis associated with mass lesions, but malignant masses ulcerate more frequendy than benign masses. The tympanic bulla is involved in up to 25% of aural neoplasms, and neurologic signs occur in 10% of dogs and 25% of cats with otic neoplasia. The biologic behavior of malignant neoplasms tends to be local invasion with a low metastatic rate (e.g. 10% in dogs) to draining lymph nodes or lung.

Surgery is the mainstay treatment of otic neoplasia. Conservative excision may be possible for benign lesions, depending on the location of the tumor. Malignancies should be removed by total ear canal ablation and lateral bulla osteotomy. Incomplete excision results in recurrence of the mass and secondary otitis externa. Malignant neoplasia is associated with a median survival time (MST) of more than 58 months in dogs and 11.7 months in cats. Extensive tumor involvement and lack of aggressive management are associated with a poor prognosis in dogs. In cats a poor prognosis is associated with neurologic signs, squamous cell carcinoma or undifferentiated carcinoma, vascular or lymphatic invasion, and lack of aggressive therapy. Ceruminous gland adenocarcinoma has a median disease free interval of more than 36 months and 42 months in dogs and cats, respectively. The MST associated with squamous cell carcinoma and undifferentiated carcinoma in cats is 4 to 6 months.


Diseases Of The Middle And Inner Ear

Normal Anatomy and Physiology

The middle ear consists of the tympanic membrane, three cavities (epitympanic, tympanic, and ventral), and the bony ossicles (malleus, incus, and stapes). The tympanic membrane has two parts: (1) the thin pars tensa that attaches to the manubrium of the malleus and (2), above the pars tensa, the thicker, pars flaccida. The main portion of the middle ear, the ventral tympanic bulla, has two compartments in the cat (ventromedial and dorsolateral). The air-filled bulla is lined with modified respiratory epithelium, which is either squamous or cuboidal and may be ciliated. The four openings in the middle ear are the (1) tympanic opening, (2) the vestibu-lar window, (3) the cochlear window, and (4) the ostium of the auditory tube. The auditory tube is the communication between the middle ear and caudal nasopharynx. The normal flora of the middle ear may be due to this pharyngeal communication, but the role of the auditory tube as a source of bacteria in otitis media is unknown. The tympanic opening is a common source of bacterial infection of the middle ear in dogs with otitis extema. The cochlear and vestibular windows are possible ports of entry for progression of otitis media or ototoxic substances into the inner ear.

Cranial nerve VII, or the facial nerve, the sympathetic innervation of the eye, and the parasympathetic innervation of the lacrimal gland are closely associated with the middle ear. The separation of the facial nerve from the middle ear is minimal along the rostral aspect of its course through the petrosal bone. The nerve supplies motor fibers to the superficial muscles of the head, the muscles of the external ear, the caudal belly of the digastricus, and the ossicular muscles. The nerve also supplies sensation of the vertical ear canal and concave surface of the pinna.

Postganglionic sympathetic nerve fibers course closely with those of the facial nerve to innervate the smooth muscles of the eye. Preganglionic parasympathetic fibers also pass through the middle ear to innervate the salivary and lacrimal glands.

The inner ear is located within the petrosal bone. The cochlea, vestibule (saccule and utricle), and semicircular canals form the membranous labyrinth, which is encased in bone, called the bony labyrinth. The vestibular system functions to maintain the position of the eyes, trunk, and limbs relative to the position of the head, responding to linear and rotational acceleration and tilting. The system consists of the saccule, utriculus, and semicircular canals and communicates with the middle ear via the vestibular window. Fluid within the semicircular canals tends to remain stationary during motion, bending the cilia of the cells in the utricle and saccule, causing depolarization. These stereocilia synapse with the dendrites of the vestibular portion of the eighth cranial nerve and the signal is conducted via cranial nerve VIII to vestibular nuclei in the myelencephalon, the spinal cord, centers in the cerebellum and cerebral cortex, and motor nuclei of cranial nerves III, IV, and VI. The result is coordination of the body, head, and eye movement. Projections to the vomiting centers are responsible for nausea and vomiting associated with vestibular disorders and motion. The cochlear system, involved with the translation of sound, consists of the spiral organ, or organ of Corti, cochlear duct, scala vestibule, and scala tympani. Transmission of sound through the tympanic membrane, ossicles, and cochlear window results in undulation of the basilar membrane of the spiral organ. Cilia bend and cause depolarization and transmission of a signal to cochlear nuclei, caudal colliculi, and cerebral cortex. The cochlear nuclei control reflex regulation of sound via projections to cranial nerves V and VII, which control the muscles of the ossicles. Other projections allow for conscious perception of sound.

Otitis Media

Neoplasia of the Middle Ear

Neoplasia of the middle ear is rare; most cases represent extension of tumors originating in the external ear canal.

Inflammatory Polyps

Inflammatory polyps are a non-neoplastic admixture of inflammatory and epithelial cells originating in the tympanic bulla in cats. Other sites of origin include the auditory tube and nasopharynx. Macrophages, neutrophils, lymphocytes, plasma cells, and epithelial cells are usually present on histopathologic examination. The cause is unknown, but ascending infection and congenital causes have been suggested. No age or sex predilection exists for the condition, but younger cats are more commonly affected (1 to 5 years of age). Signs can be unilateral or bilateral and depend on the location of the mass lesion. A single polyp can grow into the external ear canal, down the auditory tube into the nasopharynx, or both. Signs of concurrent otitis extema and media are common with polyps limited to the ear, but respiratory stridor, dyspnea, gagging, and dysphagia occur with growth into the pharynx.

Diagnosis is based on otoscopic and pharyngeal examinations. Radiographs of the bulla, nasal cavity, and pharynx may be considered, and CT or MRI can be used to diagnose the site and side of origin of inflammatory polyps/ Treatment consists of excision by traction or surgical excision via ventral bulla osteotomy. Regrowth is a problem in half of the cats treated by traction extraction alone, and Homer’s syndrome is common in cats after ventral bulla osteotomy.

Otitis Interna

Otitis interna is usually an extension of otitis media or neoplasia of the middle ear. A careful neurologic examination is imperative to the localization of vestibular signs. Clinical signs associated with otitis interna include head tilt, ataxia, horizontal or rotary nystagmus, circling or falling toward the side of the lesion, or ipsilateral nystagmus. The fast phase of nystagmus is usually away from the side of the lesion. Occasionally, animals will become nauseated or vomit. Homer’s syndrome or deficits in cranial nerve VII may accompany otitis media interna, but involvement of other cranial nerves, vertical or changing nystagmus, or the presence of conscious proprioceptive deficits or paresis indicate central rather than peripheral vestibular disease. Bilateral peripheral vestibular disease is rare, but the animal will not have a head tilt, nystagmus, or strabismus and may exhibit wide head excursions and a crouched stance or the inability to stand.

The diagnosis of otitis interna is based on history, clinical signs, and physical, neurological, and otoscopic examinations. Advanced imaging may be helpful in distinguishing the anatomic location of the disease process. Treatment with aggressive medical or surgical intervention appropriate to the localization is important in prevention of adjacent brain stem involvement.

Prognosis for Otitis Media and Interna

A fair prognosis can be given if aggressive surgical and medical therapy are possible. Cases with concurrent severe external ear canal changes require total ear canal ablation and lateral bulla osteotomy. Repeated infections after ventral bulla osteotomy or total ear canal ablation and lateral bulla osteotomy may be operated again with resolution of the condition. Resistant organisms, failure to respond to aggressive surgery, and significant osteomyelitis are associated with a poor prognosis. The neurologic signs associated with otitis media and interna may be permanent, but many animals learn to use visual cues and can compensate for vestibular deficits. Facial nerve deficits, Horner’s syndrome, and keratoconjunctivitis sicca are often permanent.


Ototoxic substances (Table Ototoxic Drugs) damage the cochlear or vestibular systems or both. Otic application of medication can also cause adverse effects through local inflammation of the tympanic membrane or the meatal window (or both), as well as resultant otitis media. Topical medications also cause adverse effects by systemic absorption. Ototoxic substances reach the inner ear after local application and absorption through the cochlear or vestibular windows or hematogenously. The most frequent cause of ototoxicity is the application of an ototoxic substance to the external ear canal in a pet with a ruptured tympanum, which results in distribution to the middle ear. Absorption by the inner ear is increased when inflammation of the cochlear window occurs with otitis media. Hematogenous distribution of otoioxins to the inner ear is inherent in some medications (e.g. aminoglycosides).

Ototoxic Drugs

Aminoglycoside Antibiotics Antiseptics
Neomycin Chlorhexidine
Dihydrostreptomycin Iodine & iodophores
Gentamicin Ethanol
Streptomycin Benzalkonium chloride
Kanamycin Benzethonium chloride
Tobramycin Cantrimide
Amikacin Antineoplastic Agents
Other Antibiotics Cisplatin
Polymixin B & E Nitrogen mustard
Minocycline Miscellaneous
Erythromycin Quinine
Chloramphenicol Solicylates
Vancomycin Propylene glycol
Loop Diuretics Detergents
Furosemide Arsenic
Bumetanide Lead
Ethacrynic acid Mercury

The development of ototoxicity also depends on the vehicle of the preparation, chemical composition, drug concentration, concurrent medications, as well as the route, frequency, and duration of administration. Examples of increased risk of ototoxicity depending on the vehicle (e.g. combination of chlorhexidine and detergents) and concurrent medications (e.g. loop diuretics and aminoglycosides) have been described. Minimization of the risk of toxicity should be considered when any potentially toxic substance is administered either topically or systemically. The integrity of the tympanic membrane should be known prior to topical administration of any potentially ototoxic drug, and consequences of each drug should be considered in light of the animal’s health and concurrent therapies.

Idiopathic Vestibular and Facial Nerve Diseases

A complete neurologic examination is key to differentiating peripheral from central vestibular disorders. Head tilt, ataxia, horizontal or rotary nystagmus, and cranial nerve VII deficits may be seen with either condition. Central vestibular disease causes paraparesis, conscious proprioceptive deficits, other cranial nerve abnormalities, and vertical or changing nystagmus. Middle ear neoplasia, otitis media interna, idiopathic vestibular syndrome, and congenital vestibular disorders result in peripheral vestibular signs. Congenital vestibular disorders have been described in the German shepherd, Doberman pinscher, English cocker spaniel, Siamese, and Burmese breeds. Bilateral congenital vestibular syndrome has been described in beagles and Akitas. Clinical signs of head tilt and ataxia in these dogs and cats may be persistent or may improve; animals can be congenitally deaf.

Otitis media interna may be associated with facial paresis or paralysis if cranial nerve VII is affected by the inflammation. Otitis should be ruled out before diagnosing any animal with idiopathic facial nerve paralysis, because otitis requires aggressive management and the idiopathic condition can only be treated symptomatically or with acupuncture.


Acquired Late-Onset Conductive Deafness

Conductive deafness is due to lack of transmission of sound through the tympanic membrane and ossicles to the inner ear. Conditions that block sound transmission through the external ear canal, tympanic membrane, or middle ear and ossicles, such as otitis externa, otitis media, and otic neoplasia, cause conductive deafness. Less common causes of conductive deafness include trauma-induced fluid accumulation in the middle ear, atresia of the tympanum or ossicles, fused ossicles, or incomplete development of the external ear canal, which results in fluid accumulation in the middle ear. An increase in hearing threshold, absence of air-conducted hearing, and the presence of bone-conducted hearing on BAER suggest conductive deafness. The application of a bone-anchored hearing aid was described in one dog with conductive deafness after total ear canal ablation. It maintained bone-conducted hearing and tolerated the hearing aid anchored to the parietal bone Use of a bone-anchored device was required, because the dog did not have an external ear canal in which to place an earpiece. The hearing aid acted as an amplifier, and the dog seemed to respond to its use.

Acquired Late-Onset Sensorineural Deafness

Presbycusis, or decline in hearing associated with aging, may be due to one of the following: loss of hair cells and degeneration in the organ of Corti, degeneration of spiral ganglion cells or neural fibers of the cochlear nerve, atrophy of the stria vascularis, or changes in the basilar membrane. Because this condition occurs in older dogs and cats from 8 to 17 years of age, animals should be evaluated for concurrent causes of conductive deafness such as chronic otitis extema or media and otic neoplasia. BAER testing may demonstrate normal waveforms in response to high-intensity sound. If conduction is intact at an increased hearing threshold, use of an amplifying hearing aid may be beneficial. Pets may not tolerate occlusive types of ear pieces often used in hearing aids, and training to the ear piece should be done prior to application of the hearing aid.

Ototoxic substances, chronic exposure to loud noise, hypothyroidism, trauma, and bony neoplasia can also cause acquired late-onset deafness in dogs and cats. Ototoxicity can result in abolition of waveforms or an increase in hearing threshold on BAER. BAER testing can be used to re-evaluate patients for return of function after withdrawal of medication after exposure to ototoxic medication.

Congenital Sensorineural Deafness

Inherited sensorineural deafness usually results in complete loss of hearing in the affected ear by 5 weeks of age. Many breeds can be affected with the condition (Box Canine Breeds Associated with Inherited Deafness). The condition has been linked to coat color in many breeds of dogs and white cats. The condition is common in white cats, and mode of inheritance is thought to be autosomal dominant with incomplete penetrance.The condition is most common in white cats with blue irides. The correlation of white coat, blue eyes, and deafness is not perfect, but cats with two blue irides have a greater risk of deafness than cats with one blue iris, which have a greater risk of deafness than cats without blue irides. Total hearing loss occurs more often in longhaired white cats. The condition is common in certain breeds of dogs, such as dalmatians, which have a nearly 30% incidence of deafness (combining unilateral and bilateral deafness).

Canine Breeds Associated with Inherited Deafness

Akita Ibizan hound
American-Canadian shepherd Italian greyhound
American cocker spaniel Jack Russell terrier
American Eskimo Kuvasz
American Staffordshire terrier Labrador retriever
Australian cattle dog Maltese
Australian shepherd Miniature pincer
Beagle Miniature poodle mongrel
Bichon frise Norwegian dunkerhound
Border collie Nova Scotia duck tolling retriever
Borzoi Old English sheepdog
Boston terrier Papillion
Boxer Pit bull terrier
Bulldog Pointer
Bull terrier Poodle (toy & miniature)
Catahoula leopard dog Puli
Chihuahua Rhodesian ridgeback
Chow chow Rottweiler
Collie Saint Bernard
Dachshund Schnauzer
Dalmatian Scottish terrier
Doberman pincer Sealyham terrier
Dogo Argentino Shetland sheepdog
English cocker spaniel Shropshire terrier
English setter Soft-coated Wheaton terrier
Foxhound Springer spaniel
Fox terrier Sussex spaniel
French bulldog Tibetan spaniel
German shepherd Tibetan terrier
Great Dane Walker American foxhound
Great Pyrenees West Highland white terrier
Greyhound Yorkshire terrier

The trait is associated with the dominant merle or dapple gene in collies, Shetland sheepdogs, Great Danes, and dachshunds. The incidence of deafness tends to increase with increasing amount of white in the coat, and dogs homozygous for the merle gene are usually deaf and may be solid white, blind, or sterile, The piebald or extreme piebald gene is associated with deafness in dalmatians, bull terriers, Great Pyrenees, Sealyham terriers, greyhounds, bulldogs, and beagles. Inheritance is thought to be autosomal recessive, but the trait may be polygenic.

Heterochromia irides and lack of retinal pigment are associated with white color in dogs and cats. Hearing loss may be associated with absence of pigment in the cochlear stria vascularis. Diminished blood supply and disorders of endolymph production, with changes in the chemical or mechanical properties of endolymph, lead to degeneration of the organ of Corti secondary to stria vascularis atrophy. Loss of hair cells and abnormalities of the cochlear duct, Reissner membrane, tectorial membrane, and internal spiral suicus are typical of cochleosaccular type of end-organ degeneration seen in these cases.,

Clinical signs of deafness may be recognized in puppies as young as 3 weeks of age by astute owners; definitive diagnosis of uni- or bilateral deafness is usually made by BAER testing at 5 to 6 weeks of age when the auditory system is completely developed and cochlear degeneration, if present, is complete.

Congenital Acquired Sensorineural Deafness Exposure to bacteria, ototoxic drugs, low oxygen tension, and trauma in utero or during the perinatal period rarely causes deafness in young animals.


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.


Normal Peripartum Procedures

The expected foaling date should be calculated as 11 months and 5 days. At approximately 30 days before foaling, booster vaccines should be given and the mare should be dewormed. If fescue toxicosis is problematic in the area, foaling mares should not be allowed to graze fescue pasture and should not be fed hay-containing fescue within 30 days of foaling. Domperidone may be given in a daily oral dose of 1.1 mg/kg, beginning 15 days before the anticipated foaling date if the threat of fescue toxicosis cannot be removed.

Weather permitting, foaling can take place outside in a grassy paddock. The mare should not be allowed to foal in the presence of other mares. If a foaling stall is used it must be prepared by thorough cleaning and disinfection. Cleaning is the first step and a detergent must be used to disperse the lipid biofilm layer that may protect some pathogens. After the stall has dried, an approved phenolic disinfectant should be used to soak all surfaces within the stall. Feed tubs and water buckets should be cleaned, rinsed, and allowed to dry before placing them back in service.

Then 2 weeks before foaling, the mare’s perineal area should be examined for conformation; the Caslick’s procedure should be opened if necessary. Signs of previous injury or tears may indicate that additional care is needed at parturition to avoid repeat foaling injuries.

On most large farms there will be a nighttime foaling attendant. This individual is trained to watch for the signs of stage 1 labor and then wrap the tail with gauze, tape, palpation sleeve, or some combination of these items, and wash the udder and perineal area with cotton soaked in a weak solution of water and povidone iodine scrub. A detergent is necessary in the scrub to break the lipid layer on the skin. The scrub should be rinsed off with water. After delivery, the umbilical cord is held close to the foal’s abdomen as the cord breaks, and the umbilical stump is immediately dipped in, or sprayed with, a navel dip solution. The application of navel dip solution is repeated again in 4 to 6 hours. Navel dip solutions such as 7% iodine are tissue destructive and should be avoided. Iodine-based solutions at 2% to 3.5%, povidone iodine solution at 2%, or chlorhexidine solution at 0.5% will reduce bacterial numbers without destroying tissue. Chlorhexidine scrub and povidone iodine scrubs have been used successfully as navel dips.

The newborn foal should be vigorously scrubbed with a towel to stimulate the foal’s movements and respirations. Then the mare and neonate should be observed from a distance for signs of normal or abnormal behavior. After the mare stands, the colostrum’s specific gravity should be tested to estimate quality. A specific gravity of 1.06 or greater is adequate. If the Eclipse refractometer is used a reading of 23% corresponds to a specific gravity of 1.06. It is good practice to collect a pint to freeze if quality is high. Colostrum with a specific gravity of 1.06 will have an approximate immunoglobulin (Ig) concentration of 3000 mg/dl. If the specific gravity of the colostrum is low, then 1 g of colostral IgG/kg of birth weight should be given by oral supplementation.

The soiled bedding should be removed and replaced with clean dry bedding. The mare and foal should still be observed for normal behavior. The foal should be in a sternal posture within 1 to 2 minutes, and a suckle reflex should be present within 2 to 20 minutes. Standing should occur within 2 hours, and the foal should nurse by 3 hours after birth. Once nursing has occurred the foal may be given a warm water or soap-based enema to facilitate passage of the meconium. Commercial phosphate enemas may be used, but repeated use should be avoided because of the absorption of the phosphate ion. If vital signs are taken, the foal’s temperature should fall in the range of 99° to 101.5° F (37.2°-38.6° C), the heart rate at 1 to 5 minutes should be greater than 60 beats per minute (bpm), and at 5 to 60 minutes it should be 80 to 130 bpm. The respiratory rate is high initially at 60 to 80 bpm in the first 30 minutes and then drops to 30 to 40 bpm within 1 to 12 hours after birth.


Management Of Thermal Burns

Skin is usually quite slow to absorb heat; as a result, its dissipation also takes much longer than might be expected. Therefore the skin is slow to burn, and the burning effects may continue for some time after the cause has been removed. Furthermore, burn injuries heal very slowly, even in sites where healing is normally good. Therefore the attending veterinarian and the owner must be willing and able to embark upon the process of healing (). The long course inevitably also means a high cost.

All burns should be treated immediately by the application of cold running water, which should be applied for at least 15 minutes. This will reduce the heat retention and limit the consequent necrosis. A protective, water-soluble emollient antibacterial cream should be applied to the burn area. Oil- or fat-based ointments should be avoided. Silver sulfadiazine (Silvadene) is a very useful topical medication for superficial or partial thickness burns over limited areas.

The metabolic status of the horse must be carefully assessed and supportive measures applied as necessary. These include fluid therapy and, if necessary, an immediate plasma transfusion to ensure that shock is controlled. The loss of plasma is maximal in the first 12 to 24 hours. In all cases, large volumes of intravenous fluid therapy are a useful first emergency measure. This applies particularly to the more severe burns (full-thickness over 5% or more of the body). If the animal is already in a state of shock, aggressive antishock therapy must be instituted immediately. This course of action may include large volumes of balanced electrolyte solutions (e.g., Hartman’s solution or lactated Ringer’s or even hypertonic [7%] saline). A helpful rule of thumb is that for each percentage body surface involved, 3 to 4 ml/kg body weight should be administered. Fluids should not be sustained if hydration is adequate because it might add to the secondary edema, particularly within the lungs.

Plasma protein estimation may identify falling total protein with albumin loss. Good-quality fresh or preserved plasma will be useful; 1 L of plasma will raise the total protein by about 0.2 g/L in a 450-kg horse. The total volume of plasma required for severe burn cases can be up to 40 L or more over 2 to 3 days. This can usefully be sustained at a low rate for as long as required.

Pain relief in the form of flunixin meglumine (1 mg/kg IV q24h) or phenylbutazone (2 mg/kg ql2h) is essential. Pain is often more severe in the more superficial types of burn. Full-thickness burns may not be very painful, but the metabolic consequences of this type of burn are usually more severe.

The topical use of dilute chlorhexidine solution in saline is controversial. Controlling infection is clearly meritorious but may be better performed simply by irrigation. However, burns are particularly liable to infection. Therefore topical water-soluble antibiotic creams may be advisable. Blisters should be left alone for at least 36 to 48 hours. No merit exists in trying to burst or drain these blisters.

Bandaging may be possible in some cases. In such a case, hydrogel should be applied liberally to the site. Burns are usually highly exudative because of the deficit in skin and the loss of cutaneous lipid. Thus an absorbent dressing should be applied. Bandages must be firm enough to provide support without slippage but loose enough as to limit any further vascular compromise. The dressing must not stick to the wound site.

Full-thickness burns must be covered immediately with a protective fluid proof dressing. In an emergency a clear plastic kitchen wrapping can be useful. Ideally, a hydro-gel should also be applied directly to the wound site from the outset.

After 24 to 36 hours, the wound can be cleaned and all damaged tissue removed. The clinician should expect further necrosis to develop over the following few days or weeks. At this stage, all hair in the affected area can usefully be clipped. This prevents matting and reduces pain.

Dressings should be replaced frequently over the first few days, and any necrotic tissue should be removed. A calcium alginate dressing (e.g., Algiderm) is a useful absorptive dressing that will maintain moist healing conditions at the site. Once healing is underway, the dressings can be left for up to 3 to 4 days, provided that no complications develop. An eschar should be left in situ until natural sloughing occurs. While it remains in situ it provides an effective biologic cover and protection for the underlying tissues. An overlying moist wound dressing (e.g., a hydrogel [Tegagel]) can sometimes reduce the time taken for the eschar to separate and may also encourage contraction of the healing edges of the wound.

The metabolic status of the horse must be regularly assessed and must include full hematologic profiles for protein analysis and electrolyte status. Hyperkalemia is a common consequence, and seriously burnt horses may show hemoglobinuria. Progressive anemia is a serious potential effect of extensive burns and requires attention as soon as it is recognized. Usually it is caused by a combination of intravascular hemolysis and bone marrow suppression.

The nutritional status of the horse is critical to its recovery. Almost all serious burn cases are in a negative protein balance (i.e., they are losing more than they are absorbing) and have a very much raised energy requirement. The early stages may require parenteral nutritional support, but if the horse will eat, gradual addition of vegetable oil into a high-quality ration (possibly of alfalfa hay) can be helpful.

In cases of extensive skin deficits, skin grafting should be considered at an early stage — as soon as the granulation tissue is healthy enough to accept a graft. Grafting is unhelpful until all necrotic tissue is removed and the bed of granulation tissue is healthy. Split-thickness, mesh grafts in single sheets or postage-stamp format can be used effectively. Cosmetic grafting with extension flaps, tube grafts, and so on are all techniques that can be applied to the healing of burn injuries of all types. Artificial skin substitutes (e.g., Integra) may be used to protect the exposed tissues and reduce the extent of plasma exudation.


Complications Of Burns

Infection is a serious and frequent complication of burns and must be addressed at an early stage. For the most part, normal skin commensal organisms such as Streptococcus equi var. zooepidemicus, Staphylococcus aureus, and Pseudomonas aeruginosa are encountered with some complicated by other gram-negative species, such as E. coli and Clostridia spp., and yeasts can be found. Silver sulfadiazine (Silvadene) is a useful broad antibacterial that has little or no harmful effects on wound healing.

Some horses suffer from renal shutdown after sustaining a severe burn and renal function must be encouraged and repeatedly checked. Diuretics such as furosemide are often indicated but should be used with considerable care.

Smoke inhalation or internal burns can cause serious pulmonary edema and thus must be controlled. Oxygen supplied directly to the trachea or nasally may be helpful. A single intravenous dose of dexamethasone (0.5 mg/kg) may assist. Intravenous administration of dimethyl sulfoxide (DMSO) at 1 g/kg over the first 2 days may be helpful in reducing the pulmonary edema. All cases in which smoke inhalation has occurred must have systemic antibiotic therapy because the respiratory tract is particularly susceptible to serious infection after inhalation damage. Obtaining a transtracheal aspirate for culture if the chosen antibiotics do not appear to be helping is justifiable. Fungal infections pose a particularly serious threat that may be untreatable.

Cornea] and eyelid damage is particularly dangerous because of the delicate nature of the tissue and their intolerance to injury. In cases in which the face has been involved in the burn (to any extent at all) the corneas should be medicated carefully with artificial tears. In all cases the cornea should be stained with fluorescein to check for ulceration and necrotic tissue. All necrotic tissue should be gently removed with a saline-soaked cotton swab. Under no circumstances should corticosteroids or any strong chemicals such as chlorhexidine or povidone iodine be applied to the eye. Topical antibiotics (e.g., triple antibiotic or gentamicin) should be applied with atropine to control any reflex uveitis. If the eyelids are involved or are suspected to be involved, then particular care must be taken to protect the corneas with artificial tears (applied every hour), and, if necessary, a third eyelid flap can be drawn over the eye to afford sustained protection.

Healing of burn sites is reported to be slower than other types of wounds. This is possibly because the full extent of the injury is not apparent from the outset; furthermore, the damaged tissue is usually slow to separate from the healthy underlying structures. Scarring is inevitable and can be either functionally limiting (e.g., the eyelids or over joints), cosmetically unacceptable, or both. Most serious burn cases have degrees of immunosuppression, which renders them liable to infection and delayed wound healing.

Healing burn sites are often pruritic, and self-inflicted damage can be severe. Suitable sedation may be required (usually acepromazine is effective) to prevent self-inflicted trauma. Cross-tying, neck cradles, or muzzles can also be useful. These measures will require extra nursing observation.

Other complications from burns include colic (usually an impaction) or laminitis. Inappetence or failure to drink are serious potential complications and must be managed early. Fresh green grass is usually a good stimulant to appetite and also provides significant water intake. Caustic burns can result in absorption of the caustic material; thus serious systemic effects may occur.



Sporotrichosis: Etiology

Sporotrichosis is a mycotic disease caused by the dimorphic fungus Sporothrix schenckii. S. schenckii exists in a mycelial form at environmental temperatures (25°-30° C) and as a yeast form in body tissues (37° C). The organism is distributed worldwide and can be found preferentially in soils that are rich in decaying organic matter. It has also been isolated from barberry and rose bush thorns, spaghnum moss, tree bark, and mine timbers. The handling of stored hay bales has also been associated with outbreaks. The traditionally accepted method of acquiring sporotrichosis is via the inoculation of the infectious organism into tissues. The disease in horses is often associated with a traumatic puncture wound on the distal extremity from a thorn, wood splinter, or barbed wire. Although contamination of a puncture wound by organisms in the environment is considered an important mechanism in acquiring this disease in people as well as horses, contact exposure to cats (especially barn cats) infected with S. schenckii is now considered a significant means by which a zoonotic infection can be established. It has also been shown that the claw of the cat may be contaminated with S. schenckii, and therefore a direct puncture wound from a claw of a cat needs to be considered a potential source of infection.

Sporotrichosis: Clinical Signs

Sporotrichosis can occur in three clinical forms: cutaneous, cutaneolymphatic, and disseminated. In horses, the cutaneolymphatic form is the most common. In most cases, the infectious organism is first inoculated into the dermis or subcutaneous tissue via a traumatic injury. The distal limbs are the most common site for the development of a cutaneous nodule or several nodules. This process extends proximally up the limb and follows the lymphatics, thus resulting in the formation of additional nodules and a “cording” of the lymphatics. Several nodules often become ulcerated and drain a purulent to hemopurulent discharge. In the more chronic cases the ulcerated nodules can develop excessive granulation tissue and take on a “proud flesh” appearance. The proximal draining lymph node may be palpably enlarged and may subsequently ulcerate and drain. Less commonly, the primary cutaneous form of sporotrichosis may be single or multiple intact or draining nodules that develop on the trunk, shoulder, hip, perineum, or face. To this author’s knowledge, the disseminated form of sporotrichosis has not yet been reported in the horse.

Differential Diagnosis of Sporotrichosis

The initial differential diagnosis for the formation of a cutaneous nodule on the distal extremity should include both bacterial (especially Corynebacterium sp., and Staphylococcus sp.) and fungal (especially blastomycosis, coccidioidomycosis, cryptococcosis, and histoplasmosis) etiologies. These differentials will vary greatly by regional differences in the incidence of these infections. It is important to obtain the horse’s geographic history for at least the past year of travel. However, in the more advanced forms of these diseases, it is very rare for the lymphatics to become “corded” proximally with the formation of additional ascending individual nodules. When individual nodules on the distal extremity become ulcera-tive, the differential diagnosis should additionally include squamous cell carcinoma, cutaneous habronemiasis (“summer sores”), exuberant granulation tissue (“proud flesh”), and the fibroblastic sarcoid.

Diagnosis of Sporotrichosis

The most reliable method for confirming the diagnosis of sporotrichosis is to biopsy an intact and nonulcerated nodule and to submit a portion for both histopathologic examination and macerated tissue culture. An 8-mm biopsy punch will give a suitable sample, and the resulting core of tissue can be cut in half longitudinally. Half of the sample should be submitted for histopathologic examination and the other half for culture. It should be requested that the tissue be macerated and cultured for both bacterial and fungal organisms as confirmed cases of sporotrichosis may have a concurrent bacterial component. In instances where only ulcerated and draining nodules occur, the tissue should still be biopsied and sampled as above. This is because both culture and cytologic examination of stained exudate from draining lesions is often negative for the presence of Sporothrix organisms. Some cases of equine sporotrichosis are initially negative on macerated tissue culture. In such cases, the histopathology report indicates the presence of a deep pyogranulomatous inflammatory reaction without the presence of any infectious agent upon examination with special fungal stains. In instances in which the lesions continue to persist and do not respond to treatment for any of the bacterial pathogens that are isolated, a portion of the biopsied tissue from a nodule should be submitted to the Centers for Disease Control and Prevention (CDC) laboratory in Atlanta, Ga., for the fluorescent antibody testing (Sporothrix antigen-specific direct immunofluorescent antibody test). This procedure is considered the most sensitive test for determining the presence of Sporothrix organisms.

Treatment of Sporotrichosis

The treatment of choice for the cutaneolymphatic or primary cutaneous form of sporotrichosis is systemic iodide therapy. The organic iodides have proven superior in efficacy to the inorganic iodides in the treatment of equine sporotrichosis; ethylene diamine dihydroiodide (EDDI Equine) is the drug of choice. This product is in the form of a feed additive and can be mixed with a small amount of grain and administered at a dosage of 1 to 2 mg/kg of the active ingredient given once to twice daily for the first week. The dosage can then be reduced to 0.5 to 1.0 mg/kg once daily for the remainder of the treatment. In general, lesions will begin to regress during the first month of treatment, and treatment should be continued for at least 1 month beyond the complete resolution of all cutaneous nodules and the healing of any ulcerated lesions. Discontinuing therapy prematurely will invariably result in an unnecessary relapse of the disease.

During treatment, the horse should be closely observed for any evidence of iodide toxicity (iodism), which includes excess scaling and alopecia, serous ocular or nasal discharge, excess salivation, anorexia, depression, coughing, nervousness, or cardiovascular abnormalities. Should any of these signs develop, the treatment should be discontinued for 1 week, and the treatment should be resumed at 75% of the dosage at which the iodism was noted. In most instances, the treatment is subsequently well tolerated. In the rare instances in which iodism is a recurrent problem or the horse fails to respond to treatment with organic iodide, griseofulvin therapy may be used. Griseofulvin has been reported to be effective when administered at a dosage of 20 mg/kg given orally once daily for the first 2 weeks and is then followed by 10 mg/kg given orally once daily for 1 month beyond apparent clinical remission. Itraconazole has also been suggested as an alternative treatment for refractory cases of sporotrichosis. The recommended dosage is 3 mg/kg twice daily mixed with grain, but its expense would be a restriction to its use in most cases.

Public Health Significance

It is important to remember that the accidental inoculation or contamination of broken skin from any contaminated tissues or exudates may infect any person who comes in contact with an animal infected with Sporothrix organisms. Several reports have documented the transmission of sporotrichosis to people by contact with a contaminated wound or the exudate from an infected cat.

In some instances, infection has occurred after exposure to an infected cat, although no known preexisting injury or penetrating wound on the person was known before the disease presented. With these considerations in mind, it is advisable that people who handle horses suspected of having sporotrichosis wear disposable gloves. Afterward, they should remove the gloves carefully and wash their forearms, wrists, and hands with either a chlorhexidine or povidone-iodine scrub.


Treatment of Pastern Dermatitis

The appropriate therapy obviously involves identification of the predisposing, perpetuating, and primary factors. In general, avoiding pastures/paddocks with mud, water, or sand may minimize predisposing factors. Keeping patients stalled during wet weather and until morning dew has dried is often rewarding. Use of alternate sources of bedding may be beneficial because the chemicals in treated or aromatic types of wood shavings may result in contact dermatitis. Lastly, clip hairs — especially feathers — to avoid moisture retention.

Perpetuating factors should be addressed according to the severity of the condition. The most conservative approach includes cleansing lesions with antimicrobial shampoos (benzoyl peroxides, chlorhexidine, ethyl lactate, imidazoles) twice daily for 7 to 10 days and then tapering in frequency. If a dry environment is not possible, the affected pastern areas can be protected with ointments (creating a moisture barrier); with padded and water-repellent bandages (changed q24-48h); or with Facilitator, a hydroxy-ethylated amylopectin liquid bandage that is replenished every 1 to 3 days. If the lesions are exudative, astringent solutions — such as lime sulfur (LymDyp), aluminum acetate solutions, black tea bag or sauerkraut poultices, or acetic acid/boric acid wipes (Malacetic Wipes, Dermapet Inc., West Plains, Mo.) — should be used after cleansing.

Topical sprays, creams, or ointments that contain antibiotics, steroids, antifungal agents, or a combination thereof may benefit the patient, depending on the diagnosis. A 2% mupirocin ointment (Bactoderm), with excellent tissue penetration, is the author’s preference for addressing localized dermatophilosis and bacterial dermatitis. A DMSO / thiabendazole / sulfa ointment has also been described in the fourth edition of Current Therapy in Equine Medicine. If generalized to all four limbs, treatment of the bacterial dermatitis is best accomplished with daily systemic antibiotics (trimethoprim/sulfa 30 mg/kg/day or cephalexin 22 mg/kg q8hrs) until 7 days after clinical resolution.

Lime sulfur dips and chlorhexidine / imidazole-containing shampoos, sprays, and residual leave-on products comprise the current antifungal arsenal in veterinary medicine. Topical enilconazole (Imaverol), labeled for use in horses in various countries other than the United States, has been used to treat fungal infections with reported success. Many veterinary dermatologists feel that systemic griseofulvin lacks efficacy for the treatment of equine dermatophytosis.

Ectoparasiticidal therapy consists of avermectins, topical organophosphates (malathion, coumaphos), pyrethroids (permethrin, flumethrin), lime sulfur, and fipronil (Frontline). The latter has had recent success in the treatment of Chorioptes bovis within a group of heavier cob and draught-cross horses. Of note was the ability of the parasite to survive off the host, enduring solely in the presence of skin debris in a moist and dark environment and thus emphasizing the need for environmental management to prevent recurrence.

Immunomodulators have been used for the condition. Interferon-a2a given at 1000 IU/ml on a cycle of 1.0 ml per horse daily for 3 weeks and then off for 1 week has been used by the author to help stimulate the local immune defense system, with very little cost or side effects. Immune-mediated conditions such as PLV, however, require a significant immunosuppressive effort to achieve resolution and control of the clinical signs. High-dose glucocorticoids, preferably dexamethasone (0.1-0.2 mg/kg q24h for 7-14 days, then taper over the next 4-6 weeks), along with reduction of UV light exposure by stabling or covering with a light bandage, appears to control — if not resolve-many cases. Should resolution of clinical signs not be achieved by 14 days, the author has achieved excellent results by adding pentoxifylline (PTX), a phosphodiesterase inhibitor. PTX has been reported to have multiple immunomodulatory effects that potentiate the effectiveness of traditional immunosuppressive drugs (i.e., steroid-sparing effect). These include inhibition of lymphocyte activation and proliferation; increased lymphocyte suppression; suppression of tumor necrosis factor (TNF)-a, lymphotoxin, and interferon-7 production; and upregulation of IL-10 mRNA that leads to increased IL-10 serum levels. Oral absorption varies considerably between individuals; thus reported dosages range between 4 to 8 mg/kg every 12 hours.

Once the skin has returned to normal, long-term control of PLV may be achieved by a combination of topical steroids (betamethasone valerate 0.1%, aclometasone 0.05%), coupled with an every other day systemic regimen of PTX and, if necessary, low-dose dexamethasone on an alternate day basis.

The prognosis and healing time of equine pastern dermatitis depends on the stage of disease when treatment begins and on the ability to identify the etiology. Ensuring that predisposing, primary, and perpetuating factors are encompassed in a diagnostic and treatment plan will optimize the likelihood of a positive outcome.