Otitis Media

Otitis media may result from extension of otitis extema through the tympanic membrane, aspiration of pharyngeal contents up the auditory tube (e.g. a sequela to upper respiratory tract (URT] infection in cats), or from hematogenous spread. Extension from otitis externa is the most common cause of otitis media, but otitis media may serve as a perpetuating factor for otitis extema. Developmental abnormalities of the external ear canal and pharynx may also result in otitis media due to the accumulation of secretions in the middle ear. Neoplasia, inflammatory polyps, and middle ear trauma may be associated with secondary otitis media or result in similar clinical signs.

Cholesteatoma is commonly associated with otitis media and chronic otitis extema. A cholesteatoma is a mass of keratinized squamous cells that accumulate within a structure lined with stratified squamous epithelium. The lesion is presumed to develop when a pocket of tympanic membrane becomes adhered to inflamed middle ear mucosa. Significant narrowing of the external ear canal is usually present. Radiographic signs of increased density and bony changes of the tympanic bulla predominate with loss of the air-filled lumen of the external ear canal and concurrent calcification. Treatment is usually limited to total ear canal ablation and lateral bulla osteotomy due to the changes of the external ear canal and mass or accumulation of debris in the tympanic bulla.

The clinical signs associated with middle ear disease often reflect concurrent otitis extema (e.g. head shaking, lethargy, exudate, otic malodor). Significant otic pain, lethargy, inappetence, and pain upon opening the mouth are more suggestive of middle ear involvement. Neurologic signs may be present due to the course of the facial nerve and sympathetic innervation of the eye. Facial nerve paresis or paralysis result in facial asymmetry (i.e. uneven position of the lip commissures, unequal ear carriage, unilateral ptyalism) and abnormal cranial nerve reflexes on neurologic examination (e.g. menace response, palpebral and corneal reflexes, abnormal ear canal, and concave pinnal sensation). Homer’s syndrome, or loss of sympathetic innervation to the eye, can also be complete or partial (i.e. ptosis, miosis, enophthalmia, prolapse of the third eyelid). Otitis interna is usually evidenced by head tilt, abnormal nystagmus, and ataxia and should be differentiated from central vestibular disease based on careful neurologic examination. Otitis interna is not usually associated with ipsilateral hemiparesis or abnormalities in level of consciousness.

Cases of paraaural abscessation usually have concurrent otitis media. The primary cause may be trauma to the external ear canal, severe otitis externa, extension of otic neoplasia, or total ear canal ablation. Signs of middle or external ear disease and soft tissue swelling in the parotid area may be accompanied by draining tracts. A head tilt and pain upon palpation of the area are usually present.

The diagnosis of otitis media is based on a thorough history and physical, neurologic, and otoscopic examinations. A ruptured tympanum strongly suggests otitis media. The pharynx should also be evaluated on physical examination; identification of specific conditions may require general anesthesia due to anatomic location (eg. inflammatory polyps) or pain associated with examination (e.g. otitis media causing temporomandibular joint (TMJ) pain, severe otitis externa). General anesthesia may also be required to perform a complete otoscopic examination in cases of severe otitis externa in which thorough cleaning of the ear is necessary for therapy and diagnosis (i.e. visualization of the tympanum). Significant otitis externa is commonly associated with otitis media; the tympanic membrane is ruptured in up to 50% of dogs with otitis externa, although 70% of dogs with otitis media had an intact tympanic membrane in one study. The tympanic membrane in dogs with otitis extema may be difficult to examine due to secondary changes of the external ear canal, pain associated with otoscopic examination, and accumulation of exudate, cerumen, and debris. Treatment to diminish the severity of otitis externa and general anesthesia may increase the ability to evaluate the tympanum in these cases.

Any case that has significant cerumen, exudate, or debris should undergo careful cleaning of the ear canal to allow evaluation of the integrity and character of the tympanic membrane The presence of a “false middle ear” occurs when large accumulations of debris lodge against the tympanic membrane, causing it to deviate medially into the middle ear. This makes the external ear canal appear elongated and leads to misdiagnosis of a ruptured tympanic membrane.

Gende probing of the tympanic membrane with a red rubber catheter under direct visualization may assist in the diagnosis of small tears in the membrane. If the catheter tip is consistently visible, rupture is unlikely. Alternatively, an aliquot of 1 mL of physiologic saline placed in the horizontal canal should remain stationary; disappearance suggests an opening in the tympanum, allowing the fluid to drain into the middle ear. Movement of the fluid may be blocked by large amounts of debris in the middle ear, even in the presence of a tear in the tympanum.

If the tympanic membrane is visible, its character should be recorded in the medical record for comparison upon re-evaluation. Bulging, increased opacity, and hyperemia may be present with otitis media. If otitis media is suspected, radiographs of the bullae may be made. Lateral oblique and open-mouth views are most helpful for evaluating the tympanic bulla, but positioning for comparison of left and right sides is difficult and requires general anesthesia. Ventrodorsal or dorsoventral views allow evaluation of the air-filled lumen and calcification of the external ear canal. Abnormalities of the bulla include increased opacity, sclerosis, and lysis. Fluid cannot be differentiated from increased soft tissue density (e.g. neoplasia), and absence of radiographic changes does not rule out otitis media. Radiographic changes were absent in 33% of the middle ears in one study of dogs with otitis media confirmed by surgical exploration. Otitis media or neoplasia and otitis interna can cause radiographic evidence of lysis of the petrosal bone.

Other diagnostic tools are available to evaluate patients with otitis media interna. Contrast introduced into the external ear canal followed by radiography, termed canalography, is used to diagnose tympanic membrane perforations. The method is useful for acute tympanic membrane rupture and increases the frequency of diagnosing tympanic membrane rupture with concurrent otitis externa and media beyond that of otoscope alone Advanced imaging with CT and MRI have been studied in normal dogs and dogs with otitis media. CT is considered superior to MRI for bony changes, whereas MRI is better for detection of soft tissue abnormalities in both dogs and cats.

If the tympanic membrane is intact in a dog with otitis media, a myringotomy is performed to obtain samples for culture and susceptibility testing and cytologic examination. Affected dogs are often more comfortable after collection of samples due to decreased pressure in the middle ear after myringotomy. The procedure must be performed with general anesthesia and is usually done after radiography or advanced imaging of the ear. The external ear canal should be thoroughly cleaned and dried prior to myringotomy to avoid contamination with external ear canal debris. Direct otoscopic visualization is used for the procedure. A 20-gauge spinal needle is used to penetrate the tympanic membrane through the caudoventral aspect of the pars tensa. Suction is applied and samples collected — culture and susceptibility takes priority over cytologic examination because cytology is frequently negative, and cultures of the external ear canal do not reflect the middle ear bacteria in the majority of cases. If fluid cannot be aspirated direcdy from the middle ear, 0.5 to 1 mL of warm, sterile saline can be infused through the needle into the middle ear cavity and aspirated. Alternatively, an open-ended tomcat catheter or small, sterile culture swab may be passed into the middle ear cautiously under otoscopic visualization. Pseudomonas species and Staphylococcus intermedium are most commonly isolated, followed by yeast, β-hemolytic Streptococcus, Corynebacterium species, Proteus species, and Enterococcus species. Surgical exploration is rarely required for the diagnosis of otitis media.

Medical therapy of otitis media should be guided by culture and susceptibility results. The external ear canal is flushed and dried as necessary to treat concurrent otitis extema. Flushing is usually performed under the same general anesthetic episode used for diagnostic testing. If the tympanic membrane is ruptured, the middle ear should be gendy lavaged with warm saline. Cytology results, when available, should be used to guide initial therapy. The integrity of the tympanic membrane must be considered when using topical agents to treat concurrent otitis extema: ototoxic medications and vehicles should be avoided if the tympanic membrane is ruptured.

Newly diagnosed cases of otitis media may be started on empiric therapy based on cytology. First-choice antimicrobials include cephalosporins, amoxicillin and clavulonic acid, and fluoroquinolones. Definitive therapy consists of administration of antibiotics based on culture and susceptibility results for a minimum of 4 to 6 weeks. Primary and perpetuating factors of otitis externa should be identified and treated or controlled. Topical medication and flushing of the external ear canal should continue until resolution of clinical signs and normalization of cytology. Gradual improvement of the otitis media is expected within 14 days. The ear canal and tympanic membrane should be evaluated prior to and after discontinuation of therapy. Small tears in the tympanic membrane after myringotomy heal rapidly with appropriate therapy within 2 to 3 weeks.Hs However, re-evaluation of the tympanic membrane in dogs with otitis externa media should precede alteration of the topical agents in the therapeutic plan.

Failure to respond to therapy or chronic or recurrent otitis media warrant re-evaluation for surgical intervention. Total ear canal ablation and lateral bulla osteotomy should be considered in cases with severe secondary changes of the external ear canal and concurrent otitis media. If the external ear canal is not affected, a ventral bulla osteotomy may be performed to remove gross exudate and establish drainage from the middle ear of dogs and cats with chronic or recurrent otitis media. Caution should be taken in considering lateral ear resection and ventral bulla osteotomy in the treatment of concurrent otitis externa and media because lateral ear resection is only an adjunct to medical management of otitis externa.


Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid (Clavamox, Augmentin)

Potentiated Aminopenicillin

Highlights Of Prescribing Information

Bactericidal aminopenicillin with beta-lactamase inhibitor that expands its spectrum. Not effective against Pseudomonas or Enterobacter

Most likely adverse effects are GI related, but hypersensitivity & other adverse effects rarely occur

What Is Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid Used For?

Amoxicillin/potassium clavulanate tablets and oral suspension products are approved for use in dogs and cats for the treatment of urinary tract, skin and soft tissue infections caused by susceptible organisms. It is also indicated for canine periodontal disease due to susceptible strains of bacteria.

Pharmacology / Actions

For information on the pharmacology/actions of amoxicillin, refer that monograph.

Clavulanic acid has only weak antibacterial activity when used alone and presently it is only available in fixed-dose combinations with either amoxicillin (oral) or ticarcillin (parenteral). Clavulanic acid acts by competitively and irreversibly binding to beta-lactamases, including types II, III, IV, and V, and penicillinases produced by Staphylococcus. Staphylococci that are resistant to penicillinase-resistant penicillins (e.g., oxacillin) are considered resistant to amoxicillin/potassium clavulanate, although susceptibility testing may indicate otherwise. Amoxicillin/potassium clavulanate is usually ineffective against type I cephalosporinases. These plasmid-mediated cephalosporinases are often produced by members of the family Enterobacteriaceae, particularly Pseudomonas aeruginosa. When combined with amoxicillin, there is little if any synergistic activity against organisms already susceptible to amoxicillin, but amoxicillin-resistant strains (due to beta-lactamase inactivation) may be covered.

When performing Kirby-Bauer susceptibility testing, the Augmenting (human-product trade name) disk is often used. Because the amoxicillinxlavulanic acid ratio of 2:1 in the susceptibility tests may not correspond to in vivo drug levels, susceptibility testing may not always accurately predict efficacy for this combination.


The pharmacokinetics of amoxicillin are presented in that drug’s monograph. There is no evidence to suggest that the addition of clavulanic acid significantly alters amoxicillin pharmacokinetics. Clavulanate potassium is relatively stable in the presence of gastric acid and is readily absorbed. In dogs, the absorption half-life is reportedly 0.39 hours with peak levels occurring about 1 hour after dosing. Specific bioavailability data for dogs or cats was not located.

Clavulanic acid has an apparent volume of distribution of 0.32 L/kg in dogs and is distributed (with amoxicillin) into the lungs, pleural fluid and peritoneal fluid. Low concentrations of both drugs are found in the saliva, sputum and CSF (uninflamed meninges). Higher concentrations in the CSF are expected when meninges are inflamed, but it is questionable whether therapeutic levels are attainable. Clavulanic acid is 13% bound to proteins in dog serum. The drug readily crosses the placenta but is not believed to be teratogenic. Clavulanic acid and amoxicillin are both found in milk in low concentrations.

Clavulanic acid is apparently extensively metabolized in the dog (and rat) primarily to l-amino-4-hydroxybutan-2-one. It is not known if this compound possesses any beta-lactamase inhibiting activity. The drug is also excreted unchanged in the urine via glomerular filtration. In dogs, 34-52% of a dose is excreted in the urine as unchanged drug and metabolites, 25-27% eliminated in the feces, and 16-33% into respired air. Urine levels of active drug are considered high, but may be only l/5th of those of amoxicillin.

Before you take Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid

Contraindications / Precautions / Warnings

Penicillins are contraindicated in patients with a history of hyper-sensitivity to them. Because there maybe cross-reactivity, use penicillins cautiously in patients who are documented hypersensitive to other beta-lactam antibiotics (e.g., cephalosporins, cefamycins, carbapenems).

Do not administer systemic antibiotics orally in patients with septicemia, shock, or other grave illnesses as absorption of the medication from the GI tract may be significantly delayed or diminished.

Do not administer penicillins, cephalosporins, or macrolides to rabbits, guinea pigs, chinchillas, hamsters, etc. or serious enteritis and clostridial enterotoxemia may occur.

Adverse Effects

Adverse effects with the penicillins are usually not serious and have a relatively low frequency of occurrence.

Hypersensitivity reactions unrelated to dose can occur with these agents and can manifest as rashes, fever, eosinophilia, neutropenia, agranulocytosis, thrombocytopenia, leukopenia, anemias, lymphadenopathy, or full-blown anaphylaxis.

When given orally, penicillins may cause GI effects (anorexia, vomiting, diarrhea). Because the penicillins may alter gut flora, antibiotic-associated diarrhea can occur and allow the proliferation of resistant bacteria in the colon (superinfections).

Neurotoxicity (e.g., ataxia in dogs) has been associated with very high doses or very prolonged use. Although the penicillins are not considered hepatotoxic, elevated liver enzymes have been reported. Other effects reported in dogs include tachypnea, dyspnea, edema and tachycardia.

Reproductive / Nursing Safety

In humans, the FDA categorizes this drug as category B for use during pregnancy (Animal studies have not yet demonstrated risk to the fetus, hut there are no adequate studies in pregnant women; or animal studies have shown an adverse effect, hut adequate studies in pregnant women have not demonstrated a risk to the fetus in the first trimester of pregnancy, and there is no evidence of risk in later trimesters.) In a separate system evaluating the safety of drugs in canine and feline pregnancy (Papich 1989), this drug is categorized as in class: A (Prohahly safe. Although specific studies may not have proved the safety of all drugs in dogs and cats, there are no reports of adverse effects in laboratory animals or women.)

Overdosage / Acute Toxicity

Acute oral penicillin overdoses are unlikely to cause significant problems other than GI distress, but other effects are possible (see Adverse Effects). In humans, very high dosages of parenteral penicillins, especially in patients with renal disease, have induced CNS effects.

How to use Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid

Note: All doses are for combined quantities of both drugs (unless noted otherwise).

Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid dosage for dogs:

For susceptible infections:

a) 13.75 mg/kg PO twice daily; do not exceed 30 days of therapy (Package insert; Clavamox — Pfizer)

b) For susceptible UTI’s: 12.5 mg/kg PO q12h for 5-7 days For susceptible skin, soft tissue infections: 12.5 mg/kg PO q12h for 5-7 days (may need to extend to 21 days; do not exceed past 30 days). Much higher doses have been recommended for resistant skin infections.

For susceptible deep pyodermas: 12.5 mg/kg PO q12h for 14-120 days

For systemic bacteremia: 22 mg/kg PO q8 – 12h for 7 days

Note: Duration of treatments are general guidelines; generally treat for at least 2 days after all signs of infection are gone. ()

c) For Gram-positive infections: 10 mg/kg PO twice daily

For Gram-negative infections: 20 mg/kg PO three times daily ()

d) For non-superficial pyoderma: 10-25 mg/kg PO twice daily for 3- 6 weeks. Maximum dose is 650 mg twice daily. Increase to three times daily if no response in 1 week. If no response by the 2nd week, discontinue. ()

e) For recurrent pyoderma: 13.75-22 mg/kg PO q8-12h ()

Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid dosage for cats:

For susceptible infections:

a) 62.5 mg PO twice daily; do not exceed 30 days of therapy (Package insert; Clavamox — Pfizer)

b) For Gram-positive infections: 10 mg/kg PO twice daily;

For Gram-negative infections: 20 mg/kg PO three times daily ()

c) For susceptible UTI’s: 62.5 mg/cat (total dose) PO q12h for 10-30 days;

For susceptible skin, soft tissue infections: 62.5 mg/cat (total dose) or 10-20 mg/kg PO q12hfor 5-7 days;

For susceptible sepsis, pneumonia: 10-20 mg/kg PO q8h for 7-10 days Note: Duration of treatment are general guidelines, generally treat for at least 2 days after all signs of infection are gone. ()

Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid dosage for ferrets:

For susceptible infections:

a) 10-20 mg/kg PO 2-3 times daily ()

Amoxicillin / Clavulanate Potassium, Amoxicillin / Clavulanic Acid dosage for birds:

For susceptible infections:

a) 50-100 mg/kg PO q6-8h ()

b) Ratites: 10-15 mg/kg PO twice daily ()

Client Information

■ The oral suspension should preferably be refrigerated, but refrigeration is not absolutely necessary; any unused oral suspension should be discarded after 10 days

■ Amoxicillin/clavulanate may be administered orally without regard to feeding status

■ If the animal develops gastrointestinal symptoms (e.g., vomiting, anorexia), giving with food may be of benefit


■ Because penicillins usually have minimal toxicity associated with their use, monitoring for efficacy is usually all that is required unless toxic signs or symptoms develop. Serum levels and therapeutic drug monitoring are not routinely performed with these agents.

Chemistry / Synonyms

A beta-lactamase inhibitor, clavulanate potassium occurs as an off-white, crystalline powder that has a pKa of 2.7 (as the acid) and is very soluble in water and slightly soluble in alcohol at room temperatures. Although available in commercially available preparations as the potassium salt, potency is expressed in terms of clavulanic acid. Amoxicillin may also be known as: amoxycillin, p-hydroxyampicillin, or BRL 2333; many trade names are available. Clavulanate potassium may also be known as: clavulanic acid, BRL-14151K, or kalii clavulanas.

Storage / Stability / Compatibility

Clavulanate products should be stored at temperatures less than 24°C (75°F) in tight containers. Potassium clavulanate is reportedly very susceptible to moisture and should be protected from excessive humidity.

After reconstitution, oral suspensions are stable for 10 days when refrigerated. Unused portions should be discarded after that time. If kept at room temperature, suspensions are reportedly stable for 48 hours. The veterinary oral suspension should be reconstituted by adding 14 mL of water and shaking vigorously; refrigerate and discard any unused portion after 10 days.

Dosage Forms / Regulatory Status

Veterinary-Labeled Products:

Oral Tablets (4:1 ratio):

62.5 mg: Amoxicillin 50 mg/12.5 mg clavulanic acid (as the potassium salt)

125 mg: Amoxicillin 100 mg/25 mg clavulanic acid (as the potassium salt)

250 mg: Amoxicillin 200 mg/50 mg clavulanic acid (as the potassium salt)

375 mg: Amoxicillin 300 mg/75 mg clavulanic acid (as the potassium salt); Clavamox Tablets (Pfizer); (Rx). Approved for use in dogs and cats.

Powder for Oral Suspension:

Amoxicillin 50 mg/12.5 mg clavulanic acid (as the potassium salt) per mL in 15 mL dropper bottles; Clavamox Drops (Pfizer); (Rx). Approved for use in dogs and cats.

Human-Labeled Products:

Note: Human-labeled amoxicillin/clavulanate products have varying ratios of amoxicillin:clavulanate ranging from 2:1 to 7:1.

Amoxicillin (as trihydrate)/Clavulanic Acid (as potassium salt) Tablets: Amoxicillin 250 mg/125 mg clavulanic acid; Amoxicillin 500 mg/125 mg clavulanic acid; Amoxicillin 875 mg/125 mg clavulanic acid; Augmenting (GlaxoSmithKline); generic (Rx)

Chewable Tablets: Amoxicillin 125 mg/31.25 mg clavulanic acid; Amoxicillin 200 mg/28.5 mg clavulanic acid; 250 mg/62.5 mg clavulanic acid & 400 mg/57 mg clavulanic acid; Augmenting (GlaxoSmithKline); generic; (Rx)

Powder for Oral Suspension — Amoxicillin/Clavulanic Acid (as potassium salt) after reconstitution: Amoxicillin 125 mg/31.25 mg clavulanic acid per 5 mL in 75 mL, 100 mL & 150 mL; Amoxicillin 200 mg/28.5 mg clavulanic acid per 5 mL in 50 mL, 75 mL &100 mL; Amoxicillin 250 mg/62.5 mg clavulanic acid per 5 mL in 75 mL, 100 mL & 150 mL; Amoxicillin 400 mg/57 mg clavulanic acid per 5 mL in 50 mL, 75 mL & 100 mL; 600 mg/42.9 mg clavulanic acid per 5 mL in 75 mL, 100 mL, 125 mL & 200 mL; Augmenting & Augmentin ES-600 (GlaxoSmithKline); Amoclan (West-ward); generic; (Rx)


Amoxicillin (Amoxil, Amoxi-Tabs)


Highlights Of Prescribing Information

Bactericidal aminopenicillin with same spectrum as ampicillin (ineffective against bacteria that produce beta-lactamase)

Most likely adverse effects are GI-related, but hypersensitivity & other adverse effects rarely occur

Available in oral & parenteral dosage forms in USA

What Is Amoxicillin Used For?

The aminopenicillins have been used for a wide range of infections in various species. FDA-approved indications/species, as well as non-approved uses, are listed in the Dosages section below.

Pharmacology / Actions

Like other penicillins, amoxicillin is a time-dependent, bactericidal (usually) agent that acts by inhibiting cell wall synthesis. Although there may be some slight differences in activity against certain organisms, amoxicillin generally shares the same spectrum of activity and uses as ampicillin. Because it is better absorbed orally (in non-ruminants), higher serum levels maybe attained than with ampicillin.

Penicillins are usually bactericidal against susceptible bacteria and act by inhibiting mucopeptide synthesis in the cell wall resulting in a defective barrier and an osmotically unstable spheroplast. The exact mechanism for this effect has not been definitively determined, but beta-lactam antibiotics have been shown to bind to several enzymes (carboxypeptidases, transpeptidases, endopeptidases) within the bacterial cytoplasmic membrane that are involved with cell wall synthesis. The different affinities that various beta-lactam antibiotics have for these enzymes (also known as penicillin-binding proteins; PBPs) help explain the differences in spectrums of activity the drugs have that are not explained by the influence of beta-lactamases. Like other beta-lactam antibiotics, penicillins are generally considered more effective against actively growing bacteria.

The aminopenicillins, also called the “broad-spectrum” or ampicillin penicillins, have increased activity against many strains of gram-negative aerobes not covered by either the natural penicillins or penicillinase-resistant penicillins, including some strains of E. coli, Klebsiella, and Haemophilus. Like the natural penicillins, they are susceptible to inactivation by beta-lactamase-producing bacteria (e.g., Staph aureus). Although not as active as the natural penicillins, they do have activity against many anaerobic bacteria, including Clostridial organisms. Organisms that are generally not susceptible include Pseudomonas aeruginosa, Serratia, Indole-positive Proteus {Proteus mirahilis is susceptible), Enterobacter, Citrobacter, and Acinetobacter. The aminopenicillins also are inactive against Rickettsia, mycobacteria, fungi, Mycoplasma, and viruses.

In order to reduce the inactivation of penicillins by beta-lactamases, potassium clavulanate and sulbactam have been developed to inactivate these enzymes and thus extend the spectrum of those penicillins. When used with a penicillin, these combinations are often effective against many beta-lactamase-producing strains of otherwise resistant E. coli, Pasturella spp., Staphylococcus spp., Klebsiella, and Proteus. Type I beta-lactamases that are often associated with E. coli, Enterobacter, and Pseudomonas are not generally inhibited by clavulanic acid.


Amoxicillin trihydrate is relatively stable in the presence of gastric acid. After oral administration, it is about 74-92% absorbed in humans and monogastric animals. Food will decrease the rate, but not the extent of oral absorption and many clinicians suggest giving the drug with food, particularly if there is concomitant associated GI distress. Amoxicillin serum levels will generally be 1.5-3 times greater than those of ampicillin after equivalent oral doses.

After absorption, the volume of distribution for amoxicillin is approximately 0.3 L/kg in humans and 0.2 L/kg in dogs. The drug is widely distributed to many tissues, including liver, lungs, prostate (human), muscle, bile, and ascitic, pleural and synovial fluids. Amoxicillin will cross into the CSF when meninges are inflamed in concentrations that may range from 10-60% of those found in serum. Very low levels of the drug are found in the aqueous humor, and low levels found in tears, sweat and saliva. Amoxicillin crosses the placenta, but it is thought to be relatively safe to use during pregnancy. It is approximately 17-20% bound to human plasma proteins, primarily albumin. Protein binding in dogs is approximately 13%. Milk levels of amoxicillin are considered low.

Amoxicillin is eliminated primarily through renal mechanisms, principally by tubular secretion, but some of the drug is metabolized by hydrolysis to penicilloic acids (inactive) and then excreted in the urine. Elimination half-lives of amoxicillin have been reported as 45-90 minutes in dogs and cats, and 90 minutes in cattle. Clearance is reportedly 1.9 mL/kg/min in dogs.

Before you take Amoxicillin

Contraindications / Precautions / Warnings

Penicillins are contraindicated in patients with a history of hyper-sensitivity to them. Because there may be cross-reactivity, use penicillins cautiously in patients who are documented hypersensitive to other beta-lactam antibiotics (e.g., cephalosporins, cefamycins, carbapenems).

Do not administer penicillins, cephalosporins, or macrolides to rabbits, guinea pigs, chinchillas, hamsters, etc. or serious enteritis and clostridial enterotoxemia may occur.

Do not administer systemic antibiotics orally in patients with septicemia, shock, or other grave illnesses as absorption of the medication from the GI tract may be significantly delayed or diminished. Parenteral (preferably IV) routes should be used for these cases.

Adverse Effects

Adverse effects with the penicillins are usually not serious and have a relatively low frequency of occurrence.

Hypersensitivity reactions unrelated to dose can occur with these agents and can manifest as rashes, fever, eosinophilia, neutropenia, agranulocytosis, thrombocytopenia, leukopenia, anemias, lymphadenopathy, or full-blown anaphylaxis.

When given orally, penicillins may cause GI effects (anorexia, vomiting, diarrhea). Because the penicillins may alter gut flora, antibiotic-associated diarrhea can occur and allow the proliferation of resistant bacteria in the colon (superinfections).

High doses or very prolonged use have been associated with neurotoxicity (e.g., ataxia in dogs). Although the penicillins are not considered hepatotoxic, elevated liver enzymes have been reported. Other effects reported in dogs include tachypnea, dyspnea, edema and tachycardia.

Reproductive / Nursing Safety

Penicillins have been shown to cross the placenta; safe use during pregnancy has not been firmly established, but neither have there been any documented teratogenic problems associated with these drugs. However, use only when the potential benefits outweigh the risks. In humans, the FDA categorizes this drug as category B for use during pregnancy () In a separate system evaluating the safety of drugs in canine and feline pregnancy (), this drug is categorized as in class: A (Probably safe. Although specific studies may not have proved the safety of all drugs in dogs and cats, there are no reports of adverse effects in laboratory animals or women.)

Overdosage / Acute Toxicity

Acute oral penicillin overdoses are unlikely to cause significant problems other than GI distress but other effects are possible (see Adverse Effects). In humans, very high dosages of parenteral penicillins, especially in patients with renal disease, have induced CNS effects.

How to use Amoxicillin

Amoxicillin dosage for dogs:

For susceptible infections:

a) For Gram-positive infections: 10 mg/kg PO, IM, SC twice daily for at least 2 days after symptoms subside.

For Gram-negative infections: 20 mg/kg PO three times daily or IM, SC twice daily for at least 2 days after symptoms subside ()

b) For susceptible UTI’s: 10-20 mg/kg PO q12h for 5-7 days. For susceptible systemic infections (bacteremia/sepsis): 22-30 mg/kg IV, IM, SC q8h for 7 days.

For susceptible orthopedic infections: 22-30 mg/kg IV, IM, SC, or PO q6-8h for 7-10 days. ()

c) For Lyme disease: 22 mg/kg PO q12h for 21-28 days ()

Amoxicillin dosage for cats:

For susceptible infections:

a) For Gram-positive infections: 10 mg/kg PO, IM, SC twice daily for at least 2 days after symptoms subside.

For Gram-negative infections: 20 mg/kg PO three times daily or IM, SC twice daily for at least 2 days after symptoms subside ()

b) For susceptible UTI’s and soft tissue infections: 50 mg (total dose per cat) or 11-22 mg/kg PO once daily for 5-7 days. For sepsis: 10-20 mg/kg IV, SC, or PO q12h for as long as necessary. Note: Duration of treatment are general guidelines, generally treat for at least 2 days after all signs of infection are gone. ()

c) C. perfringens, bacterial overgrowth (GI): 22 mg/kg PO once daily for 5 days ()

d) C. perfringens enterotoxicosis: 11-22 mg/kg PO two to three times daily for 7 days ()

e) For treating H. pylori infections using triple therapy: amoxi-cillin 20 mg/kg PO twice daily for 14 days; metronidazole 10-15 mg/kg PO twice daily; clarithromycin 7.5 mg/kg PO twice daily ()

Amoxicillin dosage for ferrets:

For eliminating Helicobacter gastritis infections:

a) Using triple therapy: Metronidazole 22 mg/kg, amoxicillin 22 mg/kg and bismuth subsalicylate (original Pepto-Bismol) 17.6 mg/kg PO. Give each 3 times daily for 3-4 weeks. ()

b) Using triple therapy: Metronidazole 20 mg/kg PO q12h, amoxicillin 20 mg/kg PO q12h and bismuth subsalicylate 17.5 mg/kg PO q8h. Give 21 days. Sucralfate (25 mg/kg PO q8h) and famotidine (0.5 mg/kg PO once daily) are also used. Fluids and assisted feeding should be continued while the primary cause of disease is investigated. ()

For susceptible infections:

a) 10-35 mg/kg PO or SC twice daily ()

Amoxicillin dosage for rabbits, rodents, and small mammals:

Note: See warning above in Contraindications a) Hedgehogs: 15 mg/kg IM or PO q12h ()

Amoxicillin dosage for cattle:

For susceptible infections:

a) 6-10 mg/kg SC or IM q24h (Withdrawal time = 30 days) ()

b) For respiratory infections: 11 mg/kg IM or SC q12h ()

c) Calves: Amoxicillin trihydrate: 7 mg/kg PO q8-12h ()

Amoxicillin dosage for horses:

For susceptible infections:

a) For respiratory infections: 20-30 mg/kg PO q6h ()

b) Foals: Amoxicillin Sodium: 15-30 mg/kg IV or IM q6-8h; amoxicillin trihydrate suspension: 25-40 mg/kg PO q8h; amoxicillin/clavulanate 15-25 mg/kg IV q6-8h ()

Amoxicillin dosage for birds:

For susceptible infections:

a) For most species: 150-175 mg/kg PO once to twice daily (using 50 mg/mL suspension) ()

b) 100 mg/kg q8h PO ()

c) 100 mg/kg q8h, IM, SC, PO ()

d) Ratites: 15-22 mg/kg PO twice daily; in drinking water: 250 mg/gallon for 3-5 days ()

Amoxicillin dosage for reptiles:

For susceptible infections:

a) For all species: 22 mg/kg PO ql2 -24h; not very useful unless used in combination with aminoglycosides ()

Client Information

■ The oral suspension should preferably be refrigerated, but refrigeration is not absolutely necessary; any unused oral suspension should be discarded after 14 days

■ Amoxicillin may be administered orally without regard to feeding status

■ If the animal develops gastrointestinal symptoms (e.g., vomiting, anorexia), giving with food may be of benefit

Chemistry / Synonyms

An aminopenicillin, amoxicillin is commercially available as the trihydrate. It occurs as a practically odorless, white, crystalline powder that is sparingly soluble in water. Amoxicillin differs structurally from ampicillin only by having an additional hydroxyl group on the phenyl ring.

Amoxicillin may also be known as: amoxycillin, p-hydroxyampicillin, or BRL 2333; many trade names are available.

Storage / Stability / Compatibility

Amoxicillin capsules, tablets, and powder for oral suspension should be stored at room temperature (15-30°C) in tight containers. After reconstitution, the oral suspension should preferably be refrigerated (refrigeration not absolutely necessary) and any unused product discarded after 14 days.

Dosage Forms / Regulatory Status/Withdrawal Times

Veterinary-Labeled Products:

Amoxicillin Oral Tablets: 50 mg, 100 mg, 150 mg, 200 mg, & 400 mg; Amoxi-Tabs (Pfizer); (Rx). Approved for use in dogs and cats.

Amoxicillin Powder for Oral Suspension 50 mg/mL (after reconstitution) in 15 mL or 30 mL bottles; Amoxi-Drop (Pfizer); (Rx). Approved for use in dogs and cats.

Amoxicillin Intramammary Infusion 62.5 mg/syringe in 10 mL syringes; Amoxi-Mast (Schering-Plough); (Rx). Approved for use in lactating dairy cattle. Slaughter withdrawal (when administered as labeled) = 12 days; Milk withdrawal (when administered as labeled) = 60 hours.

Human-Labeled Products:

Amoxicillin Tablets (chewable) (as trihydrate): 125 mg, 200 mg, 250 mg, & 400 mg; Amoxf/(GlaxoSmithKline); generic; (Rx)

Amoxicillin Tablets (as trihydrate): 500 mg & 875 mg; Amoxil (GlaxoSmithKline); generic; (Rx)

Amoxicillin Capsules (as trihydrate): 250 mg, & 500 mg; Amoxil (GlaxoSmithKline); generic; (Rx)

Amoxicillin (as trihydrate) Powder for Oral Suspension: 50 mg/mL (in 15 and 30 mL bottles), 125 mg/5 mL in 80 mL & 150 mL; 200 mg/5 mL in 50 mL, 75 mL & 100 mL; 250 mg/5 mL in 80 mL, 100 mL & 150 mL; 400 mg/5 mL in 50 mL, 75 mL & 100 mL; Amoxil & Amoxil Pediatric Drops (GlaxoSmithKline); (Apothecon), Trimox (Sandoz); generic; (Rx)

AmoxiciUin Tablets for Oral Suspension: 200 mg & 400 mg; Disper-Mox (Ranbaxy); (Rx)


Infective Endocarditis

Infective endocarditis (IE) is a life-threatening disorder that results from microorganisms that colonize the cardiac endocardium, which commonly causes destruction of valves or other structures within the heart. Bacteremia is by far the most common etiology, with the mitral and aortic valve most frequently affected. Vegetation may cause thromboembolism or metastatic infections, which involve multiple body organs and produce a large variety of clinical signs which makes diagnosis difficult. The incidence of infective endocarditis in necropsied dogs has been reported to range from 0.06% to 6.6%. Evaluation of clinical data from university animal hospitals points to infective endocarditis as a comparably rare condition with incidences ranging from 0.04% to 0.13%. Medium to large breed, mainly purebred, middle-aged male dogs are reported to be predisposed. The incidence in cats, based on clinical experience, is considered to be 7 to 10 times lower than in dogs. Animals with congenital heart disease have a low incidence of infective endocarditis”, but associations have been reported with subaortic stenosis and occasionally with PDA. infective endocarditis has not been found to have any association with chronic mitral valve insufficiency in dogs.

Infective Endocarditis: Pathology

Vegetation associated with by infective endocarditis mainly affects the left heart with the highest incidence involving the mitral valve. Involvement of the right heart or mural endocardium is uncommon. Pathologic findings vary and depend on the virulence of the infecting organism, the duration of infection, and the immunologic response. Intracardiac vegetation consists of different layers of fibrin, platelets, bacteria, red and white cells, and is often covered by an intact endothelium. Bacteria may continue to grow despite antibiotic therapy owing to the location deep within the vegetation and a slow metabolic rate. Necrosis and destruction of the valve stroma or chordae tendineae proceed rapidly in peracute or acute infective endocarditis, which causes valvular insufficiency and cardiac failure.

Infective Endocarditis: Etiology and Pathogenesis

Transient or persistent bacteremia is a prerequisite for the development of infective endocarditis. A large number of bacteria have been associated with bacteremia (see section on Blood Culture below) and some are known to cause infective endocarditis. Most bacteria require predisposing factors to cause infective endocarditis, such as depression of the immunosystem or endothelial damage, sometimes with depositions of platelet-fibrin complexes, to adhere to the valve and create infective endocarditis. The origin of the bacteremia may be active infection localized somewhere within the body. A proportion of cases with infective endocarditis has no clinically detectable source of infection. Possible routes for bacteria to reach and infect the endocardium are by direct contact with the surface endothelium via the bloodstream or from capillaries within the valve (vasculitis).

The consequences of infective endocarditis depend on several factors: virulence of the infective agent; site of infection; degree of valvular destruction; influence of vegetation on valvular function; production of exo- or endotoxins; interaction with the immunosystem with the formation of immunocomplexes; and development of thromboembolism and metastatic infections. Gram-negative bacteremia results often in a peracute or acute clinical manifestation, whereas gram-positive bacteremia typically results in a subacute or chronic condition. The vegetation may cause valvular insufficiency or obstruction. The destruction of valvular tissue is caused by the action of bacteria or the cellular response from the immunologic system. Deposition of immunocomplexes in different organs may cause glomerulonephritis, myositis, or polyarthritis. Septic embolization that produces clinical signs is uncommon but 84% of affected dogs had evidence of systemic embolization at necropsy and glomerulonephritis was reported in 16% of 44 dogs with infective endocarditis.

Infective Endocarditis: Case History and Clinical Signs

The diagnosis of infective endocarditis can easily be overlooked because the case history and clinical signs are not specific and there may be an absence of predisposing factors to raise the suspicion of infective endocarditis. Clinical signs are variable and occur in different combinations. Commonly reported signs include lethargy, weakness, fever (sometimes recurrent), anorexia, weight loss, GI disturbances, and lameness. Stiffness and pain originating from joints or muscles may be caused by immunomediated responses and abdominal pain may be caused by secondary renal or splenic infarction, septic embolization, or abscess formation. If the condition leads to severe valvular damage, especially of the aortic valve, signs of cardiac failure and syncope from arrhythmias may occur. Predisposing factors that in combination with the clinical signs above, should raise the suspicion of infective endocarditis are immunosuppressive drug therapy, such as cortico-steroids; aortic stenosis; recent surgery, especially in conjunction with trauma to mucosal surfaces in the oral or genital tract and infections in these body regions, especially prostatitis; indwelling catheters, infected wounds, abscesses, or pyoderma.

Physical Examination

Most clinical signs lack specificity for infective endocarditis. However, fever, heart murmur (particularly if newly developed), and lameness are considered classical signs. Fever is reported to occur in 80% to 90% in dogs with infective endocarditis. Absence of fever is reported to be more common in cases with aortic valve involvement but may also be attributed to treatment with antibiotics or corticosteroids.

Since aortic insufficiency is otherwise uncommon in dogs, the finding of a diastolic murmur and bounding peripheral pulse should raise the suspicion of infective endocarditis of the aortic valve. Systolic murmurs may be caused by destruction of the mitral valve, which results in mitral regurgitation or vegetations that obstruct the aortic outflow tract, which leads to stenosis. These murmurs are, in contrast to diastolic murmurs, poor indicators of infective endocarditis since they frequendy occur in dogs with other conditions, such as chronic mitral valve insufficiency and aortic stenosis. It should be noted that 26% of dogs with infective endocarditis are reported to lack audible murmurs. Lameness is also an inconsistent finding in infective endocarditis with an incidence of 34% in one study. A range of other physical findings may be present, depending on which organs are affected by circulating immunocomplexes or septic embolization. Possible findings are pain reactions from muscles or abdomen (spleen, intestines, or kidneys), cold extremities, cyanosis, and skin necrosis from severe embolization and a variety of neurologic disturbances if the central nervous system is affected.

Blood Culture

Positive blood cultures are crucial evidence of infective endocarditis. The theory that bacteremia from infective endocarditis is intermittent has changed in recent years to the opinion that, if existent, it is continuous. Thus negative or intermittent positive cultures are unusual when collection and handling of samples is conducted properly. The time for sampling is probably not critical, but a constant finding through repeat samplings is valuable to exclude sample contamination. The technique for obtaining samples aseptically and anaerobically is important and described in detail below. In cases of positive blood culture, it is important to evaluate if the microorganism is consistent with the diagnosis of infective endocarditis.

Microorganisms known to cause infective endocarditis in dogs are, in order of reported incidence, Stapkylococcus aureus, E. coli, betahemolytic streptococci, Pseudotnonas aeroginosa, Corynebacterium spp., Erysipelothrix rhusiopathiae (tonsillarium), and Bartonella irinsonii. B. vinsonii and related proteobacteria has recently been recognized as a potential cause for endocarditis in dogs. They have been found in dogs with cardiac arrhythmias, endocarditis, or myocarditis. Bartonella spp. are also a potential cause for infective endocarditis in cats. Furthermore, Bartonella spp. have been reported to occasionally cause infective endocarditis in immunocompromised (but also immunocompetent) humans, with the cat serving as the major reservoir (cat scratch disease). The recommended antibiotic therapy when the resistance is unknown is erythro-mycin or doxycycline. Immediate antibiotic therapy of humans after significant dog or cat bites may furthermore be motivated as commensals, such as Capnocytophaga canimorsus, in the saliva of dogs and cats have been reported to occasionally cause septicemia with a mortality as high as 30%. Negative blood cultures are fairly common and may be due to antibiotic therapy, chronic situations with “incapsulated” infections, noninfective infective endocarditis (only platelets and fibrin in vegetation), or failure to grow organisms from samples. Some bacteria may grow slowly and samples should not be regarded as definitely negative until they have been incubated for 10 days. More common is a rapid growth of microorganisms with 90% of cultures positive within 72 hours of incubation.

Obtaining Blood Cultures

The referral laboratory should be contacted concerning the preferred type of preprepared vials before obtaining a sample; special additives are available if the patient has been on antibiotics. Pediatric vials are useful because less blood is required but volumes in the range of 20 to 30 mL increase the chance for growth. To avoid contamination, strictly aseptic sampling should be observed which includes thorough shaving and disinfection of the sampling site and strict use of sterile gloves. Three samples with adequately filled vials from different puncture sites should be collected. If samples are collected with a syringe, suction should cease before withdrawal of the needle from the patient to avoid contamination with skin bacteria and a new sterile needle should be used for the transfer of blood into the bottles. The bottles should be prewarmed to 37° C and, after sampling, incubated at the same temperature. Sampling through indwelling catheters should be avoided but may be used as a second choice. The former recommendation to draw samples over 24-hour periods has changed, since multiple simultaneously drawn samples in humans have been shown to be equally sensitive.

Electrocardiographic Findings

Arrhythmia is reported to occur in 50% to 75% of dogs with infective Ventricular premature beats and tachyarrhythmias are the most commonly encountered arrhythmias, but they are usually not life threatening. Deviation in the ST-segment suggests myocardial hypoxia and may indicate coronary artery embolism or ischemia from heart failure. Evidence of chamber enlargement may occur in chronic infective endocarditis. All the mentioned ECG abnormalities are, however, nonspecific.

Radiogrophic Findings

Radiography often does not add any information specific for infective endocarditis. In cases of chronic infective endocarditis with aortic or mitral insufficiency, left-sided cardiac enlargement may be detected. Calcified deposits on the valve leaflets are occasionally observed in chronic cases.

Echocardiographic Findings

Echocardiography has significantly improved the possibility of diagnosis and monitoring of animals with infective endocarditis. Valvular vegetations may be detected using two-dimensional echocardiography, although minor lesions may be difficult to distinguish from myxomatous lesions. M-mode can be used to measure secondary changes in cardiac size and to detect abnormal mitral valve motion such as fluttering from aortic regurgitation. Mitral or aortic regurgitation may be detected using continous or color-flow Doppler echocardiography.

Other Laboratory Findings

Mild anemia is found in 50% to 60% of cases with infective endocarditis. The anemia is similar to those from other infections, usually being normocytic and normochromic. Leukocytosis is found in about 80% of dogs with infective endocarditis, usually due to neutrophilia and monocy-tosis (left shift). Other findings that may be encountered include elevated blood urea nitrogen (BUN) due to embolization, metastatic infection, heart failure, or immune-mediated disease.

Urine analysis may reveal pyuria, bacteriuria, or proteinuria. Elevated serum alkaline phosphatase may be found, probably caused by circulating endotoxins and reduced hepatic function, which may cause hypoalbuminemia. The serum glucose concentration may be decreased and serologic tests for immuno-mediated disease, such as Coombs test, may be positive.

Diagnosis of Infective Endocarditis

Since the clinical signs of infective endocarditis are often a result of complications, rather than reflecting the intracardiac infection, the diagnosis may easily be overlooked. Major criteria for infective endocarditis are positive blood cultures with typical microorganisms for infective endocarditis from two separate samples plus evidence of cardiac involvement. The localization and severity of cardiac lesions is confirmed by echocardiographic visualization of vegetations. In the absence of positive cultures, a tentative diagnosis of infective endocarditis can be made if there is clinical and laboratory evidence of systemic infection, such as fever and leukocytosis plus cardiac involvement and possibly signs of embolization.

Management of Infective Endocarditis

The goal of therapy is to eradicate the infective microorganism and to treat all secondary complications. A successful outcome of the therapy is based on early diagnosis and immediate and aggressive treatment. Only bactericidal antibiotics capable of penetrating fibrin should be considered. The antibiotic concentration in serum and deep within vegetations should exceed the organisms minimal inhibitory concentration (MIC), but preferably also the minimum bactericidal concentration (MBC), continuously or throughout most of the interval between doses. Treatment should continue for at least 6 weeks to eradicate dormant microorganisms.

Management of Cases with Tentative Diagnosis of infective endocarditis

A blood culture (see section above) and an antibiotic sensitivity profile should be obtained. While results from cultures and sensitivity tests are awaited, intravenous treatment with a high dosage of bactericidal antibiotic IV, such as cephalosporins (second generation), should be initiated. Alternatives to cephalosporins are combinations of ampicillin or amoxicillin for gram-positive organisms and gentamicin or amikacin for gram-negative organisms. An alternative to gentamicin and amikacin, which are potentially toxic and only recommended to be used for at most one week, is enrofloxacin for suspected gram-negative infective endocarditis. Enrofloxacin is bactericidal and may penetrate myocardium and heart valves and is also indicated for treating Bartonella infections. Choice of antibiotic should preferably depend on the suspected source of infection and the estimated resistance pattern for the primary infection. Practitioners should try to identify the source of infection and treat it as aggressively as possible, such as use of surgical drainage or debridement. Possible secondary problems should be identified, such as heart or renal failure that need therapy or may impair the prognosis.

For dogs with heart failure from aortic regurgitation, hydralazine titered to an adequate reduction of arterial blood pressure is effective and should be considered as a part of medical therapy. When results are available from blood cultures, appropriate antibiotics are selected and aggressive IV treatment continued for 5 to 10 days while renal function is monitored. If results from cultures are negative, the decision to continue antibiotic therapy should be based on clinical improvement. Depending on the early outcome of therapy, subcutaneous administration may substitute a 5 to 10 days IV treatment, and later be superseded by oral preparations. The duration of therapy should be at least 6 weeks on the effective antibiotic. Frequent clinical examinations, blood screening, and urine analyses should be performed during that period.

Prognosis of Infective Endocarditis

Factors that indicate a poor prognosis include late diagnosis and late start of therapy; vegetations on valves (especially the aortic); gram-negative infections, heart or renal failure that do not respond to therapy; septic embolization or metastatic infection; elevation of serum alkaline phosphatase and hypoalbuminemia (70% mortality is reported if this is found in cases with infective endocarditis); concurrent treatment with corticosteroids, regardless if antibiotics are given simultaneously; treatment with bacteriostatic antibiotics or premature termination of antibiotic therapy. Factors that indicate a more favorable prognosis include only mitral valve involvement (47% of dogs are reported to survive); gram-positive infections, origin of infection being the skin, abscesses, cellulitis, or wound infections.

Prevention of Infective Endocarditis

Prophylactic antibiotics may be indicated 1 to 2 hours before and 12 to 24 hours after diagnostic or surgical procedures when turbulent blood flow is suspected to have damaged the endocardium, such as aortic stenosis, patent ductus arteriosus (PDA), or ventral septal defect (VSD). In these cases, early treatment of all manifest infections is important to avoid bacteremia and reduce the risk for infective endocarditis, and caution should be observed when bleeding or infection is anticipated or evident in the oral, urogenital, intestinal, or respiratory tract. Amoxicillin may be the first choice, but other antibiotics, such as clindamycin or cephalosporins, may also be considered depending on the organ system involved and site of infection.


Aglepristone (Alizin, Alizine)

Injectable Progesterone Blocker

Highlights Of Prescribing Information

Injectable progesterone blocker indicated for pregnancy termination in bitches; may also be of benefit in inducing parturition or in treating pyometra complex in dogs & progesterone-dependent mammary hyperplasia in cats

Not currently available in USA; marketed for use in dogs in Europe, South America, etc.

Localized injection site reactions are most commonly noted adverse effect; other adverse effects reported in >5% of patients include: anorexia (25%), excitation (23%), depression (21%), & diarrhea (13%)

What Is Aglepristone Used For?

Aglepristone is labeled (in the U.K. and elsewhere) for pregnancy termination in bitches up to 45 days after mating.

In dogs, aglepristone may prove useful in inducing parturition or treating pyometra complex (often in combination with a prostaglandin F analog such as cloprostenol).

In cats, it may be of benefit for pregnancy termination (one study documented 87% efficacy when administered at the recommended dog dose at day 25) or in treating mammary hyperplasias or pyometras.


Aglepristone is a synthetic steroid that binds to the progesterone (P4) receptors thereby preventing biological effects from progesterone. It has an affinity for uterine progesterone receptors approximately three times that of progesterone. As progesterone is necessary for maintaining pregnancy, pregnancy can be terminated or parturition induced. Abortion occurs within 7 days of administration.

Benign feline mammary hyperplasias (fibroadenomatous hyperplasia; FAHs) are usually under the influence of progesterone and aglepristone can be used to medically treat this condition.

When used for treating pyometra in dogs, aglepristone can cause opening of the cervix and resumption of miometral contractility.

Within 24 hours of administration, aglepristone does not appreciably affect circulating plasma levels of progesterone, cortisol, prostaglandins or oxytocin. Plasma levels of prolactin are increased within 12 hours when used in dogs during mid-pregnancy which is probably the cause of mammary gland congestion often seen in these dogs.

Aglepristone also binds to glucocorticoid receptors but has no glucocorticoid activity; it can prevent endogenous or exogenously administered glucocorticoids from binding and acting at these sites.


In dogs, after injecting two doses of 10 mg/kg 24 hours apart, peak serum levels occur about 2.5 days later and mean residence time is about 6 days. The majority (90%) of the drug is excreted via the feces.

Before you take Aglepristone

Contraindications / Precautions / Warnings

Aglepristone is contraindicated in patients who have documented hypersensitivity to it and during pregnancy, unless used for pregnancy termination or inducing parturition.

Because of its antagonistic effects on glucocorticoid receptors, the drug should not be used in patients with hypoadrenocorticism or in dogs with a genetic predisposition to hypoadrenocorticism.

The manufacturer does not recommend using the product in patients in poor health, with diabetes, or with impaired hepatic or renal function as there is no data documenting its safety with these conditions.

Adverse Effects

As the product is in an oil-alcohol base, localized pain and inflammatory reactions (edema, skin thickening, ulceration, and localized lymph node enlargement) can be noted at the injection site. Resolution of pain generally occurs shortly after injection; other injection site reactions usually resolve within 2-4 weeks. The manufacturer recommends light massage of the injection site after administration. Larger dogs should not receive more than 5 mL at any one subcutaneous injection site. One source states that severe injection reactions can be avoided if the drug is administered into the scruff of the neck.

Systemic adverse effects reported from field trials include: anorexia (25%), excitation (23%), depression (21%), vomiting (2%), diarrhea (13%) and uterine infections (3.4%). Transient changes in hematologic (RBC, WBC indices) or biochemical (BUN, creatinine, chloride, potassium, sodium, liver enzymes) laboratory parameters were seen in <5% of dogs treated.

When used for pregnancy termination, a brown mucoid vaginal discharge can be seen approximately 24 hours before fetal expulsion. This discharge can persist for an additional 3-5 days. If used in bitches after the 20th day of gestation, abortion maybe accompanied with other signs associated with parturition (e.g., inappetance, restlessness, mammary congestion).

Bitches may return to estrus in as little as 45 days after pregnancy termination.

Overdosage / Acute Toxicity

When administered at 3X (30mg/kg) recommended doses, bitches demonstrated no untoward systemic effects. Localized reactions were noted at the injection site, presumably due to the larger volumes injected.

How to use Aglepristone

WARNING: As accidental injection of this product can induce abortion; it should not be administered or handled by pregnant women. Accidental injection can also cause severe pain, intense swelling and ischemic necrosis that can lead to serious sequelae, including loss of a digit. In cases of accidental injection, prompt medical attention must be sought.

Aglepristone dosage for dogs:

To terminate pregnancy (up to day 45):

a) 10 mg/kg (0.33 mL/kg) subcutaneous injection only. Repeat one time, 24 hours after the first injection. A maximum of 5 mL should be injected at any one site. Light massage of the injection site is recommended after administration. (Label information; Alizin — Virbac U.K.)

To induce parturition:

a) After day 58 of pregnancy: 15 mg/kg subcutaneously one time. 24 hours after aglepristone injection, give oxytocin 0.15 Units/kg every 2 hours until the end of parturition. ()

b) On or after day 58 of pregnancy: 15 mg/kg subcutaneously; repeat in 9 hours. In treated group, expulsion of first pup occurred between 32 and 56 hours after treatment. Use standard protocols to assist with birth (including oxytocin to assist in pup expulsion if necessary) or to intervene if parturition does not proceed. ()

As an adjunct to treating pyometra/metritis:

a) For closed cervix: 6 mg/kg twice daily on the first day followed by the same dose once daily on days 2, 3, and 4. Some prefer using larger doses (10 mg/kg) once daily on days 1, 3,and 8, then follow up also on days 15 and 28 depending on the bitch’s condition. ()

b) For metritis: 10 mg/kg subcutaneously once daily on days 1,2 and 8.

For open or closed pyometra: aglepristone 10 mg/kg subcutaneously once daily on days 1,2 and 8 and cloprostenol 1 meg/ kg subcutaneously on days 3 to 7. Bitches with closed pyometra or with elevated temperature or dehydration should also receive intravenous fluids and antibiotics (e.g., amoxicillin/clavulanate at 24 mg/kg/day on days 1 – 5). If pyometra has not resolved, additional aglepristone doses should be given on days 14 and 28. ()

Aglepristone dosage for cats:

For treating mammary fibroadenomatous hyperplasia: a) 20 mg/kg aglepristone subcutaneously once weekly until resolution of signs. Cats who present with heart rates greater than 200 BPM should receive atenolol at 6.25 mg (total dose) until heart rate is less than 200 BPM with regression in size of the mammary glands. ()


■ Clinical efficacy

■ For pregnancy termination: ultrasound 10 days after treatment and at least 30 days after mating

■ Adverse effects (see above)

Client Information

■ Only veterinary professionals should handle and administer this product

■ When used for pregnancy termination in the bitch, clients should understand that aglepristone might only be 95% effective in terminating pregnancy when used between days 26-45

■ A brown mucoid vaginal discharge can be seen approximately 24 hours before fetal expulsion

■ Bitch may exhibit the following after treatment: lack of appetite, excitement, restlessness or depression, vomiting, or diarrhea

■ Clients should be instructed to contact veterinarian if bitch exhibits a purulent or hemorrhagic discharge after treatment or if vaginal discharge persists 3 weeks after treatment

Chemistry / Synonyms

Aglepristone is a synthetic steroid. The manufactured injectable dosage form is in a clear, yellow, oily, non-aqueous vehicle that contains arachis oil and ethanol. No additional antimicrobial agent is added to the injection.

Aglepristone may also be known as RU-534, Alizine, or Alizin.

Storage / Stability/Compatibility

Aglepristone injection should be stored below 25°C and protected from light. The manufacturer recommends using the product within 28 days of withdrawing the first dose.

Although no incompatibilities have been reported, due to the product’s oil/alcohol vehicle formulation it should not be mixed with any other medication.

Dosage Forms / Regulatory Status

Veterinary-Labeled Products:

Note: Not presently available or approved for use in the USA. In several countries:

Aglepristone 30 mg/mL in 5 mL and 10 mL vials; Alizine or Alizin (Virbac); (Rx)

The FDA may allow legal importation of this medication for compassionate use in animals; for more information, see the Instructions for Legally Importing Drugs for Compassionate Use in the USA found in the appendix.

Human-Labeled Products: None

Veterinary Medicine

Canine Hemorrhagic Gastroenteritis

1. What is canine hemorrhagic gastroenteritis (HGE)?

Canine hemorrhagic gastroenteritis is a syndrome characterized by the acute onset of profuse vomiting and bloody diarrhea with significant hemoconcentration.

2. What is the cause3 of hemorrhagic gastroenteritis?

The cause is unknown. Although the term hemorrhagic gastroenteritis implies an inflammatory condition, the disease is more likely due to altered intestinal mucosal permeability and perhaps mucosal hypersecretion. Cultures of GI contents from HGE-affected dogs have yielded large numbers of Clostridium perfringens, leading to speculation that this organism or its exotoxins are the cause.

3. Which dogs are most likely to be affected with HGE?

Toy and miniature breeds seem particularly prone to HE (hemorrhagic gastroenteritis), especially toy and miniature poodles and schnauzers, but the syndrome may affect any breed.

4. What are the clinical signs of hemorrhagic gastroenteritis?

• Acute onset of vomiting

• Profuse, bloody, fetid diarrhea

• Severe depression

• Shock

5. How is the diagnosis of HGE made?

• Extreme hemoconcentration (packed cell volume > 50-60%)

• Bloody, fetid diarrhea

• No leukopenia

• Fecal cytology with increased numbers of clostridial organisms

6. Describe the treatment for hemorrhagic gastroenteritis.

• Intensive fluid therapy until the packed cell volume is in the normal range and then continued intravenous crystalloid fluids (Normosol-R + potassium chloride) until vomiting is controlled.

• Antibiotics to control C. perfringens (ampicillin or amoxicillin)

• Restriction of food and water

• Antiemetic drugs (metoclopramide)

7. What is the prognosis of hemorrhagic gastroenteritis?

• Early, aggressive fluid therapy consistently results in significant improvement within 24 hours.

• If vomiting and diarrhea are not resolved in 48 hours, a search for other causes mimicking hemorrhagic gastroenteritis should be conducted (parvovirus, coronavirus, GI foreign bodies, intussusception, intestinal volvulus, clostridial enteritis, lymphocytic-plasmocytic enteritis).



Cause of Salmonella

Salmonella spp. are predominandy motile, gram-negative facultative anaerobic rod-shaped bacteria found in the feces of normal and diarrheic animals. As with many other commensal organisms of the gastrointestinal tract, the high prevalence of these organisms complicates diagnosis. From 1% to 30% of the fecal samples or rectal swabs taken from healthy domestic pet dogs, 16.7% of dogs boarded in kennels, and 21.5% of hospitalized dogs were found to be positive on bacteriologic culture for Salmonella. From 1% to 18% of healthy cats and 10.6% of random source research colony cats are also culture-positive for Salmonella. Despite these findings, several species of Salmonella have been impugned in the pathogenesis of acute enterocolitis in dogs and cats. S. typhimurium is the species most commonly isolated from diarrheic feces of dogs and cats, although other species have been identified.

Salmonella: Pathophysiology

Those most at risk for Salmonella infection are young and immunoincompetent animals, those with concurrent gastrointestinal infections (e. g., parvoviral or parasitic infections), and those animals who have had prior therapeutic interventions (e. g., antibiotics or glucocorticoids). Salmonella is an enteroinvasive organism that induces an acute inflammatory response resulting in enterocolitis, mucosal sloughing, and secretory diarrhea. Most Salmonella infections are resolved via the local immune response, but bacterial translocation and septicemia may evolve into systemic inflammatory response syndrome and multiple organ dysfunction syndromes (MODS) in some patients. Early recognition is important in preventing this sequela.

Clinical examination

The main clinical signs of Salmonella enterocolitis are anorexia, lethargy, fever, vomiting, diarrhea with mucous and blood pigments, dehydration, abdominal pain, and tenesmus. With bacterial translocation and septicemia, affected animals may have evidence of pale mucous membranes, weakness, tachycardia, tachypnea, and vascular collapse.

Diagnosis of Salmonella

Culture, serotyping, and polymerase chain reaction are the best methods oi diagnosing Salmonella infections.

Treatment of Salmonella

Treatment varies according to the severity of the clinical signs. Mild, self-limiting forms of enterocolitis may in fact resolve with little more than supportive therapy. Antibiotic therapy in such cases may prolong fecal shedding and encourage development of the carrier state. In animals with severe hemorrhagic diarrhea, history of immunosuppression, suspected or documented septicemia, evidence of systemic inflammatory response syndrome, or a combination of these symptoms, parenteral antibiotics should definitely be used. If culture results are unavailable, therapy should include enrofloxacin, amoxicillin, or trimethoprim-sulfa, all of which are effective against Salmonella. Post-treatment cultures should be performed to confirm eradication, and pet owners should be advised of the public health importance of the disease.

Prognosis of Salmonella

The prognosis for recovery in nonsepticemic patients is generally good, although some animals may remain chronic carriers with recrudescence during periods of stress or unrelated disease. The prognosis for the septicemic patient is more guarded.


Acute Small Intestinal Disease

Potential causes of acute diarrhea are listed in Table Causes of Acute Diarrhea, but whether a complete diagnosis is pursued and when therapy is instituted are clinical judgments. The diagnostic approach to acute diarrhea is discussed elsewhere.

Patients that are bright, alert, and not dehydrated may require no further investigation, because signs are often self-limiting. Further investigation of acute diarrhea is indicated under the following circumstances:

• The patient is dull or depressed, febrile, dehydrated, tachycardic or bradycardic or is having abdominal discomfort, melena, bloody mucoid stools, or frequent vomiting.

• Obvious physical abnormalities (e. g., intestinal masses, thickening, or plication) localize the problem to the small intestine, and diagnostic imaging, noninvasive biopsy, or surgery can define the cause.

• Systemic abnormalities are present, as defined by a minimum database and other clinicopathologic tests.

It is important that the patient be regularly re-evaluated to monitor the response to therapy and to detect any new abnormalities that may arise.

Causes of Acute Diarrhea

Causes Examples
Dietary Hypersensitivity (allergy), intolerance, sudden diet change, food poisoning (poor quality, spoiled foods / bacterial)
Toxic Food or other sources
Infectious Parvovirus, coronavirus, paramyxovirus, adenovirus, may or may not be feline leukemia virus / FIV related; also Salmonella, Campylobacter, Clostridium spp. (?) and Escherichia coli (?) 

Helminths; Coccidia, Ciardia spp.

Acute pancreatitis
Anatomic Intussusception
Metabolic Hypoadrenocorticism

FeLV, feline leukemia virus, FIV, feline immunodeficiency virus

Treatment of Acute Diarrhea

The initial management of acute diarrhea associated with systemic illness is symptomatic and supportive, and is commenced on the basis of clinical findings, in particular the presence of dehydration, while the results of the initial data base and further tests are pending.

Fluid therapy Oral fluid and electrolyte replacement therapy may be sufficient if acute diarrhea is associated with only mild or insignificant dehydration, and if vomiting is infrequent or absent, although its efficacy should still be monitored. However, when diarrhea is accompanied by significant vomiting or dehydration, parenteral fluids should be administered at a rate that replaces deficits, supplies maintenance needs and compensates for ongoing losses. Patients with marked hypovolemia require more aggressive support.

The type of fluid and requirement for potassium supplementation is best judged by performing a minimum data base and blood gas analysis. Parenteral fluids are usually best given intravenously. The intraosseous route can be used if venous access is unavailable, but subcutaneous administration of fluids is likely to be inadequate.

Diet Studies examining the role of diet in the treatment of acute diarrhea in dogs and cats are scarce. Current recommendations are based on common sense and anecdotal evidence Best practice generally is considered to be withholding food for 24 to 48 hours and then feeding a bland diet, given little and often, for 3 to 5 days. Thereafter the original diet is gradually reintroduced. In animals with no other significant clinical findings, this may be the only therapy required. Common choices of a bland, fat-restricted diet for dogs are boiled chicken or white fish or low-fat cottage cheese with boiled rice. Cats seem to have a lower tolerance to dietary starch and may benefit from a diet with a higher fat content. Little attention is paid to the overall nutritional adequacy of home-prepared bland diets when fed in the short term.

This dogma of intestinal rest has been challenged by studies that demonstrate that feeding human infants during diarrhea promotes recovery. The success of such feeding through diarrhea varies depending on the cause, with most benefit seen in secretory diarrhea. However, in dogs and cats, secretory diarrhea is less common, the increased volumes of diarrhea may be cosmetically unacceptable, and the frequently contemporaneous vomiting may preclude this approach. The inclusion of glutamine, a nutrient utilized preferentially by enterocytes, may also promote recovery and decrease bacterial translocatjon, although experimental proof of improved intestinal integrity in animals is lacking.

Theoretically, any intestinal disease may predispose the animal to the development of a food sensitivity, therefore feeding of a novel protein source during these periods may preclude the development of sensitivity to the staple diet. However, this concept of feeding of a sacrificial protein is supported only by circumstantial evidence.

Protectants and adsorbents Bismuth-subsalicylate, kaolin-pectin, montmorillonite, activated charcoal and magnesium, and aluminum and barium-containing products are often administered in acute diarrhea to bind bacteria and their toxins and to coat and protect the intestinal mucosa, but they may also have an antisecretory effect. Therapy for acute diarrhea with protectants, absorbents or motility modifying agents should not exceed 5 days.

Motility- and secretion-modifying agents Anticholinergics and opiates or opioids (loperamide, diphenoxylate) are frequently used for the symptomatic management of acute diarrhea, but anticholinergic agents can potentiate ileus and are not recommended. Opiate analgesics were thought to exert their effects by stimulating segmental motility, but they actually act mainly by decreasing intestinal secretion and promoting absorption and can be used in the short-term symptomatic management of acute diarrhea in dogs. They are contraindicated in cases involving obstruction or an infectious etiology.

Antimicrobial therapy Antimicrobials are indicated only in animals with a confirmed bacterial or protozoal infection, those in which a breach of intestinal barrier integrity is suspected from evidence of gastrointestinal bleeding, and hence in those at risk of sepsis. Leukopenia, neutrophilia, pyrexia, the presence of blood in the feces, and shock all are indications for prophylactic antibiotics in animals with diarrhea. Initial choices in these situations include ampicillin or a cephalosporin (effective against gram-positive and some gram-negative and anaerobic bacteria). If systemic translocation of enteric bacteria is suspected, antimicrobials effective against anaerobic organisms (e. g., metronidazole or clindamycin) and “difficult” gram-negative aerobes (e. g., an aminoglycoside or a fluoroquinolone) are indicated. Intravenous quinolones have been shown to reach therapeutic concentrations in the canine gut lumen and can be effective against enterococci, E. coli, and anaerobes. Oxytetracycline, tylosin, and metronidazole are suitable for the treatment of SIBO.

A four-quadrant, intravenous antibacterial regimen may be required if septicemia is likely, and suitable combinations would be a cephalosporin (or amoxicillin) or a fluoroquinolone (or amikacin) with metronidazole or clindamycin. However, aminoglycosides should not be given until the patient is volume-expanded.

Probiotics Traditionally, many practitioners have recommended feeding live yogurt as a way of repopulating the intestine with beneficial lactobacilli after an acute gastrointestinal upset. There is evidence in other species that probiotics do exert a positive effect on intestinal permeability and mucosal immune responses, although the effects may be species specific and present only while the probiotic is continuously administered. Probiotics are now available for use in dogs and cats, and emerging data exist to support their use.

Acute Diarrhea Induced by Diet, Drugs, or Toxins

Altered food intake, probably the most common cause of acute, self-limiting diarrhea in dogs, includes rapid diet change, dietary indiscretion, dietary intolerance, hypersensitivity, and food poisoning. Dietary hypersensitivity (food allergy) is probably rare. Ingestion of drugs (e. g., nonsteroidal anti-inflammatory drugs [NSAIDs] or antibacterials) or toxins (e. g., insecticides) also may cause vomiting and diarrhea. The history may allow an educated, presumptive diagnosis to be made. However, the exact cause often is never determined because the patient is not systemically unwell and responds to symptomatic therapy. The prognosis usually is excellent, and only if the diarrhea does not respond or the patient deteriorates is further investigation necessary.

Hemorrhagic Gastroenteritis

There are numerous potential causes of bloody vomiting and diarrhea, but hemorrhagic gastroenteritis (HGE) is the name given to a syndrome characterized by acute hemorrhagic diarrhea accompanied by marked hemoconcentration. The cause of the syndrome is unknown. It may represent an intestinal type 1 hypersensitivity reaction or could be a consequence of C. perfringens enterotoxin production.

Clinical findings Dogs present with acute hemorrhagic diarrhea, with small breed dogs most frequently affected. Pyrexia is unusual, but vomiting, depression, and abdominal discomfort are common. The onset may be peracute and can be associated with marked fluid shifts into the small intestine, leading to severe hypovolemic shock even before signs of dehydration (e. g., decreased skin turgor) appear.

Diagnosis A presumptive diagnosis of hemorrhagic gastroenteritis can be made on the basis of appropriate clinical findings associated with a packed cell volume (PCV) of 55% to 60% or more. Total protein is often normal or not as high relative to the packed cell volume. probably because ot intestinal plasma loss. Radiographs may demonstrate ileus. The absence of leukopenia and the presence of marked hemoconcentration help distinguish hemorrhagic gastroenteritis from parvovirus. Positive fecal tests may support a diagnosis of clostridiosis, but direct evidence of small intestine infection is rarely obtained.

Treatment Intravenous fluids are essential in treating patients with hemorrhagic gastroenteritis. Some patients become hypoproteinemic, and plasma or colloid support may be required. Parenteral antibiotics are often administered because of potential clostridial infection and the high risk of sepsis. Clinical improvement is usually noted within a few hours, though the diarrhea may take several days to resolve. Close patient monitoring is essential; patients that have not responded within 24 hours should be re-evaluated for parvovirus, intussusception, or foreign objects. Once the patient is in the recovery phase, standard dietary therapy for acute diarrhea can be instigated. The prognosis for most animals with hemorrhagic gastroenteritis is good, but if hemorrhagic gastroenteritis is complicated by severe hypoproteinemia or sepsis, the prognosis is more guarded.

Infectious and Parasitic Causes of Acute Diarrhea

Diarrhea caused by infectious and parasitic agents is considered common in animals that are young, immunologically naive or immunocompromised, housed in large numbers, or housed in unsanitary conditions. Parvovirus, Giardia, Salmonella, and Campylobacter spp., and some helminths can be significant causes of diarrhea. The importance of coronavirus, C. perfringens, and E. coli as causes of diarrhea has yet to be defined. The zoonotic potential of many of these infections has not been clearly elucidated, but basic hygienic precautions should always be adopted. Specific small intestine infections are discussed below, but the reader is referred elsewhere for detailed information on other viruses such as paramyxoviruses, adenoviruses, feline leukemia, and immunodeficiency viruses, which also cause diarrhea but affect many other organ systems apart from the gastrointestinal tract.


Chronic Gastritis

Gastritis is a common finding in dogs, with 35% of dogs investigated for chronic vomiting and 26% to 48% of asymptomatic dogs affected. The prevalence in cats has not been determined. The diagnosis of chronic gastritis is based on the histologic examination of gastric biopsies and it is usually subclassified according to histopathological changes and etiology.

Histopathologic Features of Gastritis

Gastritis in dogs and cats is usually classified according to the nature of the predominant cellular infiltrate (eosinophilic, lymphocytic, plasmacytic, granulomatous, lymphoid follicular), the presence of architectural abnormalities (atrophy, hypertrophy, fibrosis, edema, ulceration, metaplasia), and their subjective severity (mild, moderate, severe). A standardized visual grading scheme has been proposed by Happonen et al and has been adapted for pathologists.

The most common form of gastritis in dogs and cats is mild to moderate superficial lymphoplasmacytic gastritis with concomitant lymphoid follicle hyperplasia. Eosinophilic, granulomatous, atrophic, and hyperplastic gastritis are less common.


Despite the high prevalence of gastritis an underlying cause is rarely identified, and in the absence of systemic disease, ulcerogenic or irritant drugs, gastric foreign objects, parasites (Physalloptera and Ollulanus spp. ), and in rare instances fungal infections (Pythium insidiosum, Histoplasma spp. ), it is usually attributed to dietary allergy or intolerance, occult parasitism, or a reaction to bacterial antigens, or unknown pathogens. Treatment is often empirical but can serve to define the cause of gastritis, such as diet responsive, antibiotic responsive, steroid responsive, or parasitic.

Although the basis of the immunologic response in canine and feline gastritis is unknown, recent studies in experimental animals have shed light on the immunologic environment in the gastrointestinal tract and reveal a complex interplay between the gastrointestinal microflora, the epithelium, immune effector cells such as lymphocytes and macrophages, and soluble mediators such as chemokines and cytokines. In health, this system avoids active inflammation by antigen exclusion and the induction of immune tolerance. The development of intestinal inflammation in mice lacking the cytokines IL-10, TGFP, or IL-2 indicates the central importance of cytokines in damping down mucosal inflammation. In many of these murine models gastrointestinal inflammation only develops in the presence of indigenous intestinal microflora, leading to the hypothesis that spontaneous mucosal inflammation may be the result of a loss of tolerance to the indigenous gastrointestinal microflora. The role of these mechanisms in outbred species such as the dog and cat remains to be determined, but clearly loss of tolerance to bacterial of dietary antigens should be considered.

The epithelial cell is also emerging as a “general” in the inflammatory response, with gram-negative or pathogenic bacteria inducing proinflammatory cytokine (e. g., IL-8, ILI-p) secretion from epithelial cells, whereas commensal or bacteria such as S. fecium or Lactobacillus spp. induce the production of the immunomodulatory cytokines TGFB or IL-10.The pro-inflammatory cytokines produced by epithelial cells are modulated by the production of IL-10 from macrophages and potentially by the epithelial cells themselves. In this context, dogs with lymphoplasmacytic gastritis of undetermined etiology showed a correlation between the expression of the immunomodulatory cytokine IL-10 and proinflammatory cytokines (IFN-γ, IL-1β, IL-8). Simultaneous expression of IL-10 and IFN-y mRNA has also been observed in the intestines of beagle dogs (lamina propria cells and the intestinal epithelium) in the face of a Iuminal bacterial flora that was more numerous than that of control dogs. Thus it is tempting to visualize a “homeostatic loop” consisting of proinflammatory stimuli and responses, countered by immunomodulation and repair, with an imbalance in either of these arms manifested as gastritis.

The importance of unknown pathogens in the development of mucosal inflammation is best demonstrated by the gastric bacterium Helicobacter pylori, a gram-negative bacterium, which chronically infects more than half of all people worldwide. Chronic infection of human adults with H. pylori is characterized by the infiltration of polymorphonuclear and mononuclear cells and the up-regulation of pro-inflammatory cytokines and the chemokine IL-8.Mucosal T cells in infected individuals are polarized toward the production of gamma interferon (IFN-γ), rather than IL-4 or IL-5 indicating a strong bias toward a TH-1 type response). This sustained gastric inflammatory and immune response to infection appears to be pivotal for the development of peptic ulcers and gastric cancer in people.

There is also a high prevalence of gastric Helicobacter spp. infection in dogs (67% to 100% of healthy pet dogs, 74% to 90% of vomiting dogs, 100% of laboratory beagles) and cats (40% to 100% of healthy and sick cats). In contrast to people, in whom Helicobacter pylori infection predominates, dogs and cats are colonized by a variety of large spiral organisms (5 to 12 n). In cats from Switzerland, United States, and Germany, Helicobacter heilmannii is the predominant species, with Helicobacter bizzozeronii and Helicobacter felis much less frequent. In dogs from Finland, Switzerland, the United States, and Denmark Helicobacter bizzozeronii and Helicobacter salomonis are most common followed by Helicobacter heilmanni and Helicobacter felis. Helicobacter bilis and Flexispira rappini have also been described. Cats can also be colonized by Helicobacter pylori (2 to 5 p) but infection has been limited to a closed colony of laboratory cats.

Ownership of dogs and cats has been correlated with an increased risk of infection of Helicobacter heilmannii in people. Case reports have also suggested the transmission of Helicobacter spp. from pets to man. Recent studies clearly confirm that dogs and cats harbor H. heilmannii, but the subtypes of H. heilmannii present in dogs and cats (types 2 and 4) are of minor importance (approximately 15% of cases) to humans, who are predominantly colonized by H. heilmannii type 1 (the predominant Helicobacter sp. in pigs).

The effect of eradicating Helicobacter spp. on gastritis and clinical signs, the main form of evidence supporting the pathogenic role of H. pylori in human gastritis, has not been thoroughly investigated to date in dogs and cats. An uncontrolled treatment trial of dogs and cats with gastritis and Helicobacter spp. infection showed that clinical signs in 90% of 63 dogs and cats responded to treatment with a combination of metronidazole, amoxicillin, and famotidine, and that 14 of the 19 animals re-endoscoped had resolution of gastritis and no evidence of Helicobacter spp. in gastric biopsies.

Controlled clinical trials are required to confirm these observations but have been hampered by a much higher apparent recrudescence or re-infection rate than the 1 % to 2% per year observed after treatment of H. pylori-infected people. With such limited information from eradication trials, most current knowledge about the pathogenicity of Helicobacter spp. for dogs and cats comes from the evaluation of animals with and without infections and clinical signs, and a small number of experimental infections.

The large Helicobacter species found in dogs an cats do not attach to the epithelium but colonize the superficial mucus and gastric glands, particularly of the fundus and cardia, and may also be observed intracellularly. Degeneration of gastric glands, with vacuolation, pyknosis, and necrosis of parietal cells is more common in infected than uninfected animals. Inflammation is generally mononuclear in nature and ranges from mild to moderate in severity. Gastric lymphoid hyperplasia is common and can be extensive in dogs and cats infected with Helicobaaer spp. (particularly when full thickness gastric biopsies are evaluated). In addition to this local gastric immune response, a systemic response characterized by increased circulating anti-Helicobacter IgG has been detected in sera from naturally infected dogs and cats. However, the gastritis observed in cats and dogs infected with large HLOs is generally less severe than that observed in Helicobacter pylori infected humans (where neutrophilic aggregates, and moderate to severe gastritis, are commonly encountered), and gastro-intestinal ulcers, gastric neoplasia, or changes in serum gastrin or acid secretion have not been associated with Helicobacter spp. infection in dogs and cats.

These differences between people, dogs, and cats may be attributed to differences in the virulence of the infecting Helicobacter spp., or the host response. Studies that address this issue indicate that Helicobacter pylori evokes a more severe pro-inflammatory cytokine and cellular response in dogs and cats than natural or experimental infection with large Helicobacter spp. The limited mucosal inflammatory response and absence of clinical signs in the vast majority of dogs and cats infected with non-H. pylori Helicobaaer spp., despite significant antigenic stimulation (evidenced by seroconversion and lymphoid follicle hyperplasia) suggest that large gastric Helicobacter spp. are more commensal than pathogenic. With this in mind, it is interesting to speculate that it is the loss of tolerance to gastric Helicobacter spp., rather than the innate pathogenicity of these bacteria, that explains the development of gastritis and clinical signs in some dogs and cats. However, much still remains to be learned about the role of Helicobacter spp. in canine and feline gastritis.

Clinical Findings

The major clinical sign of chronic gastritis is vomiting of food or bile. Decreased appetite, weight loss, melena, or hematemesis is variably encountered. The concurrent presence of dermatoIogic and gastrointestinal signs raises the likelihood of dietary sensitivity. Access to toxins, medications, foreign bodies, and dietary practices should be thoroughly reviewed. The signalment should not be overlooked as it may increase the probability that chronic gastritis is the cause of vomiting. Hypertrophy of the fundic mucosa is frequendy associated with a severe enteropathy in basenjis and stomatocytosis, hemolytic anemia, icterus, and polyneuropathy in Drentse Patrijshond. Hypertrophy of the pyloric mucosa is observed in small brachycephalic dogs such as Lhasa apso and is associated with gastric outflow obstruction (see Delayed Gastric Emptying and Motility Disorders). Atrophy of the gastric mucosa that may progress to adenocarcinoma has been reported in Lundehunds with protein-losing gastroenteropathy.

Young, large breed, male dogs in the Gulf states of the United States may have granulomatous gastritis caused by Pythium spp. with infection more prevalent in fall, winter, and spring. Physical examination is often unremarkable. Abdominal distension may be related to delayed gastric emptying caused by obstruction or defective propulsion. Abdominal masses, lymphadenopathy, or ocular changes may be encountered in dogs with gastric fungal infections.


A biochemical profile, complete blood count, urinalysis, and T4 (cats) should be performed as a basic screen for metabolic, endocrine, infectious, and other non-GI causes of vomiting, as well as the acid base and electrolyte changes associated with vomiting, outflow obstruction, or acid hypersecretion. Clinicopathologic tests are often normal in patients with chronic gastritis.

Eosinophilia may prompt the consideration of gastritis associated with dietary hypersensitivity, endoparasites, or mast cell tumors. Hyperglobulinemia and hypoalbuninemia may be present in basenjis with gastropathy / enteropathy, or dogs with gastric pythisosis. Panhypoproteinemia is a feature of gastroenteropathy in Lundehunds, moderate to severe generalized inflammatory bowel disease, gastrointestinal lymphoma, and gastrointestinal histoplasmosis. More specific testing, such as an adrenocorticotropic hormone stimulation test, or serology for Pythium isnsidiosum, is performed based on the results of these initial tests. Determination of food specific IgE has not been shown to be useful in the diagnosis of dietary sensitivity in dogs or cats. The utility of noninvasive tests, such as serum pepsinogen and gastric permeability to sucrose, used to diagnose gastritis in people has not been determined in dogs and cats.

Abdominal radiographs are frequently normal in dogs and cats with gastritis but may show gastric distention or delayed gastric emptying (food retained more than 12 hours after a meal). Contrast radiography may reveal ulcers or thickening of the gastric rugae or wall but has largely been supersceded by the combination of ultrasonography to detect mural abnormalities and endoscopy to observe and sample the gastric mucosa.

Endoscopic examination enables the visualization of foreign bodies, erosions, ulceration, hemorrhage, rugal thickening, lymphoid follicle hyperplasia (evident as mucosal pock marks), increased mucus or fluid (dear or bile stained), and increased or decreased mucosal friability. Discrete focal or multifocal mucosal nodules may be observed with Ollulanus spp. infection.

Gastric phycomycosis can be associated with irregular masses in the pyloric outflow tract and may prompt serologic testing by ELISA, Western blotting, and culture of fresh gastric biopsies. Parasites such as Physalloptera spp. may be observed as 1- to 4-cm worms. Large amounts of bile stained fluid is suggestive of duodenogastric reflux-associated gastritis, whereas lots of clear fluid my indicate hypersecretion of gastric acid. Gastric fluid can be aspirated for cytology (Helicobacter spp., parasite ova or larvae) and pH measurement. Impression smears of gastric biopsies are an effective way of looking for Helicobacter spp. (5 to 12 u spirals) and are more sensitive than the biopsy urease test (Helicobacter spp. produce urease). Serum gastrin should be measured in the face of unexplained gastric erosions, ulcers, fluid accumulation, or mucosal hypertrophy.

The endoscopic procedure of dribbling dietary antigens onto the gastric mucosa to ascertain the presence of food allergy has not been useful in dogs or cats: it is highly subjective, detects only immediate hypersensitivity, and does not correlate with the results of dietary elimination trials. The stomach should be biopsied even when it looks grossly normal (usually three biopsies from each region- pylorus, fundus, and cardia). Thickened rugae may require multiple biopsies, and a full-thickness biopsy is often required to differentiate gastritis from neoplasia or fungal infection and to diagnose submucosal or muscular hypertrophy. The results of gastric ultrasonography can help to forewarn the clinician of these possibilities and are complement to the endoscopic findings.

Gastric sections should be stained with H&E for evaluation of cellularity and architecture, and modified Steiner stain for gastric spiral bacteria. Further special stains, such as Gomori’s methenamine silver, are indicated if pyogranulomatous inflammation is present to detect fungi. Masson’s trichrome can be used to highlight gastric fibrosis, whereas sirius red and alcian blue help to reveal eosinophils and mast cells, respectively. Immunocytochemistry can be employed to help distinguish lymphoma from severe lymphocytic gastritis. Mucin staining has been performed in Lundchunds with gastric atrophy and showed an abnormal presence of mucus neck cells and pseudo-pyloric metaplasia.

The interpretation of gastric biopsies has important implications for patient care because biopsy findings are often used to guide treatment. For example, moderate lymphoplasmacytic gastritis without Helicobacter spp. infection is often treated with corticosteroids, whereas mild lymphoplasmacytic gastritis may be treated with a change in diet. As the histopathologic evaluation of gastric biopsies has not been standardized, the prudent clinician should carefully review histologic sections to get a feel for the pathologist’s interpretation. Even with optimum evaluation similar histologic changes can be observed in patients with different underlying etiologies, so well-structured treatment trials often form the basis of an etiologic diagnosis.


Treatment of gastritis initially centers on the detection and treatment of underlying metabolic disorders and the removal of drugs, toxins, foreign bodies, parasites, and fungal infections.

Parasitic Gastritis

Ollulanus tricuspis is a microscopic worm (0.7 to 1 mm long, 0.04 mm wide) that infects the feline stomach. Its predominant cat-to-cat transmisison is through ingestion of vomitus. It can also undergo internal autoinfection with worm burdens reaching up to 11, 000 per stomach. Mucosal abnormalities range from none, to rugal hyperplasia, and nodular (2 to 3 mm) gastritis.

Histologic findings include lymphoplasmacytic infiltrates, lymphoid follicular hyperplasia, fibrosis, and up to 100 / hpf globular leukocytes. Ollulanus spp. are not detected by fecal examination and require evaluation of gastric juice, vomitus, or histologic sections for larvae or worms. Gastric lavage and xylazine-induced emesis have been described to aid diagnosis. Treatment with fenbendazole 10 mg / kg PO SID 2d may be effective.

Physalloptera spp. are about 2 to 6 cm long worms that are sporadically detected in the stomachs of dogs and cats. Physalloptera rara are most commonly described and appear to be primarily a parasite of coyotes. Diagnosis is difficult as worm burden is often low and the eggs are transparent and difficult to see in sugar floatation. Treatment with pyrantel pamoate (5 mg / kg PO:dogs single dose; cats two doses 14 days apart) may be effective. Control of infection may be difficult due to the ingestion of intermediate hosts, such as cockroaches and beedes, and paratenic hosts, such as lizards and hedgehogs.

Given the difficult diagnosis of Ollulanus and Physalloptera spp., empirical therapy with an anthelminthic such as fenbendazole may be warranted in dogs and cats with unexplained gastritis.

Gastric infection with Gnathostoma spp. (cats), Spirocerca spp. (dogs), and Aonchotheca spp. (cats) has been associated with gastric nodules that have been treated by surgical resection of affected gastric tissue.

Gastric Pythiosis

The presence of transmural thickening of the gastric outflow tract and histology that indicates pyogranulomatous inflammation raise the possibility of infection with fungi such as Pythium insidiosum. Special staining (Gomoris methenamine silver), culture, serology, and PCR of infected tissues can be used to help confirm the diagnosis. Treatment consists of aggressive surgical resection combined with itraconazole (10 mg / kg PO SID) and terbinafine (5 to 10 mg / kg PO SID) for 2 to 3 months post-surgery. ELISA titers of pre- and post-treatment samples may show a marked drop during successful treatment and drugs can be stopped. Medical therapy is continued for another 2 to 3 months if titers remain elevated. The prognosis is poor and fewer than 25% of afflicted animals are cured with medical therapy alone.

Helicobacter-Associated Gastritis

The general lack of knowledge of the pathogenicity of gastric Helicobacter spp. has meant that veterinarians are faced with the dilemma of either treating or ignoring spiral bacteria observed in biopsies from patients with chronic vomiting and gastritis. In light of their pathogenicity in man, ferrets, cheetahs, and mice, it would seem prudent that eradication of gastric Helicobacter spp. is attempted prior to initiating treatment with immunosuppressive agents to control gastritis. However, this must be decided on an individual basis. For example, in the patient with a lvmphoplasmacytic infiltrate of the stomach and small intestine with a concomitant gastric Helicobacter spp. infection, should one treat for inflammatory bowel disease, Helicobacter, or both?

The author recommends treating only symptomatic patients that have biopsy-confirmed Helicobacter spp. infection and gastritis. Current treatment protocols are based on those found to be effective in humans infected with Helicobacter pylori. An uncontrolled treatment trial of dogs and cats with gastritis and Helicobacter spp. infection showed that clinical signs in 90% of 63 dogs and cats responded to treatment with a combination of metronidazole, amoxicillin, and famotidine, and that 74% of 19 animals re-endoscoped had no evidence of Helicobacter spp. in gastric biopsies.

Unfortunately these promising results regarding the eradication of Helicobacter spp. have not been borne out by more controlled studies in asymptomatic Helicobacter-infected dogs and cats. Treatment combinations that have been critically evaluated are (1) amoxicillin (20 mg / kg PO BID 14d), metronidazole (20 mg / kg PO BID 14d), and famotidine (0.5 mg / kg PO BID 14d) in dogs; (2) clarithromycin (30 mg PO BID 4d), metronidazole (30 mg PO BID 4d), ranitidine (10 mg PO BID 4d), and bismuth (20 mg PO BID 4d) (CMRB) in H. heilmannii infected cats and (3) azithromycin (30 mg PO SID 4d), tinidazole (100 mg PO SID 4d), ranitidine (20 mg PO SID 4d) and bismuth (40 mg PO SID 4d)(ATRB) in H. heilmannii-infecled cats. Re-evaluation of infection status at 3 days (dogs) or 10 days (cats) after treatment revealed six of eight dogs and 11 of 11 CMRB and four of six ATRB-treated cats to be Helicobacter spp. free on the basis of histology and urease testing (dogs) or C-urea breath test (dogs and cats). However, at 28 days (dogs) or 42 days (cats) after completing antimicrobial therapy, eight of eight dogs and four of eleven cats that received CMRB, five of six cats that received ATRB were found to be re-infected. A transient effect of combination therapy (amoxicillin 20 mg / kg PO TID 2Id, metronidazole 20 mg / kg PO TID 21d, and omeprazole 0.7 mg PO SID 2Id) on bacterial colonization has also been observed in six cats with H. pylori infection.

Further analysis of gastric biopsies from infected dogs and H. pylori infected cats using PCR and Helicobacter-specific primers revealed persistence of Helicobacter DNA in gastric biopsies that appeared negative on histology and urease testing. These studies suggest that antibiotic regimens that are effective against H. pylori in people may only cause transient suppression, rather than eradication, of gastric Helicobacter spp. in dogs and cats.

The author has recendy employed the combination of amoxicillin (20 mg / kg PO BID), clarithromycin (7.5 mg / kg PO BID) and metronidazole (10 mg / kg PO BID) for 14 days to eradicate Helicobacter pylori infection in cats. Further controlled trials of antibiotic therapy in infected dogs and cats, particularly symptomatic patients with gastritis and Helicobacter spp. infection, are clearly required before guidelines regarding the treatment of gastric Helicobacter spp. in dogs and cats can be made.

Chronic Gastritis of Unknown Cause

Lymphocytic plasmacytic gastritis of unknown cause is common in dogs and cats. It may be associated with similar infiltrates in the intestines, particularly in cats (who should also be evaluated for the presence of pancreatic and biliary disease). The cellular infiltrate varies widely in severity and it may be accompanied by mucosal atrophy or fibrosis, and less commonly hyperplasia.

Patients with mild Iymphoplasmacytic gastritis are initially treated with diet. The diet is usually restricted in antigens to which the patient has been previously exposed, such as a lamb-based diet if the patient has previously been fed chicken and beef, or contains hydrolyzed proteins (usually chicken or soy) that may be less allergenic than intact proteins. Many of these diets are also high in carbohydrate and restricted in fat, which facilitates gastric emptying, and may contain other substances such as menhaden fish oil or antioxidants that may alter inflammation.

The test diet is fed exclusively for a period of about 2 weeks while vomiting episodes are recorded. If vomiting is improved a challenge with the original diet is required to confirm a diagnosis of food intolerance. The introduction of a specific dietary component to the test diet, such as beef, is required to confirm dietary sensitivity. If vomiting is unresponsive the patient may be placed on a different diet for another 2 weeks, usually the limit of client tolerance, or started on prednisolone (1 to 2 mg / kg / day PO, tapered to every other day at the lowest dose that maintains remission over 8 to 12 weeks).

Patients with moderate to severe Iymphoplasmacytic gastritis are usually started on a combination of a test diet and prednisolone. If the patient goes into remission they are maintained on the test diet while prednisolone is tapered and potentially discontinued. Antacids and mucosal protectants are added to the therapeutic regimen if ulcers or erosion are detected at endoscopy or if hematemesis or melena is noted.

If gastritis is unresponsive to diet, prednisolone, and antacids, additional immunosuppression may be indicated. Gastric biopsies should be carefully re-evaluated for evidence of lymphoma. In dogs immunosuppression is usually increased with azathioprine (PO 2 mg / kg SID for 5d then EOD, on alternating days with prednisolone). Chlorambucil may be a safer alternative to azathioprine in cats (PO) and has been successfully employed in the management of inflammatory bowel disease and small cell lymphoma. Prokinetic agents such as metoclopramide, cisapride, and erythromycin can be used as an adjunct where delayed gastric emptying is present. These are discussed below.

Diffuse eosinophillic gastritis of undefined etiology is usually approached in a similar fashion to Iymphoplasmacytic gastritis. The presence of eosinphilia, dermatologic changes, and eosinophilic infiltrates may be even more suggestive of dietary sensitivity. In cats it should be determined if it is part of a hypereosinophilic syndrome. Treatment for occult parasites, dietary trials, and immunosuppression can be carried out as described above. Focal eosinophilic granulomas can be associated with parasites or fungal infection that should be excluded prior to immunosuppression with corticosteroids.

Atrophic Gastritis

Atrophic gastritis in dogs and cats is often associated with a marked cellular infiltrate. In people atrophy is associated with Helicobacter spp. infection and inflammation, and immune-mediated destruction. Gastric disease is often not discovered until the patient presents with pernicious anemia secondary to cobalamin deficiency caused by a lack of gastric intrinsic factor. In people, atrophic gastritis, intestinal metaplasia of the gastric mucosa, and hypochlorhydia are thought to precede the development of gastric cancer. The host inflammatory response is also thought to contribute to the development of atrophy and pro-inflammatory IL-1β and IL-10 gene polymorphisms in people are associated with increased inflammation, gastric atrophy, hypochlorhydria, and gastric cancer.

Atrophic gastritis has been infrequently described in dogs and cats but does share some similarities with people. Atrophy has been associated with gastric adenocarcinoma in Lundehunds and in dogs with Iymphoplasmacytic gastritis of undetermined cause atrophy correlates with the expression of mRNA for IL-1 B and IL-10 and the presence of neutrophils. However, there is no clear evidence that Iymphoplasmacytic gastritis progresses to atrophy and gastric cancer in dogs or cats, and the role of Helicobacter spp. or antigastric antibodies in the development of atrophy in dogs and cats remains to be determined.

In contrast to humans, dogs and cats with atrophic gastritis have not been reported to develop cobalamin deficiency. This is probably because the pancreas, rather than the stomach, is the main source of intrinsic factor in these species. Achlorhydria has been described in dogs and may enable the proliferation of bacteria in the stomach and upper small intestine, although this has not been proven. The treatment of atrophic gastritis has received limited attention, but Helkobacter spp. eradication and immunosuppression have been effective in people.

Hypertrophic Gastritis

Hypertrophy in the fundic mucosa is uncommon and is often part ol the breed-specific gastropathies or gastroenteropathies mentioned above. Concurrent hypergastrinemia should prompt consideration of underlying hepatic or renal disease, achlorhydria, or gastrin-producing tumors, which should be pursued appropriately. Basenji gastoenteropathy is variably associated with fasting hypergastrinemia and exaggerated secretin stimulated gastrin, and anecdotal reports suggest that affected basenjis may respond to antimicrobial therapy. Antral hypertrophy of brachycephalic dogs causes outflow obstruction and is treated with surgery.

Veterinary Medicine

Selected Acquired Diseases Of The Lips, Cheeks, And Palate

Feline eosinophilic granuloma complex (FEGC) comprises an eosinophilic ulcer, plaque, and a linear granuloma. Oral lesions are usually a linear granuloma or an eosinophilic ulcer; the latter has a predisposition for the maxillary lips (80%). Intraoral lesions appear as one or more discrete, firm, raised nodules. Clinical signs include dysphagia and / or ptyalism. Although the etiology of this disease is unknown, bacterial and viral infections and immune-mediated and hypersensitivity diseases have been associated with feline eosinophilic granuloma complex. Biopsy of the lesion, with the aforementioned diseases in mind, should be performed to confirm the diagnosis and to differentiate it from neoplastic disease. Ancillary tests should include a complete blood count, which usually shows an absolute eosinophilia. Concurrent or potentially causative hypersensitivity diseases should be considered during the diagnostic phase of treatment. The mainstay of feline eosinophilic granuloma complex treatment is corticosteroid therapy. Intralesional triamcinolone (3 mg weekly), oral prednisolone (1. 0 to 2. 0 mg / kg given twice daily), and subcutaneous methylprednisolone acetate (20 mg every 2 weeks), administered until feline eosinophilic granuloma complex resolves, are efficacious treatments. Progestational compounds (progesterone or medroxyprogesterone) are often used to treat feline eosinophilic granuloma complex These compounds are not approved for use in cats and have potential side effects that make them undesirable, including adrenocortical suppression, polydipsia, polyuria, polyphagia, obesity, personality change, reproduction abnormalities, mammary hypertrophy, neoplasia, and diabetes mellitus. Cats with untreated chronic lesions, responsive previous lesions, and lesions refractory to corticosteroid therapy have a 50% recurrence rate within 5 months. Failure of treatment is usually related to inadequate dosage or premature cessation of therapy. Animals that do not respond to either corticosteroids or progestational compounds have a poor prognosis and are candidates for more aggressive therapy, such as irradiation, cryosurgery, laser therapy, or immunotherapy.

The term stomatitis refers to an inflammation of the oral mucosa. Oral inflammatory lesions in dogs and cats have multiple causes, necessitating a consistent and logical diagnostic approach. A complete history and thorough physical examination are essential. Dogs and cats with no evidence of debilitating systemic disease should receive a short-acting intravenous anesthetic to allow an unimpeded visual and tactile oral examination. Oral ulcerations occur in at least four different immunemediated diseases, including systemic lupus erythematosus, bullous (pemphigus) disease, idiopathic vasculitis, and toxic epidermal necrosis. The many infectious diseases that are manifested by lesions in the oral cavity include feline leukemia virus, feline immunodeficiency virus, feline syncytium-forming virus, feline calicivirus, feline herpes virus, and feline infectious peritonitis. fis Canine distemper and feline panleukopenia virus may cause stomatitis, although other organs are more severely affected. Candidiasis (infection with Candida albicans) may cause severe stomatitis in dogs and cats. Many cats with stomatitis have immunosuppressive disease or systemic debilitation or have received chronic immunosuppressive therapy. Although the oral manifestation may appear as a white, pseudomembranous covering of the tongue, the lesions are usually irregular, ulcerated areas in zones of inflamed mucosa.

Feline oral inflammatory disease ranges from simple gingivitis to varying degrees of stomatitis in which inflammation extends beyond the mucogingival junction into the oral mucosa. Cats with chronic gingivitis / stomatitis may have ulceration and extension of granulation tissue involving the palatoglossal folds and fauces. Clinical signs include halitosis, ptyalism, dysphagia, inappetence, and weight loss. Extensive disease is marked by root resorption and possibly bony sequestrae in edentulous areas. Unfortunately, because the causation is usually unknown, treatment is symptomatic, including professional cleaning of the teeth, therapy with antimicrobials or with systemic or local corticosteroid agents similar to those used for feline eosinophilic granuloma complex, and laser therapy to stimulate re-epithelialization over inflamed, ulcerated areas. It is not unusual for refractory cases to require extraction of all molars and premolars or all teeth to alleviate the symptoms of this disorder.

Stomatitis may be described as idiopathic despite a thorough diagnostic evaluation. Immunemediated ulcerativc gingivitis / stomatitis afflicts Maltese terriers, although the etiology is verified in only 20% of animals. If diagnostic test results are negative in idiopathic stomatitis, it is appropriate to assume a possible immune-mediated component. A prudent treatment plan includes regular cleaning of the teeth, oral preventive medicine at home, and intermittent or chronic provocative corticosteroid therapy. Antimicrobial therapy emphasizing anaerobic pathogens (e. g. , metronidazole, amoxicillin, clavulanic acid / amoxicillin) may he administered on an intermittent, chronic basis.