On presentation, Ruby was lethargic but alert and responsive. Abdominal pain, pyrexia, and hyporexia were of main concern. Her BCS was 5/9, her rectal temperature was 104.4°F (40.2°C), and she was ≈7% dehydrated. She demonstrated apparent generalized discomfort on abdominal palpation. No peripheral lymphadenopathy was noted, and the rest of the physical examination was unremarkable.

Differential diagnoses for abdominal pain should include any infectious, inflammatory, or neoplastic disease that affects the abdominal organs; rupture, volvulus, or obstruction of an abdominal organ; and referred pain from other muscular, neurologic, or bony structures. Differential diagnoses for pyrexia should include drug administration; toxin ingestion; and infectious, inflammatory/immune-mediated, and neoplastic disease. Differential diagnoses for hyporexia are broad and can be related to abdominal pain or pyrexia; however, other causes may include primary GI or systemic disease; local disease affecting structures involved in prehension, mastication, or swallowing (ie, oral, dental, muscular, bone, neurologic conditions); pain; and behavior.¹

CBC and serum chemistry profile revealed a moderate neutrophilic leukocytosis (23,300/μL; reference interval, 3,000-11,500/μL) without toxic changes. The remaining CBC and serum chemistry results were within normal limits. C-reactive protein (CRP) levels were also assessed for systemic inflammation, and levels were markedly elevated at 29.4 mg/dL (reference interval, <1).

Abdominal ultrasonography revealed the presence of multiple enlarged, heterogeneous, and rounded cranial mesenteric lymph nodes (Figures 1, above, and 2). These findings were suggestive of round cell neoplasia, lymphadenitis (infectious or noninfectious), or reactive lymphadenopathy. There was a small amount of free abdominal fluid, which was sampled and submitted for analysis; this was compatible with a nonseptic suppurative exudate. Culture results of the fluid were negative. Fine-needle aspirates and cytology of the abdominal lymph nodes were consistent with neutrophilic lymphadenitis (Figure 3). No infectious agents were seen. The rest of the abdominal ultrasound was unremarkable.

Differential diagnoses for neutrophilic lymphadenitis include infectious disease, immune-mediated disease, neoplasia, and sterile lymphadenitis.² Thoracic radiography was performed to assess for any distant cause of infection, neoplasia, or inflammation, but the results were unremarkable. Urinalysis was unremarkable, urine culture was negative, and serology for arthropod-borne disease (eg, Ehrlichia spp, Anaplasma spp, Borrelia spp) was negative.

Ultrasound-guided trucut biopsies of the abdominal lymph nodes were submitted for histopathology and culture to exclude infectious lymphadenitis. Histopathology confirmed neutrophilic lymphadenitis (Figure 4), and lymph node tissue cultures (bacterial and fungal) were negative.

Ruby was initially treated with IV crystalloid fluid therapy for dehydration and opioid pain relief (methadone at 0.2 mg/kg every 4 hours) for abdominal discomfort. Antibiotic therapy with potentiated amoxicillin was continued.

Because an infectious cause was not found, treatment with glucocorticoids was started (prednisolone at 1.5 mg/kg every 24 hours). Ruby remained normothermic 48 hours after treatment was initiated; she had a good appetite and there was no evidence of abdominal pain. She was discharged, and treatment with glucocorticoids was continued.

Recheck examinations were scheduled to occur every 3 to 4 weeks. Ruby remained clinically well, and her CRP levels normalized (0.4 mg/dL; reference interval, <1). Repeat CBC also revealed resolution of the inflammatory leukogram. Glucocorticoid treatment was gradually decreased by 25% to 30% every 3 to 4 weeks, on condition that clinical signs remained resolved and physical examination was normal. Abdominal ultrasonography was performed 1 month after initial diagnosis, revealing resolution of the mesenteric lymphadenopathy.

Treatment with glucocorticoids was discontinued ≈4 months after diagnosis because the patient remained clinically well. Ruby remained healthy without treatment 2.5 years after initial presentation.

Sterile steroid-responsive lymphadenitis (SSRL) is not well-characterized in dogs; however, it should be considered in dogs with pyrexia of unknown origin and inflammatory lymphadenopathy for which no underlying cause can be found.

There is little information available about SSRL. Two retrospective, multicentric studies discussed this condition (including clinical signs, diagnostic approach, and treatment) in different breeds,³ particularly in English springer spaniels.⁴ Other case reports/series have also reported on SSRL, especially in English springer spaniels with or without concurrent dermatologic signs.^5-8

A diagnosis of SSRL can be determined via cytology and/or histopathology by documenting an inflammatory infiltrate in lymph nodes without evidence of infectious agents or other underlying disease. It is key that infectious causes be excluded before SSRL is diagnosed; geographic variations are important considerations in these cases, as prevalence of infectious disease can vary depending on the region or country. Good clinical response is initially seen in most patients after glucocorticoid initiation, but other immunosuppressants can be considered if patient response is suboptimal or steroid adverse effects are severe. Relapses are possible after discontinuation of therapy, so monitoring may be necessary.^3,4 CRP evaluation has been used as a monitoring technique. Young female dogs, particularly English springer spaniels, seem to be predisposed to lymphadenitis.^3,4 Previous reports of lymphadenitis in English springer spaniels included a case of sterile neutrophilic-macrophagic lymphadenitis associated with nodular panniculitis,⁶ granulomatous necrotizing lymphadenitis,⁷ and mineral-associated lymphadenopathy.⁸

SSRL should be on the differential diagnosis list for young adult dogs presented with pyrexia and lymphadenopathy.

Following are the top 5 consequences of brachycephaly according to the author.

Shortening of the facial skeleton leads to crowding and compression of the upper airway, resulting in upper airway resistance that increases the work of breathing and leads to air hunger during exertion; this syndrome is known as brachycephalic obstructive airway syndrome (BOAS).² Anatomically, brachycephaly is associated with stenotic nares, hypertrophy of nasal turbinates, an elongated and thickened soft palate, a thickened tongue, everted laryngeal saccules, everted palatine tonsils, and a hypoplastic trachea; this can lead to partial or complete airway obstruction.³ Clinical signs range from mild stridor to severe dyspnea and collapse. Affected dogs may experience sleep apnea.^4,5 Increased airway resistance can also result in soft tissue swelling and laryngeal collapse, exacerbating BOAS and potentially causing complete obstruction of the upper airway, necessitating emergency opening of the airway (Figure 1).^6,7

Because owners are often unaware that BOAS indicates underlying pathology, and because upper airway noises may be considered normal for the breed, there can be delays in seeking veterinary care.⁸ Clinicians should advise owners on when to seek veterinary care and educate them on the sensitivity of brachycephalic breeds to heat stress (Figure 2).⁹

Early surgical correction is recommended to minimize progression of airway pathology in BOAS patients.^10,11 Surgery typically addresses stenotic nares, elongated soft palate, everted laryngeal saccules, and palatine tonsils. Care must be taken when anesthetizing brachycephalic dogs with BOAS because of increased risk for complications (eg, regurgitation, aspiration pneumonia, respiratory distress).^12,13 Perioperative administration of a prokinetic and a histamine blocker, minimal use of opioids, and recovery in an intensive care unit reduced the incidence of postoperative regurgitation in brachycephalic dogs.¹³

Chiari-like malformation of the skull and craniocervical junction is most commonly seen in Cavalier King Charles spaniels, Brussels Griffons, and affenpinschers. Compression of neural tissue and disruption of CSF circulation can result and lead to development of fluid-filled cavities in the spinal cord (ie, syringomyelia [ie, neck scratcher’s disease]).¹⁴ Affected dogs may have concurrent ventriculomegaly,³ a painful condition most commonly seen in smaller brachycephalic breeds.¹⁴

Abnormalities of the vertebral column, including spondylosis deformans and vertebral malformations (eg, hemivertebrae), predispose brachycephalic dogs to intervertebral disk disease.¹⁵ Brachycephalic screw-tailed dogs (eg, French bulldogs, pugs, English bulldogs) are more commonly affected by spinal malformations, including kyphosis and scoliosis, which can increase the risk for intervertebral disk disease.³

Prognosis for affected dogs varies according to the severity of the underlying abnormality. Medical management with analgesic and anti-inflammatory drugs may relieve signs in mildly affected dogs. Surgery may be of benefit for some dogs; however, the prognosis is poor for dogs with marked scoliosis, intractable spinal pain, and/or neurologic signs refractory to medical management.³

Brachycephaly is associated with dental malocclusion, overcrowding, and misalignment of teeth. A number of brachycephalic breeds have mandibular mesioclusion (ie, an undershot jaw), which is specified in breed standards; for example, French bulldog breed standards include an underjaw that is deep, square, broad, undershot, and well turned up.¹⁶

Malocclusion is associated with difficulty chewing food, temporomandibular joint dysfunction, trauma to soft tissue of the oral cavity, and premature tooth loss.¹⁷ Brachycephaly may also predispose dogs to dentigerous cysts, supernumerary incisors, and rotated, fused, or unerupted teeth (Figure 3).¹⁸ Some brachycephalic breeds (eg, boxers, bulldogs) may have prominent palatal rugae, in which plaque, hair, and food become trapped, leading to inflammation and development of granulomas; surgical correction may be required.¹⁸ Dental radiography is essential in the diagnosis and management of dentigerous cysts and supernumerary teeth. Owners should be advised to maintain dental hygiene and pursue regular dental examinations for their pet.

Brachycephalic dogs are at increased risk for GI disease, including hiatal hernia, gastroesophageal reflux, esophagitis, delayed esophageal transit time, and redundant esophagus.^19,20 French bulldogs have a higher incidence of hiatal hernia as compared with other dogs.²⁰ Brachycephalic dogs presented with regurgitation and/or dysphagia are more likely to have esophageal motility disorders as compared with nonbrachycephalic dogs.²¹ Clinical signs of GI disease (eg, dysphagia, vomiting, regurgitation) are common in brachycephalic dogs with clinical upper respiratory tract disease.¹⁹ Of note, GI lesions were seen endoscopically in brachycephalic dogs that did not have GI signs.

Surgical management of respiratory disease may reduce GI signs.^19,20 Because patients with hiatal hernia and esophageal disease are at greater risk for esophagitis and aspiration during and after anesthesia, patients with a history of regurgitation or reflux require close monitoring, and owners should be advised of increased risks.

Brachycephalic breeds may have a variety of conformational defects (eg, medial canthal entropion, trichiasis, inappropriate tear fluid drainage leading to epiphora, qualitative and/or quantitative tear deficiencies, shallow orbits, proptosis, reduced corneal sensitivity) that compromise ocular health.²² These abnormalities increase the risk for ocular disease (including corneal ulcerations and erosions, vascular keratitis, pigmentary keratitis, corneal fibrosis, and keratoconjunctivitis sicca), leading to pain and vision deficits.²² Brachycephalic dogs are 11 to 20 times more likely to be affected by corneal ulcers than are nonbrachycephalic dogs.^23,24 Nasal folds, visible sclera, and an increased eyelid aperture have been identified as risk factors.²³

In some cases, surgical management (eg, medial canthoplasty) can reduce the risk for corneal exposure and irritation. Medical and surgical management may be required to manage acute and chronic conditions (eg, corneal ulceration). Careful attention should be paid to the ocular conformation of these dogs during examination so that owners can be advised accordingly.

Brachycephalic conformation predisposes dogs to respiratory, neurologic, dental, GI, ocular, and other disorders—including dermatologic abnormalities. Because these conditions affect the health and welfare of dogs, it is important they are not dismissed as normal for the breed.

The most common dermatophyte pathogens in small animals are Microsporum canis, M gypseum, and Trichophyton spp.¹ M gypseum is found in and contracted from soil; Trichophyton spp are presumably transmitted through contact with large animals or rodents.¹

Dermatophytosis is more common in warm, humid climates and is seasonal in temperate climates.¹ The primary risk factor is exposure to another infected animal. Group housing of animals (eg, pet shops, animal shelters, animal rescue situations, hoarding) can also increase risk for disease.¹

Successful development of an infection nidus requires exposure of the skin surface to a critical load of infective spores (ie, arthrospores), increased moisture on the skin surface to facilitate sporulation, and microtrauma. The latter is believed to be important in facilitating infection, as experimental infections in cats have been difficult to establish without it.¹ Under optimal conditions, infective spores can adhere to skin within 6 hours, germinate within 24 hours, and begin shedding within 7 days.¹ Lesion foci are detectable within 7 days; however, pet owners may not notice lesions until 14 to 21 days postexposure.¹

The primary mode of transmission is via direct contact with an infected animal. Transmission can also be from contaminated fomites, especially those that can cause microtrauma (eg, clipper blades, grooming tools, collars).

Transmission via a contaminated environment has been shown to be inefficient in establishing active lesions.¹ In the author’s experience, exposure to a contaminated environment most commonly results in culture-positive, lesion-free fomite carriage; however, this exposure can be a risk factor for transmission if the patient is debilitated, has chronic skin issues, and/or has any inflammatory skin disease.

Patients of all ages and breeds can be affected, but young animals and those under severe physiologic stress are predisposed.¹ Persian cats, Yorkshire terriers, and Jack Russell terriers appear to be overrepresented; subcutaneous nodular lesions have been observed almost exclusively in Persian cats and Yorkshire terrier dogs.¹ Working and hunting dogs are predisposed to kerion lesions, which are focal areas of dermatophytosis that resemble nodular draining lesions of deep pyoderma.¹

Clinical history of dermatophytosis is variable. A history of skin lesions in a patient recently adopted from a high-risk situation should raise suspicion. Owners may also report skin lesions on other animals in the home and/or on themselves.

Hair loss, scaling, crusting, and erythema are the most common clinical lesions. Lesions tend to be asymmetric and can affect any area on the body but often appear first on the face, ears, and distal extremities. Lesions may be difficult to find in longhaired animals. Facial lesions appearing as pustular dermatophytosis (ie, resembling pemphigus foliaceus) are rare. Lesions may be focal, multifocal, or widespread, and disease may be mild to severe; presentation reflects the health of the host.

Pruritus may be present and is highly variable; some lesions may be intensely pruritic and exudative and may resemble superficial pyoderma in dogs or exudative eosinophilic lesions in cats.

Nodular inflammatory lesions may be observed in dogs or cats, especially working dogs; nodular lesions in cats may be exudative or subcutaneous.

Purulent paronychia may be observed. Concurrent bacterial infection may worsen clinical signs.

Although dermatophytosis is a differential diagnosis for any inflammatory follicular disease, it is more commonly diagnosed in kittens and puppies and less frequently diagnosed in adult animals. In adult dogs, superficial pyoderma, demodicosis, and other ectoparasitic infestations that can cause hair loss, erythema, and/or scaling should be ruled out. In adult cats, flea infestation, flea allergy dermatitis, and other infestations and allergies associated with generalized scaling should be ruled out.

Dermatophytosis and pemphigus foliaceus can have similar presentations in adult animals and can be differentiated by histopathology, cytology, direct examination of hairs, and fungal culture.

No gold-standard diagnostic test is available for dermatophytosis.¹ Histopathology (± tissue culture) can confirm nodular or pustular dermatophytosis, but fungal culture must be performed to determine fungal species (see Point-of-Care Diagnostics). In a study, direct microscopic examination of hairs (ie, trichogram) and skin scrapings from lesions confirmed infection in >85% of cases.¹ Fungal culture can be used to identify spores on the hair coat and confirm disease if the sample is from a lesion site. Positive fungal cultures from whole-body toothbrush samples may be due to true disease or fomite carriage; thus, sampling should be limited to lesion sites. Dermatophyte PCR is sensitive and specific for fungal DNA but detects both viable and nonviable fungal DNA.¹ PCR has a quicker turnaround time than does fungal culture, but laboratory access may be limited. Field studies comparing fungal culture and PCR are few, so it is not possible to comment on how concordant results are between the tests. If PCR is pursued, a large number of hairs and crusts only from the target lesions should be submitted to the laboratory. A toothbrush fungal culture should also be sent to the laboratory in case it is needed to confirm the infection. The author’s first choice is a fungal culture.

Hairs should be plucked in the direction of growth and skin scrapings obtained from the lesion margins. Mineral oil should be used for mounting. Clearing agents used to soften nail keratin in human medicine (eg, potassium hydroxide) are not needed and can add artifacts or damage microscope lenses, as they are caustic. Cover slips should be placed on slide specimens and viewed at 4× and 10× magnification. Infected hairs are wider and paler in appearance, and ectothrix spores cuffing the hair may be present (Figures 1 and 2).

Wood’s lamp examination is recommended when dermatophytosis is suspected. This is not a diagnostic test but rather a diagnostic tool that helps find suspect M canis-infected hairs for direct examination. In recent studies, 91% to 100% of patients with untreated spontaneous infections showed positive fluorescence.¹ Historic studies reporting 30% to 50% positive fluorescence were from retrospective laboratory studies and not from in vivo studies.

A plug-in Wood’s lamp with a wavelength of 320 to 400 nm and built-in magnification should be used. During examination, the clinician should hold the lamp 2 to 4 cm from the patient’s skin and proceed slowly, starting at the head. Apple-green fluorescence on the hair shaft is suggestive of M canis infection. Crusts do not glow and may need to be lifted (Figure 3) to find infected hairs underneath.

If infection is confirmed with point-of-care diagnostics, treatment can be initiated.

Fungal culture can be performed via point-of-care or laboratory diagnostics. The most commonly used fungal culture medium is dermatophyte test medium (DTM). If fungal culture is performed in-house, easy-open or petri dish-type plates with the largest surface area possible should be used. Fungal culture jars can be difficult to inoculate, are prone to increased bacterial overgrowth due to increased humidity, and can be difficult to obtain samples from the surface. Removing media from the jars is not recommended, as this increases exposure to a possible pathogen.

An untreated lesion should be sampled using a soft-bristled toothbrush, which is mycologically sterile if prepackaged. It is important to sample the skin surface and hairs; materials will be entrapped in toothbrush bristles. Crusts can be gently lifted to sample beneath them using the edge of a skin scraping spatula or other blunt-edged instrument.

In-house fungal cultures can be performed on site, and one study showed that the difference in results between point-of-care testing and a reference laboratory was <3% if proper procedures were followed.² In-house testing practices should follow laboratory biohazard practices.

Fungal cultures should be incubated at room temperature and examined daily; darkness is not necessary. M canis cultures can be finalized at day 14 if no growth is observed.^3,4 Studies have not been conducted for M gypseum, but this pathogen is not difficult to isolate, and it is reasonable to extrapolate studies from M canis to this pathogen. Regarding Trichophyton spp, a study in humans examined 5549 samples and found only 16 required >17 days of incubation, and only 1 of 16 was a veterinary pathogen (T mentagrophtyes).^3,4 The authors concluded 17 days is adequate for finalizing a diagnosis for human pathogens.⁴

Microscopic identification must be performed to confirm diagnosis (see Suggested Reading). Pale, flat, and fluffy gross colonies should be sampled for microscopy. If DTM is used, clinicians should look for pale colonies with a red color change in the medium around them as they grow. It is important to remember that the red color change is not diagnostic of a dermatophyte. Colony morphology may also be suggestive of a positive fungal culture, but it is not diagnostic; microscopic identification is always needed.

In otherwise healthy animals, dermatophytosis usually self-resolves; treatment is intended to shorten the course of disease and limit contagion. Owners should be informed that dermatophytosis is a non-life–threatening zoonotic disease that causes easily treatable skin lesions and be instructed to consult their personal physician if they have questions or suspect they may have skin lesions. Misinformation regarding cleaning, disinfection, and environmental contamination is pervasive; owners should be advised that fungal spores do not invade home surfaces as do other molds (eg, mildew), do not cause respiratory disease, and can be easily removed.

Owners should also be informed that treatment is multimodal and includes reasonable confinement, cleaning, topical therapy, systemic therapy, and monitoring.

Confinement of patients limits the area that requires cleaning and helps prevent the spread of disease. Confinement alone is not curative and should be implemented with care to ensure the welfare of the patient; the area should be a single room large enough to allow eating, sleeping, and exercise.

Dermatophytosis can occur in young animals during key socialization and bonding times; owners should continue to socialize and play with the infected pet. Owners should wear gloves and washable clothing and avoid direct skin-to-skin contact. Hand hygiene is important; owners should wash hands or use hand sanitizer (found to be sporicidal in the author’s laboratory) if soap and water are not available. Safe, washable, interactive toys should be provided. Recommendations for other animals in the home are similar to those for any infectious disease: Direct contact should be avoided, and bowls, brushes, leashes, and bedding should not be shared among animals. Keeping animals physically separated can be challenging; in-contact animals can be bathed with a topical antifungal shampoo (see Topical Therapy) or treated with lime sulfur and watched closely for development of lesions.

Cleaning removes shed-infective material in the environment. Cleaning minimizes false-positive fungal cultures (ie, a culture-positive but lesion-free patient) that complicate determination of mycologic cure and prolong treatment. If cleaning is regularly performed while the patient receives topical therapy, most homes can be decontaminated with 1 or 2 cleanings after cure.⁵ Any items that can be mechanically washed can also be decontaminated.¹

Homes do not need to be aggressively cleaned every day; twice-weekly thorough cleaning of the confinement area is usually sufficient. It is important to mechanically remove gross debris (ie, hairs) on a daily basis. The most efficient method is vacuuming, provided the vacuum has a filter to trap debris and is emptied and cleaned after each use. Removal of debris with disposable dust cloths or wet wipes is adequate between aggressive cleanings.

After removal of gross debris, hard surfaces should be washed with detergent until visibly clean, then rinsed, dried, and sprayed with a disinfectant. Over-the-counter, ready-to-use bathroom disinfectant cleaners with a label claim that it is an antifungal against Trichophyton spp or products containing accelerated hydrogen peroxide are effective against dermatophytes.⁶ Bleach should be avoided, as it can be an irritant, can damage surfaces, and has no detergent properties.

Exposed textiles and soft items should be washed twice with any common laundry detergent on the longest wash cycle possible. Bleach and/or hot water have not been found to be superior to cold water without bleach.⁷ Fabric should be dried according to its label instructions. Agitation from washing (not drying) is antifungal; household dryers do not reach temperatures that are sporicidal. Carpets can be decontaminated by being washed with a beater-brush rug cleaner twice or steam cleaned once. If a disinfectant is desired, antifungal pet shampoo can be substituted for carpet detergent, but color testing should be performed prior to use.⁸

Pet food bowls can be decontaminated via thorough washing with hot, soapy water.⁹

Topical treatment of the hair coat is not considered an optional part of therapy. Topical therapy decreases shedding of infective material, kills ectothrix spores (not affected by systemic therapy¹) on the hair coat, helps prevent development of new lesions, and decreases contagion and environmental contamination. Clipping of the hair coat is not routinely needed, but infected hair may be clipped with metal blunt-tip scissors (ie, to avoid microtrauma to skin from electric clippers). The coat should be combed before application to remove loose hairs. Whole-body hair coat disinfection twice weekly is recommended. Patients should be kept warm (eg, with warm blankets) following whole-body treatment to prevent hypothermia.

In vitro and in vivo studies have shown lime sulfur, miconazole/chlorhexidine gluconate, and enilconazole to be consistently effective.¹ Leave-on rinses are preferred because of their residual activity and should be applied to the face with a sponge.

Leave-on lime sulfur rinse should be applied twice weekly at a 1:16 dilution. This product, which is not available in all countries, may discolor the hair coat and is somewhat odorous. It will also stain fabric and discolor items in contact with it; owners must wear gloves when applying the product and should not let it come into contact with watches or jewelry. Owners should be educated about proper dilution, as concentrated application can be irritating to the skin. Lime sulfur is efficacious, is immediately sporicidal, and has residual activity. Lime sulfur can be drying to the hair coat or footpads when used for prolonged periods. In the author’s experience in shelters, oral ulcers were never observed as a result of use of lime sulfur; cats with oral ulcers had concurrent respiratory infections.

Miconazole (2%)/chlorhexidine gluconate (2%) shampoo is widely available and is sporicidal but does not have residual activity. Although no in vivo studies have determined the optimal contact time, an in vitro study found that 3 minutes of contact was sporicidal¹⁰; therefore, 3 to 10 minutes of contact is recommended. This shampoo can also be used for treatment of exposed but uninfected animals in the home.

Enilconazole leave-on emulsion (1:50 or 1:100) is only labeled for use in cats in France and is not available in all countries. The emulsion is slightly odorous and may be greasy.

Adjuvant focal topical therapy applied once daily is recommended for focal lesions and/or lesions in areas that are difficult to treat (eg, face, ears). This is in contrast to recommendations in most veterinary dermatology textbooks that recommend application twice daily. A recent in vitro study demonstrated good residual activity of clotrimazole (1%), terbinafine (1%), miconazole (0.2%, 1%, or 2%), and 3 leave-on mousse products containing chlorhexidine and climbazole, miconazole, or ketoconazole.¹⁰ Mousse products may be suitable for animals that cannot be wetted or are difficult to treat. Care must be taken to use the product as directed by the manufacturer. For periocular lesions, 2% miconazole nitrate vaginal cream is recommended^11,12; this product is widely used with proven safety by ophthalmologists to treat fungal keratitis. Lesions on or in the ears are best treated with otic preparations with antifungal efficacy.

Owners should always wear gloves when applying topical therapy.

Systemic therapy eradicates infection in the hair follicle and is considered to be an important and necessary part of therapy. Itraconazole (noncompounded) and terbinafine are the most effective and safe treatments for dermatophytosis and have residual activity in the skin and hair, allowing for pulse therapy. Compounded itraconazole should not be used due to poor bioavailability.^13,14 In cats, itraconazole should be administered at 5 mg/kg PO once daily on a week on/week off basis until mycologic cure (see Guidelines for Determining Mycologic Cure). Because this drug is difficult to get into a suspension, the veterinary or human pediatric liquid suspension should be used. If neither is available, 100-mg capsules can be repackaged into 25-mg capsules.

Experimental and field studies have found this drug to be well tolerated, with the most adverse side effects being vomiting and/or decreased appetite.¹ In a licensing study, elevations of liver enzymes posttreatment were noted but deemed to be of little clinical significance, and most remained within normal laboratory values.¹⁵ In a shelter study, 21 cats with dermatophytosis had serum chemistry profile results monitored pre- and posttreatment (ie, itraconazole at 5 mg/kg PO once daily for 21 days). No cats became ill or anorexic. A statistically significant increase in alanine aminotransferase was noted, but no values were outside normal reference ranges.¹⁶ Based on these findings, routine monitoring of liver enzymes is not routinely needed in otherwise healthy animals. Monitoring serum chemistry profiles is important in animals with comorbidities.

Alternatively, terbinafine (30-40 mg/kg PO once daily) may be administered until mycologic cure. Some cats given terbinafine may experience GI effects. In small dogs, itraconazole at 5 mg/kg PO may be administered once daily. Pulse therapy options (eg, week on/week off) may likely be appropriate for dogs, but this has not been confirmed. In larger dogs, terbinafine at 30-40 mg/kg PO should be administered once daily.

Griseofulvin, which is fungistatic and teratogenic and requires intensive monitoring, is no longer recommended because safer choices exist. Ketoconazole and fluconazole are not recommended because they do not have residual activity in the skin and have higher MIC than do itraconazole and terbinafine.

Infected patients should be treated until mycologic and clinical cure are achieved (see Guidelines for Determining Mycologic Cure). Clinical cure commonly precedes mycologic cure and can occur within weeks of treatment initiation. A recent study showed that when compliance with treatment and environmental cleaning was high, 1 negative fungal culture was predictive of mycologic cure.¹⁶ If 2 negative fungal cultures are needed (eg, patient has underlying illness, compliance issues are suspect), they should be obtained at weekly intervals.

Prognosis is good in patients with superficial dermatophytosis and dogs with kerion reactions. Prognosis is less certain in patients with SC nodular lesions; these patients, particularly cats, often require surgical intervention and long-term therapy.

Immunocompromised humans should avoid contact with infected animals during treatment, as is the case in any animal-acquired disease.¹⁷

Culture-positive, lesion-free cats are typically fomite carriers or have subtle lesions not detected at initial examination and should be carefully re-examined (including the head and between the digits) with a Wood’s lamp. Wood’s lamp examination may identify lesions or sites of early infection not visible under examination room light. If no lesions are found, the cat should be bathed with an antifungal shampoo or treated with a lime sulfur rinse; the fungal culture should be obtained again when the cat is dry. Topical therapy should be continued until fungal culture results are available. Commonly, these cats rapidly become fungal culture-negative when allowed to groom and moved to a clean area.

A persistent positive fungal culture following clinical cure has 3 common causes: inadequate disinfection of the hair coat, fomite carriage due to insufficient cleaning of the home, and development of new lesions in areas that are difficult to treat due to inadequate hair coat disinfection. In the case of fomite carriage, owners should be instructed to aggressively clean the patient’s living area and repeat topical therapy; culture testing should be performed again when the cat is dry or within 24 hours.

Now, the benefits of AI can also be applied to veterinary medicine. With the introduction of RenalTech, available exclusively from Antech Diagnostics, AI is set to transform the way we care for pets. Through deep analysis of large sets of health data collected as part of routine diagnostics, RenalTech can predict whether a cat will develop chronic kidney disease (CKD) within two years with greater than 95% accuracy. As the industry’s first predictive diagnostic tool, RenalTech is the future of veterinary care. For the first time, veterinarians can provide care before CKD strikes.

CKD is a multifactorial disease that is difficult to detect early enough to positively impact a cat’s health and longevity. Traditional diagnostics find disease when about 75% of kidney function is lost, while the SDMA biomarker finds disease when about 25% of kidney function is lost. Nonetheless, by the time either of these diagnostics detect disease, organ damage is underway. With RenalTech, veterinarians can intervene early, deliver highly personalized care plans, and inspire better pet owner compliance. 

Initially, researchers at WALTHAM Centre® identified 35 data points as possible predictors of CKD and over time, leveraging machine learning, were able to narrow the list to 6 routine analytes (creatinine, BUN, urine specific gravity, urine protein, urine pH, WBC) and the pet’s approximate age. Powered by data from 150 000 cats seen by Banfield Animal Hospital veterinarians over 20 years, the RenalTech algorithm is the result of collaborative research led by the world’s largest pet care company, Mars Petcare. The vast repository of historical patient data and RenalTech algorithm combine to produce a RenalTech value that allows veterinarians to predict whether cats are likely or not likely to develop CKD.

Antech will offer RenalTech at no additional cost as part of routine feline diagnostic panels. In addition to predicting CKD, the new test helps support the value of preventive care, offering a compelling reason for ongoing diagnostics for comorbidities and other undiagnosed conditions.

RenalTech is the first of a new generation of predictive diagnostic tools poised to ensure veterinary care continues to develop parallel to human healthcare. The ability to predict disease offers veterinarians a powerful, tangible way to inspire pet owner compliance with personalized care plans that maintain pet quality of life and the bond between pet owners and their pets.

• Acute infection, particularly viral 
• Cytotoxic or immunosuppressive drug (eg, chlorambucil, cyclophosphamide) 
• Destruction/disruption of lymphoid tissue (eg, multicentric lymphoma)
• Immunodeficiency (rare; eg, severe combined immunodeficiency syndrome in basset hounds, Cardigan Welsh corgis, Jack Russell terriers, Frisian water dogs)
• Increased exposure to endogenous corticosteroids
  • Acute illness
  • Hyperadrenocorticism
  • Stress (eg, surgery)
• Increased exposure to exogenous corticosteroids
  • Steroid therapy
  • Contact with steroid creams in household
• Loss of lymphocytes (eg, into intestinal tract with lymphangiectasia, into pleural cavity with chylothorax)
• Whole body irradiation

Septic peritonitis is an infection of the peritoneal cavity and often occurs secondary to a ruptured abdominal viscus. Recommendations for treatment have been extrapolated from human medicine and include prompt antimicrobial intervention, surgical treatment for source control, and peritoneal lavage (200-300 mL/kg) to remove/dilute the infectious organisms.^1-3

Forty dogs diagnosed with first-time septic peritonitis between 2011 and 2015 were enrolled in this prospective study evaluating bacterial isolate type, susceptibility, and change in resistance between pre- and postlavage samples. Culture samples were collected intraoperatively before and after lavage. Swabs contacted both the body wall and affected viscera during collection. Prelavage samples were collected on entry to the abdomen, and postlavage samples were collected before closure following sterile glove change. All swabs were submitted for aerobic and anaerobic culture testing.

Empiric antimicrobial therapy was instituted in all dogs preoperatively, with 39 out of 40 (97.5%) receiving appropriate antimicrobials based on pre- and postlavage culture results. Prelavage cultures were positive in 37 out of 40 (92.5%) cases, whereas postlavage cultures were positive in 35 out of 40 (87.5%) cases. Forty-six new isolates were identified in 20 out of 40 dogs; however, a decrease in total number of bacterial isolates was noted in postlavage cultures. The most common bacterial isolates included Escherichia coli, Clostridium perfringens, and Enterococcus faecalis. There was no significant difference in overall resistance between pre- and postlavage samples, although multidrug resistance was identified less commonly postlavage. Survival to discharge occurred in 35 out of 40 (87.5%) dogs, including 1 dog that received inappropriate empiric antimicrobial therapy.

Peritoneal lavage has an effect on both the number and type of bacteria isolated in patients with septic peritonitis. In this study, source control and lavage successfully reduced the overall number of bacterial isolates between pre- and postlavage samples. However, new isolates identified postlavage likely represent mobilization of bacteria during lavage that were not accessible at the time of prelavage sampling. The reduction in multidrug-resistant isolates between pre- and postlavage samples is attributed to source control and lavage, as these were the only interventions performed between sample collections. Early empiric antimicrobial therapy must be initiated for all cases of septic peritonitis; however, critical use of culture results for rapid de-escalation of antimicrobial therapy is paramount. Overall survival (87.5%) to discharge for septic peritonitis was higher than previously reported.

Key pearls to put into practice:

Clinicians should continue to follow previous recommendations for septic peritonitis, including prompt antimicrobial intervention, surgical treatment for source control, and peritoneal lavage.

When collecting culture swabs, clinicians should ensure swabs contact not only the abdominal fluid but also the body wall and affected viscera.

To guide appropriate de-escalation of antimicrobial therapy, clinicians should consider collecting both pre- and postlavage samples to ensure all bacterial isolates are identified. If the pet owner has financial constraints, pooling pre- and postlavage samples can be considered to reduce cost while not compromising the identification of bacterial isolates.

Understanding the causes and factors contributing to development of otitis externa is critical to successful management.^2-4 Causes may directly induce inflammation (primary causes [eg, allergy or endocrine disease]) or lead to disease in abnormal ears (secondary causes [eg, yeast or bacterial overgrowth, overcleaning]).^1-7 Predisposing factors such as conformation of the ear are present before disease develops, and perpetuating factors such as edema or proliferative changes to the ear canal and changes in the tympanic membrane occur as a result of inflammation and can prevent resolution.^1,7 Once identified, primary causes should be addressed, secondary infections resolved, and predisposing and perpetuating factors managed.^1,7

Ear cleaning is an important part of managing otitis externa and should be performed routinely in dogs prone to otitis externa to prevent recurrence.^3,6,7 When performed well, ear cleaning helps maintain a normal ear environment by removing debris, microbes, small foreign bodies, and biofilm that could result in otitis externa.^6,8 By eliminating exudate and debris, a proper assessment of the ear canal and the tympanic membrane can be performed and the inactivation of some antimicrobials by inflammatory material prevented.⁶ Ear cleaning is beneficial in dogs with inflamed ears secondary to allergies, seborrheic ears with excessive cerumen production, and/or stenotic or pendulous ears in which normal epithelial migration is impeded.^6-8

Several cleaning techniques (eg, manual cleansing, bulb syringes, ear flushing) can be used.^1,6 Ear wash or rinse is the most common method used at home; this technique should be clearly demonstrated to the owner to ensure effective cleaning. When performed poorly, ear cleaning can cause trauma, ongoing inflammation, and discomfort, leading to decreased compliance.⁷ To prevent maceration and secondary infection, ear cleaning should generally be performed every 48 hours.⁶ Follow-up examinations are important to assess whether the ears are being cleaned effectively or whether the owner should receive better instructions or employ a new technique.

Many ear cleaning solutions with different active ingredients are available.^1,6,7 The clinician should understand the purpose of each ingredient to recommend the appropriate product.^1,3 It is important that no harm is caused when using an ear cleanser, particularly in ears with a ruptured tympanic membrane. Saline and water should be used in patients without intact tympanic membranes, as many ear cleaning solutions are potentially ototoxic.⁶ Ear cleansers may be used more frequently in certain circumstances (eg, infected ears) or less frequently if used as maintenance to prevent recurrence.

Some ear cleaners, such as those with chlorhexidine or tris-EDTA, have been shown to have antimicrobial activity, which may be due to active ingredients or a low pH.^9-11 Products like these can limit bacterial and yeast proliferation, helping prevent recurrent infections.^12,13

Ceruminolytics emulsify waxes and lipids, which are then more readily flushed from the ear,^1,6 and are commonly used to break up waxy or purulent debris prior to ear flushing or other cleaning under sedation.^6,7,14,15 Ceruminolytics can be ototoxic if left in the middle ear; flushing with water or saline after use may decrease the chance of ototoxicity.

Astringents are used to dry ears and prevent maceration and secondary infection.^6,7 They are often combined with ceruminolytics and surfactants in drying/cleaning products but can be used as sole therapy.^6,7 Common astringent ingredients such as isopropyl alcohol, boric acid, benzoic acid, and salicylic acid can be useful after the ear has been cleaned or prophylactically after bathing, swimming, or the application of aqueous-based solutions in dogs prone to otitis.^6-8

Some ceruminolytics and astringents can cause pain when used in sensitive ears.⁶ Choosing a nonirritating formula for maintenance use is essential to prevent complications and improve compliance. EPIOTIC® Advanced Ear Cleanser is cleanser that breaks down wax and has astringent properties, with a neutral pH and nonirritating formula. With once to twice daily use, EPIOTIC® Advanced Ear Cleanser has been shown to reduce microbial adhesion and work as well as acidic cleaners, with no noted adverse effects.^6,9,16

Effective ear cleaning plays an essential part in otitis externa by helping to manage inflammation and infection that can lead to more complicated otic disease. The product chosen should be based on the individual patient and the underlying causes and factors at play. Demonstrating proper application of the ear cleanser for the owner can help prevent trauma and ongoing inflammation of the canal, which can inhibit resolution of otitis. Frequent follow-up and ongoing training for owners is important to ensuring good compliance and long-term outcomes.

Although less common than atopic dermatitis caused by environmental allergens, cutaneous adverse food reaction (ie, food allergy, food-induced atopic dermatitis) is an important differential diagnosis in a pruritic dog when infections, including parasitic infections, have been ruled out. Diagnosis is typically based on the results of an 8- to 12-week dietary elimination trial, which can often be difficult to perform due to a lack of appropriate pet owner compliance.

The aim of this prospective study* was to evaluate the usefulness of administering a short course of oral prednisolone for a minimum of 2 weeks of a dietary elimination trial to reduce the time required to confirm or refute a food allergy diagnosis. Fifty-three dogs with a diagnosis of nonseasonal atopic dermatitis were fed a commercially available, extensively hydrolyzed diet and given anti-inflammatory dosages of prednisolone for ≥2 weeks. Two weeks after prednisolone was discontinued, 10 of the 53 dogs did not show signs of flare-up; these dogs were then challenged with their original diets and experienced relapse of signs. When the extensively hydrolyzed diet was reintroduced to this group, clinical signs improved. These dogs were subsequently diagnosed with food-induced atopic dermatitis. Median duration of the food elimination trial for the food-allergic dogs was 28 days (range, 28-44 days). In the other 43 dogs, pruritus could not be controlled without concurrent prednisolone administration and clinical signs did not worsen with diet challenge. In these dogs, elimination diet trials lasted a median of 60 days (range, 54-70 days).

Key pearls to put into practice:

Historically, atopic dermatitis has been referred to as allergic skin disease triggered by environmental allergens. There is growing evidence that supports food as a possible trigger for some dogs with atopic dermatitis.¹ In these dogs, pruritic skin disease with clinical characteristics of atopic dermatitis can develop with ingestion of various foods. Canine food allergy is believed to involve both immunologic and nonimmunologic pathomechanisms of development²; this could support shared immunologic responses in the conditions previously considered to be separate.

A strict dietary elimination trial remains the gold standard for diagnosing food allergy in dogs and cats. A recent literature review evaluated information on in vitro and in vivo testing for food allergies in veterinary species³; results lack support for any evaluated test other than diet trial. However, these trials can be frustrating for owners and often lead to noncompliance. Concurrent administration of anti-inflammatory steroids as described in the present study may allow for reduced time to achieve a diagnosis in truly food-allergic dogs.

Reducing inflammation is key when managing allergic skin disease in dogs and cats. Focusing on this in the early stages of dietary elimination trials can help dampen pathways that lead to itch and inflammation. Steroids have a broad effect with regard to inflammation; whether these same effects would be noted with more targeted therapeutics (eg, oclacitinib, lokivetmab) is unknown.

In this study, the authors describe the temporal changes in clinical and radiographic variables prior to development of congestive heart failure (CHF) in dogs with stage B2 myxomatous mitral valve disease. Dogs developing CHF showed increased heart rates, respiratory rates at home and in the clinic, and vertebral heart sums. Rectal temperatures and body weights were decreased. Vertebral heart sums gradually increased over 12 months, whereas the other variables changed in the 2 to 10 months prior to developing CHF. The variables with the highest absolute change and rate of change were observed with respiratory rates at home and in the clinic, suggesting monitoring of these variables may enable earlier detection and management of CHF.

The prevalence of inflammatory bowel disease (IBD) in dogs warrants an easier route for diagnosis, as the current path is costly, invasive, and time-consuming. In this study, researchers explored the possibility of using serologic markers for diagnosis of IBD in dogs, similar to what is done in human medicine. Serologic markers represent the patient’s reaction to translocation of GI pathogens in the bloodstream when the gut mucosal barrier breaks down. Three cohorts were studied: dogs diagnosed with IBD via biopsy, dogs with acute GI signs from causes other than IBD, and a normal cohort. ELISA methods were developed to detect autoantibodies against canine polymorphonuclear leukocytes (ie, antipolymorphonuclear leukocytes antibody [APMNA]) and calprotectin, microbial outer membrane porin C (OmpC), antibodies against food-derived gliadins, and flagellins isolated from diseased dogs. Of these, antibodies against APMNA and Escherichia coli OmpC exhibited the highest single-marker performance for discriminating IBD from other acute GI conditions and normal cohorts.

Feline parasitism not only has the potential to produce disease and unthriftiness in cats but can also cause zoonotic disease in humans (eg, ocular or visceral larval migrans, toxoplasmosis). Therefore, identifying the prevalence and types of parasites seen in cats can be beneficial.

The objective of this retrospective study was to comprehensively evaluate the prevalence and trend of parasitism in client-owned cats over a 12-year period. Results of fecal examinations performed at 2 locations between 2007 and 2018 were evaluated. Results came primarily from the examination of centrifugal flotation with either Sheather’s sugar or zinc sulfate solutions but also included saline direct smears, sedimentation, and Baermann tests. Of the 2,586 samples tested, parasites were observed in 24.5% of samples, with multiple parasites identified in 5.7% of samples. Twenty-three different types of parasites were identified, with the most common being Cystoisospora spp (9.4%), Toxocara cati (7.8%), Giardia spp (4%), Alaria spp (3.5%), Ancyclostoma spp (1.2%), taeniid (1.2%), Dipylidium caninum (1.1%), and Eucoleus (syn Capillaria) aerophilus (0.7%). A significant difference in prevalence was identified between age categories, with the youngest group (<6 months of age) having the highest infection rate (ie, 41%). Prevalence of parasites decreased in each subsequent older age group. The prevalence of Cystoisospora spp and T cati increased in summer months through fall; this seasonality is likely due to the litters of kittens born in spring and summer. The prevalence rate of parasitism increased over the 12-year period.

Key pearls to put into practice:

Although fecal flotation is the most common method of parasitism testing, it is not always the best technique for all parasites. Heavy trematode eggs do not reliably float and are better identified through fecal sedimentation. The Baermann technique is the best test for identifying lungworm larvae. Fecal flotation techniques also differ in their ability to reveal various ova and protozoa¹; the specific gravity of the solution affects the variety of ova and protozoa that float and can also cause distortion, making them harder to detect, and centrifugal flotation is more sensitive than passive flotation.² The type of test should be selected based on the patient’s history and expected findings.

Review of medical records in this study revealed that all cats positive for the rare parasites Trichuris felis and Platynosomum fastosum had recently moved from the Caribbean. When animals are imported from outside the United States or travel with their owners, they may transport exotic diseases. Therefore, clinicians should be familiar with nonendemic parasites to avoid overlooking or misdiagnosing them. 

This study showed that the prevalence rate of feline parasitism continued to increase over the 12-year study period. Along with owner education and year-round, broad-spectrum parasite control, it is vital that clinicians continue to conduct parasite testing and treatment, especially in kittens and young cats.

Although UTIs are relatively common in female dogs and other reviews have been published, the authors of this study from Beijing, China, recognized a possibility for variance in the local and regional population. To further understand the prevalence and antimicrobial resistance of bacteria associated with canine UTIs, analysis of urine samples from dogs with UTIs or other urinary tract diseases was performed.

In processing 326 samples, 129 bacterial isolates were identified from 103 samples. The isolated organisms were similar to other reports¹; Escherichia coli was identified most commonly but only represented approximately one-third of total positive cultures. A variety of other gram-positive and gram-negative organisms comprised the remaining isolates, with E coli, Klebsiella spp, and Staphylococcus spp comprising ≈70% of positive cultures. More than one pathogen was isolated in ≈33% of positive cases. Resistance to common antimicrobials was also common in positive samples. In E coli isolates, the highest rates of resistance were recorded for ampicillin, ceftazidime, and florfenicol. The highest rates of resistance in Staphylococcus spp isolates were recorded for erythromycin, trimethoprim/sulfamethoxazole, and penicillin. These results reinforce the importance of culture and antimicrobial susceptibility testing when planning appropriate UTI treatment. 

The results of this study suggest the existence of rampant multidrug-resistant urinary tract pathogens in the region in which the study took place; however, they are also fairly consistent with what is seen in typical small animal practice.¹ Even with expert screening of urine sediment, only ≈33% of urine cultures grew organisms; negative urine cultures may represent prior antimicrobial treatment, true negatives, or misidentification of sediment artifacts. However, samples from dogs with signs of UTI but unremarkable sediments were not cultured; thus, other infected dogs may have been missed. In addition, although E coli may be expected to cause a proportion of UTIs, two-thirds of cases are caused by other bacteria, and a significant variance in antimicrobial susceptibility can be expected.

Key pearls to put into practice:

This study provides a glimpse of the value of antibiograms and the need for antimicrobial stewardship. An antibiogram assimilates the susceptibility patterns from large numbers of samples at a single laboratory, region, or hospital. Antibiograms are particularly pertinent for hospital-acquired infections, both for planning treatment and tracking resistance patterns. Although an antibiogram does not replace individual susceptibility testing in the management of infection, it does provide some generalizable information to guide empiric treatment selection.

Nearly all urinary pathogens in this study remained susceptible to amikacin and meropenem; however, cost, toxicity, and practicality of these medications limit their clinical value. Similarly, doxycycline appears promising based solely on the antibiogram in this study, but it is not excreted at high levels in urine and is usually reserved for infections resistant to other treatment options.

Antimicrobial stewardship entails limiting antimicrobial exposure and reducing the risk for resistant organisms. Consensus guidelines are available for shorter, targeted, and selective management of UTIs in dogs and cats²; although these guidelines rely heavily on human medical literature and practice and are yet to be tested in veterinary practice, they provide a conservative view of antimicrobial treatment worth adopting. By prioritizing stewardship, reasonable empiric antimicrobial choices, and short treatments based on culture and susceptibility results, the veterinary profession can help support good patient care while blunting induced resistance.

Interdog aggression in a home can be disturbing and frustrating to pet owners, disruptive to everyday life, and potentially dangerous to both the owner and the dogs. The more that can be understood about this problem, the better advice a clinician can give the owner.

This review presented the results of a large, well-designed study that evaluated 305 pairs of dogs (217 included in outcome analysis) presented to a behavior referral practice for aggression toward each other. Cases reviewed had ≥6 months of follow-up or ≥1 of the dogs euthanized or permanently removed from the home. Multiple factors were assessed to determine correlations between interdog aggression and long-term outcome. Many of the results also support previous studies.^1,2

Intrahousehold interdog aggression is typically associated with dog pairs in which resource guarding is a trigger, a fighting pair of dogs that includes ≥1 female dog,¹ dogs of the same sex,¹ situations in which the aggressor dog was acquired after the recipient dog and is younger,² and aggressor dogs that are purebred but not breed-specific.^1,2 Several of these correlations were seen in ≥50% of the cases.

For the 217 pairs that were followed long-term, 55 pairs (25.3%) had poor outcomes, which included 23 pairs that required complete separation from one another, 24 involving ≥1 dog being euthanized, and 8 involving ≥1 dog being rehomed.² Of the remaining 162 pairs with a better outcome, 100 (61.7%) did not have to be separated following behavioral intervention, 32 (19.8%) were separated during triggers, 21 (13%) were kept separate when unsupervised and during triggers, and 9 (5.6%) were kept muzzled when together and supervised.²

Key pearls to put into practice:

Risk factors significantly associated with a poor outcome (eg, euthanasia, permanent separation of the dogs) in dogs with interdog aggression include^1,2:

• Dogs of the same sex, particularly female–female
• A bite serious enough to puncture the skin of the recipient
• The aggressor is ≥2 years younger than the recipient.
• The aggressor was introduced into the household after the recipient.
• An aggressor that is heavier than the recipient
• The aggression is triggered by the sight of the recipient, even without other triggers.
• The owner uses positive-punishment/negative-reinforcement training techniques.

Management is a particularly important part of treatment and should be strongly encouraged when clinicians become aware of the problem. Triggers should be removed if possible. The dogs should be kept separate from each other—particularly if eye contact alone triggers the aggression, when triggers are present, and when unsupervised. Muzzles are recommended, and appropriate muzzle training is emphasized. A variety of psychopharmacologic medications may be helpful. In this study, such medications were prescribed for 82.4% of aggressors and 32.7% of recipient dogs.

Ultimately, when historical information points to risk factors associated with poor outcomes (as described above), strong and immediate intervention is called for by the clinician, often including referral to a board-certified veterinary behaviorist.

Sacrocaudal luxations (ie, tail-pull injuries) are relatively common in cats and present as a limp, sometimes painful, tail. Assessing nerve function is important during physical examination of patients presented with this condition; testing should be performed for distal and proximal tail sensation, anal tone, and perineal reflex. Damage to the caudal nerves can cause decreased sensory and motor function to the tail, and damage to the pelvic and pudendal nerves affects urine and fecal continence.

The goal of this study was to assess long-term outcome and prognostic factors in cats with sacrocaudal luxation. Seventy cats were evaluated retrospectively; 60 had absent tail tone and 53 had absent tail-base sensation. Anal tone was absent in 20 cats and decreased in an additional 13 cats. Inability to urinate voluntarily was noted in 53 cats; inability to defecate voluntarily was observed in 29 cats. Twenty-one of the cats with an inability to defecate voluntarily were constipated, whereas 8 were fecally incontinent.

Of the 61 cats for which urinary outcomes were available, 90% regained voluntary urinary function; 87% of those regained it in <30 days. Cats with a flaccid incontinent urinary bladder had a significantly worse prognosis, with only 50% regaining urinary control at a median of 33 days. With regard to fecal continence, 25% of those incontinent at presentation remained so, and 68.4% of those experiencing constipation at the time of injury continued to experience it at the time of follow-up. Age, sex, tail-base sensation, anal tone, perineal sensation, fecal continence, degree of vertebral displacement, and tail amputation did not affect outcome. Despite nerve dysfunction commonly being noted at the time of injury, most cats regained function with time. Because early tail amputation did not affect outcome, the authors did not recommend this as a treatment for cats with sacrocaudal luxation.

Key pearls to put into practice:

Cats with sacrocaudal luxation should undergo a thorough neurologic examination, including careful evaluation of bladder and anal tone.

Overall, cats with sacrocaudal luxation have a good prognosis for return to function; urinary incontinence with a flaccid bladder may be associated with a worse prognosis.

Based on the high percentage of cats that returned to function in this study, aggressive decisions about euthanasia or tail amputation should not be made until at least 6 weeks postinjury.

Although CT is considered the gold standard for detecting vertebral fractures, human studies have suggested that MRI may be used as a single modality to evaluate spinal cord and soft tissue injuries in addition to vertebral fractures. In this study of 128 vertebrae in 33 dogs, only moderate agreement between 2 expert observers was achieved when evaluating vertebral fractures using MRI, although agreement was substantial with structurally unstable fractures. Fractures in the transverse process were particularly more likely to be missed. It was concluded that MRI is a poor modality for assessing fracture morphology and that, although MRI may be useful for detecting unstable fractures, it should not replace CT for complete evaluation when this modality is available.

Diagnostic surveillance performed at the authors’ laboratory over the past few years suggests the presence of multidrug-resistant (MDR) hookworms (ie, A caninum) likely evolved on greyhound breeding farms and in racing kennels. Most, if not all, actively racing and/or recently adopted greyhounds appear to be infected with MDR hookworms; however, many cases of MDR hookworms have been diagnosed in non-greyhound breeds, suggesting MDR hookworms are spreading to the general canine population.

Hookworms (Figure 1) have a direct life cycle, with adult females releasing a large number of eggs (up to 10,000/day). Once passed in the feces, development of eggs to third-stage infective larvae (L3) typically takes ≈5 days, although this will vary depending on temperature. Dogs may be infected via both the oral and percutaneous routes. L3 larvae are ingested either directly or by ingestion of paratenic hosts carrying L3 tissue larvae. After penetrating the skin, L3 larvae migrate via the bloodstream to the lungs, penetrate the alveoli, migrate up the bronchial tree to the trachea, are expectorated via coughing, are swallowed, and enter the small intestine, where they complete development into the adult stage. The prepatent period for either route of infection is 15 to 26 days. Following skin penetration in dogs older than 3 months of age, A caninum L3 larvae often undergo somatic migration to the muscle, fat, and other organs; encyst; and enter a hypobiotic state.

Encysted somatic larvae may become reactivated under 2 conditions: host pregnancy or larval leakage (ie, when arrested somatic larvae continuously leak from tissue and complete migration to the intestine, where they develop into adults and begin a new round of egg shedding). An important mode of A caninum transmission is the transmammary route, in which puppies become infected by reactivated larvae that migrate to the mammary tissue of the dam.

Although direct evidence is lacking, based on the authors’ observations and previous research on this issue in the sheep parasite, Haemonchus contortus,² it is probable that macrocyclic-lactone resistance has worsened as the use of moxidectin has become more common. Further, drug resistance in nematodes is typically a slow evolutionary process, requiring many years of drug selection to reach levels that are clinically apparent.³ This was most likely the case for A caninum; thus, the level of resistance seen in any particular case and to any particular drug will depend in part on the time frame of the animal’s adoption and previous anthelmintic treatments.

The emergence and spread of MDR hookworms that are poorly responsive to typical anthelmintic treatments necessitate a different management approach.

When addressing persistent cases of A caninum infection, the clinician should first differentiate between larval leakage⁴ with drug-susceptible A caninum and infection with MDR hookworms.

There are 3 methods to diagnose anthelmintic resistance: performing a fecal egg count reduction test (FECRT), submitting a sample to a laboratory that can perform in vitro drug bioassays with hookworms, and submitting a sample to a laboratory that can perform molecular testing for resistance.

FECRT is the ideal practical approach, as laboratory expertise and facilities may not be readily available for the other diagnostic methods. FECRT can be easily accomplished at the clinic level for minimal cost.

To perform an FECRT, the number of eggs per gram of feces must be quantified pre- and posttreatment. Fecal flotations, which are frequently performed in small animal practice, are inadequate for an FECRT. The pretreatment fecal sample can be collected either the day before or the day of treatment and should be kept refrigerated until submission to the laboratory to prevent development and hatching of eggs prior to testing.

To evaluate for resistance, a quantitative fecal egg count (FEC) method (eg, McMaster,⁵ Mini-FLOTAC⁶; Figures 2 and 3) is needed. A double-centrifugation method (eg, modified Wisconsin [Figure 4]) could also be used but is more time consuming, more labor intensive, less accurate, and less precise.^7,8 A quantitative method is necessary to assess response to treatment. This approach is standard for diagnosis of anthelmintic resistance in livestock nematode parasites, for which drug resistance has been a long-standing problem. Further, because only 1 dog is typically being tested, as compared with groups of 10 to 20 livestock, the authors also recommend performing 2 separate FECs on the pretreatment sample and 2 separate FECs on the posttreatment sample. The FEC reduction is then calculated by comparing the average eggs per gram (EPG) for the 2 pretreatment FECs and the average EPG for the 2 posttreatment FECs. By repeating the FEC, the variability of each FEC measurement will be reduced by half, thus improving the accuracy of the measured FEC reduction.

A specialized laboratory that offers the service should be contacted if in-clinic or diagnostic laboratory FEC testing is not an option. McMaster slides, Mini-FLOTAC reading discs, and Fill-FLOTAC devices are available for purchase (see Fecal Egg Count Reduction Test Resources).

During testing, the most-recently used anthelmintic should be readministered, even if it elicited poor therapeutic results. Use of an alternate drug will not allow differentiation between larval leakage and resistance as the cause of treatment failure, as the worms may not be resistant to the new drug. However, this may only be a theoretical concern, as the authors’ experience indicates that drug-resistant hookworm infrapopulations most likely will be MDR to all 3 anthelmintic classes.

Multiple days (≥3) are needed for eggs already shed in the intestine to be fully cleared.⁹ In addition, the authors have observed a temporary, but high, level of suppression on worm fecundity following fenbendazole treatment. A 99% reduction in FEC has been observed by 3 days posttreatment, with egg counts rapidly rising again after ≈10 days.^1,10 This phenomenon has been reported rarely in sheep after treatment with benzimidazole anthelmintics¹¹ and on multiple occasions in strongylids of ruminants^12-14 and pigs¹⁵ following treatment with ivermectin and moxidectin. Consequently, checking FEC too soon posttreatment can yield a false-negative result for resistance.

In contrast, if too much time passes, larvae arrested in somatic tissue could repopulate the intestine and begin a new round of egg shedding, leading to a false-positive result. The prepatent period for A caninum has been reported to be as early as 14 days,^16-18 but this time frame is from studies in immune-naive puppies following primary infection. Few data exist on time to worm maturity and egg production in older dogs with chronic infections; however, based on clinical data and other reports,¹⁹ new worms take 3 to 4 weeks to repopulate the lumen of the small intestine and initiate a new round of egg shedding.

The following timeframes are thus recommended for posttreatment FEC: 10 to 14 days after treatment with pyrantel, 14 days after treatment with fenbendazole/febantel, and 14 days after treatment with moxidectin.

The following formula can be used to calculate FECRT percentage:

FECRT results should be interpreted conservatively, as FEC can be highly variable. It is important to note, however, that when commonly used anthelmintics were first approved, high efficacies were reported based on worm counts (>99% for febantel, moxidectin, and milbemycin oxime^20-22; >98% for fenbendazole²³; variable for pyrantel, with a mean across studies of ≈94% and over half of studies yielding >99%²⁴).

In contrast, in a recent study using an MDR A caninum isolate (Worthy 4.1F3P), the efficacies based on worm counts were 23%, 9%, and 26% for pyrantel, milbemycin oxime, and fenbendazole, respectively.¹⁰ The corresponding FEC reductions measured 10 days posttreatment for these same treatments were 13%, 0%, and 46%, respectively.¹⁰ These data demonstrate that poor FEC reduction can be expected against an MDR A caninum isolate following treatment with typical commercial products.

The following interpretation of FEC reduction results are suggested:

• <75% reduction: indicative of resistance (larval leakage is highly unlikely to be the cause of persistent egg shedding)
• 75% to 89% reduction: suggestive of resistance (larval leakage is unlikely to be the cause of persistent egg shedding)
• 90% to 95% reduction: suggestive of reduced efficacy and should raise suspicion for resistance, but results should be viewed as inconclusive (persistent egg shedding could be due to resistance or larval leakage)
• >95% reduction: suggestive of effective treatment (larval leakage is likely the cause of persistent egg shedding)

FECRT is sensitive for detecting resistance (ie, dead worms do not shed eggs); consequently, effective treatment will produce a high reduction in the number of eggs shed, and a poorly effective treatment will yield a low reduction in eggs shed. However, egg-shedding levels on a per-worm basis can vary greatly, and egg production per worm can increase following treatment that kills some of the worms (referred to as density dependent fecundity).²⁵ Therefore, the actual percentage for reduction should not be overinterpreted; for example, 25% and 70% reduction both indicate resistance, but the results should not be interpreted as being greatly different. Likewise, given the expected variability, it should not be assumed that the reduction in FEC will be the same in each case of resistance or even in the same dog if the FECRT is repeated.

FEC reduction between 75% and 95% yields an inconclusive result; repeating FECRT at the next treatment is advised.

A diagnosis of resistance should only be established if all of the following are true:

• The patient was treated with the proper dosage.
• The drug administered was within the expiration date and stored properly.
• Fecal samples were labeled and stored correctly prior to fecal analysis.
• An FECRT was performed.
• Proper laboratory techniques were applied when conducting the FECRT, and the same method was used on both the pre- and posttreatment samples.

The treatment plan should depend on the results of the FECRT. If FEC reduction is >95%, treatment should be considered effective. Drugs are not 100% effective, even against drug-susceptible worms; thus, some eggs may be seen, particularly when pretreatment FEC is high. Because resistance can be ruled out, eggs seen on previous fecal examinations are most likely a result of larval leakage. The patient should be treated with an anthelmintic monthly, and fecal examinations should be conducted every few months. Moxidectin can be a good choice in dogs with larval leakage, although any effective anthelmintic should work.

If FEC reduction is between 90% and 95%, FECRT should be repeated a few weeks later at the next treatment.

If FEC reduction is between 75% and 90%, FECRT can be repeated for more conclusive results, or because there is a high chance the worms are resistant, the treatment plan suggested below can be followed.

If FEC reduction is <75%, treatment should be considered ineffective and adjusted to a triple anthelmintic combination with all drugs administered concurrently at the labeled doses. Drugs should be administered sequentially on the same day and not mixed together.

This treatment plan has been successful in eliminating active infections in persistent hookworm cases²⁶:

• Febantel (25 mg/kg PO)/pyrantel pamoate (5 mg/kg PO)/praziquantel (5 mg/kg PO) + moxidectin (2.5 mg/kg topical), or
• Fenbendazole (50 mg/kg PO once daily for 3 days) + pyrantel pamoate (5 mg/kg PO) + moxidectin (2.5 mg/kg topical)

Treatment success using this triple-drug combination depends on whether the hookworms are moxidectin-resistant. MDR hookworms studied by the authors were all ivermectin-resistant but may still be moxidectin-sensitive. The aforementioned regimen should be effective if hookworms are moxidectin-sensitive; however, this approach may be ineffective if the infecting source hookworms were previously treated with moxidectin. The authors have diagnosed recent cases of moxidectin-resistant A caninum in greyhounds in which monthly moxidectin treatments offered little benefit. If this monthly treatment regimen is effective in eliminating egg shedding, the patient will need to remain on this treatment for several months, or possibly for life, as somatic tissue stores will continually leak and repopulate the intestine for an extended time.

Extra-label administration at higher-than-label doses might improve efficacy, but there are currently no data to support such a recommendation. In addition, some parasitologists recommend repeating moxidectin treatment every 2 weeks for the first 4 treatments, then treating monthly, as this allows the moxidectin to rapidly reach a steady-state tissue concentration due to the long half-life of moxidectin in dogs.^27-29 This is reasonable and potentially beneficial, although no specific data presently exist. Other products containing moxidectin may also be effective, but there are no published data to support the effectiveness of those products against MDR A caninum isolates.

If the triple combination approach is ineffective, emodepside is the only potentially effective alternative treatment, based on a recent study evaluating the efficacy of emodepside and praziquantel against an MDR A caninum isolate.¹⁰ Oral emodepside (1 mg/kg) with praziquantel (5 mg/kg) demonstrated an efficacy of 99.6% with a 100% reduction in FEC at 10 days posttreatment.

Emodepside is not currently approved for use in dogs in the United States; however, emodepside (with praziquantel) is FDA-approved as a topical solution for cats. The authors have determined that extra-label use of emodepside topical solution for cats, administered PO at a different dose than is recommended for this product, has high efficacy in MDR A caninum isolates refractory to triple combination treatment.

The authors have monitored FEC in 17 client-owned dogs, both greyhounds and nongreyhounds, treated by private practice clinicians using 1 mg/kg PO emodepside administered once. In all cases, FEC reduction was 100% at 14 days, and no adverse effects were observed. However, there are a number of important factors to consider with extra-label emodepside use (see Considerations for Extra-Label Emodepside).

If extra-label use of the emodepside topical solution for cats is warranted, the patient should be fasted overnight prior to administration, and food should not be provided until 4 hours posttreatment. The dose should be given at the clinic and not dispensed to the owner. The product should be drawn into a syringe with a needle; then, the needle should be removed and the syringe administered as distal orally as possible to decrease the ability of the patient to taste the product.

Precise dosing of emodepside is critical and cannot be readily achieved without careful calculations. The correct canine dose cannot be estimated based on the feline label. The product comes in 3 sizes: small cat (5.5 lb [2.5 kg]), medium cat (11 lb [5 kg]), and large cat (17.6 lb [8 kg]), all of which have a different volume but the same concentration of emodepside (21.4 mg/mL). The following formula should be used to determine the correct dose for dogs:

For example, an 8.8-lb (4-kg) dog would receive 0.19 mL, and a 66-lb (30-kg) dog would receive 1.4 mL.

The authors have not evaluated nor are aware of any evidence regarding the use of other concurrent and/or supportive treatments (eg, probiotics) and cannot provide recommendations on their use.

The authors strongly recommend that FEC (not just flotations) be evaluated monthly to monitor egg shedding.

It is critical that strict environmental hygiene is practiced. Feces of a dog shedding hookworm eggs (or any helminth parasite) should be picked up immediately and properly disposed of to eliminate the potential for reinfection or spread.

It takes ≥5 days for hookworm eggs to develop to the infective third-stage larvae in ideal temperature and humidity conditions³⁰; therefore, fecal pickup even every few days can be highly effective in preventing environmental contamination. However, waiting can result in feces breakdown, allowing the hookworm larvae (or other parasite eggs/larvae) to contaminate the environment.

If reinfection with ivermectin-resistant worms from the environment is permitted to occur in dogs treated with moxidectin, resistance to moxidectin can rapidly develop.³¹

There are several methods for killing hookworm larvae in the environment, but their effectiveness is undetermined.