Familial polymyositis in humans has predisposing genetic factors.1 Descriptions of breed-specific variants of polymyositis in dogs are limited to Newfoundlands and Boxers.2 A breed-specific myositis in Hungarian Vizslas, marked by pharyngeal dysphagia and masticatory muscle atrophy, occurs in the United Kingdom.3 In a survey funded by the Vizsla Club of America Welfare Foundation, 16 of 2,505 (0.64%) Vizsla dogs were diagnosed with myositis. Myositis was the 3rd most common disorder affecting nervous tissue or muscles after epilepsy (77 of 2,505; 3.04%) and other cases without a definitive diagnosis (22 of 2,505; 0.88%).4 In this report we describe 3 unrelated Hungarian Vizslas with masticatory muscle atrophy and pharyngeal dysphagia from across the United States (Pennsylvania, California, Georgia) (Fig 1).
major histocompatability complex
A 9-year-old castrated male Vizsla was presented for evaluation of dysphagia, regurgitation, coughing, and vomiting of 3 weeks' duration. The dog displayed ptyalism after eating and drinking. Treatment before presentation included thyroid hormone supplementation (levothyroxine,a 0.01 mg/kg PO q12h), metoclopramideb (0.23 mg/kg PO q12h), metronidazolec (11.9 mg/kg PO q12h), nutriceutical supplementation (HepatoGen,d 2 tablets PO q12h), enrofloxacine (6 mg/kg PO q24h), and maropitant citratef (1.4 mg/kg SC, twice over 4 days). At the time of presentation, the dog was thin. There were several areas of alopecia and seborrhea along the lateral aspect of both elbows, shoulders, and ventrum of the neck. Auscultation of the thorax disclosed harsh lung sounds with tachypnea and increased respiratory effort. Complete blood count (CBC) revealed a normocytic, normochromic anemia (hematocrit 33.4%; reference range 40.3–60.3%). Serum chemistry disclosed increases in activity of alanine aminotransferase (515 U/L; reference range 16–91 U/L), aspartate aminotransferase (736 U/L; reference range 23–65 U/L), and creatine kinase (CK, 9,758 U/L; reference range 46–467 U/L). Serum titers for Toxoplasma gondii, Neospora caninum, Ehrlichia canis, Ehrlichia equi, and Borrelia burgdorferi were all negative. A serum acetylcholine receptor (AChR) antibody titer was negative. Testing for antibodies against masticatory muscle type 2M fibers was not performed. Serum total T4 and cortisol levels were normal. Thoracic radiographs revealed megaesophagus and an interstitial pattern in the right middle lung lobe consistent with aspiration pneumonia. Abdominal ultrasound disclosed an enlarged, hyperechoic liver, a small nodule in the spleen, and abdominal lymphadenopathy. A trichogram, skin scrape, skin and ear cytology, and fungal culture of epilated hair revealed no etiology for the skin lesions. Chronic superficial pyoderma and folliculitis were observed in skin biopsies. Supportive care consisted of polyionic fluids administered IV and feeding small meatballs of canned dog food. Antibiotic therapy (amoxicillin-clavulanateg; dose not recorded) was started for the suspicion of aspiration pneumonia based on thoracic radiographs. Electromyography (EMG) was not performed. Because of a poor quality of life, the owner elected euthanasia. A necropsy was performed, and on microscopic examination severe, chronic esophageal, myocardial, and skeletal (triceps, temporalis muscles) myositis with necrosis, and fibrosis, was identified (Fig 2A–C). The inflammatory infiltrates in the affected muscles were characterized as lymphoplasmacytic and histiocytic.
A 5-year-old castrated male Vizsla was presented for evaluation of masticatory muscle atrophy, difficulty drinking, and reduced physical activity of 2 months duration. The dog had been treated previously with cyclosporineh (5.17 mg/kg PO q24h) and chloramphenicoli (51.7 mg/kg PO q8h) prescribed by a dermatologist for alopecia and pododermatitis. Marked increase in thirst, ptyalism while drinking, masticatory muscle (temporalis and masseter) wasting, a change in bark, and lethargy were all noticed within days of starting the cyclosporine. Discontinuation of the cyclosporine resulted in improvement in attitude and appetite. Although the atrophy did not progress, muscle mass did not return. Previous diagnostic testing performed by the primary veterinarian included CBC and serum chemistry analysis, which were within normal limits, including CK; measurement of serum antibody titers for N. caninum and T. gondii, which were negative; measurement of antinuclear antibody titer, which was negative; assessment of thyroid function, which was consistent with euthyroid sick syndrome (serum T4 was low [0.4 μg/dL; reference range 1–4 μg/dL] with normal free T4 [21.6 pmol/L; reference range 8–40 pmol/L], and serum thyroid stimulating hormone concentration [0.16 ng/mL; reference range 0–0.60 ng/mL]); and a positive contrast esophagram, which indicated esophageal dysfunction. On examination, the dog had moderate, bilateral temporalis, and masseter muscle atrophy with normal range of jaw motion. A decreased gag reflex was noted. When offered food, swallowing of a food bolus subjectively appeared delayed, consistent with dysphagia. Right flank alopecia and bilateral thoracic limb pododermatitis also were present. A focal grade I/VI murmur was auscultated with a point of maximal intensity over the left heart apex. Serum titers for B. burgdorferi (via quantitative C6 antibody), E. canis, Anaplasma phagocytophilum, and Rickettsia rickettsii were negative. Serum AChR and 2M antibody titers were negative. Thoracic radiographs and abdominal ultrasound were unremarkable. Echocardiography was normal. EMG revealed positive sharp waves and fibrillation potentials in all appendicular muscles tested (left interosseous-pelvic, gastrocnemius, cranial tibial, quadriceps, semimembranosus, infraspinatus, supraspinatus, triceps, biceps, flexor carpi radialis, extensor carpi radialis, interosseous-thoracic). Positive sharp waves and fibrillation potentials were also noted in the temporalis muscles, the tongue and pharynx. Epaxial muscles along the vertebral column were normal. Histopathologic evaluation was performed on fresh frozen muscle biopsy specimens obtained from the cranial tibial, temporalis, and triceps muscles. Abnormalities were most severe in the cranial tibial and temporalis muscles. Microscopic changes included excessive variability in myofiber size, numerous atrophic fibers having a polygonal to anguloid shape of both fiber types, and multifocal areas of mixed mononuclear inflammatory cell infiltration having an endomysial distribution with invasion of nonnecrotic fibers. Immunofluorescent staining was performed on cryosections of the triceps muscle using previously characterized monoclonal antibodies against canine leukocytej and major histocompatibility complex (MHC)k antigens.5 Scattered CD4+ T cells with lesser numbers of CD8+ T cells and rare CD21+ B cells and CD1c positive macrophage/dendritic cells were identified with increased expression of MHC Class I and Class II antigens (Fig 3). Additional stains were performed using various monoclonal and polyclonal antibodies against muscular dystrophy associated proteins including dystrophin, laminin α2, spectrin, utrophin, α, β, and γ sarcoglycans, caveolin 3, dysferlin, and emerin.l Intensity of staining for all dystrophy associated proteins was similar to control tissue (not shown).
Therapy with prednisonem (1 mg/kg PO q12h) was initiated and the dog appeared to improve within 2 weeks. The prednisonem was tapered slowly over 6 months to a maintenance dose (0.5 mg/kg PO q48 h). At the 9-month recheck the dog had a normal gait and strength with good appendicular muscling. The masticatory muscles were less atrophied than previously but moderate atrophy remained. Jaw function, jaw range of movement, and the ability to swallow were normal with no apparent dysphagia.
A 2-year-old castrated male Vizsla was presented for atrophy of the temporalis and masseter muscles, ptyalism, and dysphagia. Six months prior, the dog had been evaluated for regurgitation by the same hospital. At that time, the serum CK activity was increased (591 U/L; reference range 63–350 U/L). Endoscopic assisted biopsy specimen and histopathologic evaluation of the gastric and duodenal mucosa revealed multifocal lymphonodular gastritis and moderate diffuse plasmacytic enteritis. Spiral-shaped bacteria consistent with Helicobacter sp. also were observed. Empirical treatment for helicobacter-induced gastritis was initiated along with symptomatic therapy for regurgitation and ptyalism. Treatment consisted of metoclopramide,b famotidine,n sucralfate,o omeprazole,p metronidazole,c and amoxicillin-clavulanateg (doses not recorded). Regurgitation resolved with therapy. Six months later the dog developed bilateral atrophy of the temporalis and masseter muscles. There was bilateral enophthalmus secondary to atrophy of the masticatory muscles. CBC and serum chemistries, including CK activity, were within reference ranges. Serum titers for T. gondii and N. caninum were negative. Testing for AChR antibodies was not performed. The serum titer for antibodies against type 2M fibers was negative. Thoracic radiographs were normal and abdominal imaging was not performed. EMG (temporalis and appendicular muscles) performed under general anesthesia revealed no abnormal spontaneous activity. Histopathologic evaluation performed on fresh frozen muscle biopsy specimens obtained from the temporalis muscle revealed moderate variability in fiber size with atrophy involving both fiber types. More than 50% of the muscle fibers contained internal nuclei. Multifocal areas of mild to moderate mononuclear, predominately lymphocytic, cellular infiltrates with an endomysial, perimysial, and perivascular distribution were present. Immunofluorescence staining for canine leukocyte antigens showed a pattern similar to Case 2 with CD4+ T cells in greater number than CD8+ T cells (not shown). The dog was treated with prednisonem (2 mg/kg PO q12h). Progressive atrophy of the masticatory muscles and ptyalism continued over the next month. Consequently, azathioprinen (2 mg/kg PO q24h for 7 days followed by 2 mg/kg PO q48h) was initiated adjunctively with prednisone therapy. Clinical signs stabilized and the dog was maintained on prednisonem (1 mg/kg PO q24h) and azathioprineq (2 mg/kg PO q48h) for 6 months with no further progression. The dog was euthanized 1 year after diagnosis for unrelated reasons.
Fourteen Hungarian Vizslas in the United Kingdom have been described with a unique form of polymyositis.3 The clinical signs in affected dogs included ptyalism, dysphagia, marked temporalis, and masseter muscle atrophy, increase in serum CK activity, and megaesophagus.3 Generalized loss of muscle mass and exercise intolerance were also described.3 The clinical signs observed in the dogs reported here were similar to those reported in the United Kingdom. Males have been affected (all dogs of this report; 10/14 in the United Kingdom3) more than females. However, a definitive predisposition for male dogs cannot be determined based on the small sample size. The dogs in this report were older (2, 5, and 9 years of age) than those in the United Kingdom3 (mean age 14 months, range 9–30 months), but similar in age to dogs with generalized inflammatory myopathies (mean age 5.09 years) as reported by Evans et al.2 Serum CK activity was increased in 85% of the Hungarian Vizslas reported from the United Kingdom3 but inconsistently observed in the dogs described here. In dogs from both the United Kingdom and the United States, the apparent distribution of affected musculature based on clinical signs and clinicopathologic findings combined with the exclusion of other disease processes suggests a unique breed-associated syndrome of polymyositis manifested primarily in the masticatory, pharyngeal, and occasionally the esophageal musculature.
The diagnosis of polymyositis in the Hungarian Vizsla is based on exclusion of other conditions with similar signs and confirmation of cellular infiltrates within muscle biopsies. Other neuromuscular diseases, such as myasthenia gravis and masticatory myositis, should be excluded based on serum AChR and masticatory muscle type 2M antibody titers, respectively. Infectious causes of myositis should be excluded based on serology. Muscle inflammation may be a prominent feature in some muscular dystrophies such as dysferlin and dystrophin deficiency.6 These forms of muscular dystrophy were ruled out by immunohistochemical staining of frozen muscle biopsy sections using available monoclonal and polyclonal antibodies. Interestingly, EMG of the affected muscles did not reveal abnormalities in all dogs. However, the presence of EMG abnormalities in cases of polymyositis depends on disease duration and the degree and distribution of inflammation within the muscle at the time of testing. Magnetic resonance imaging may be useful as an adjunctive modality in establishing a diagnosis.7 Although hyperintensity within the muscle on T2-weighted images represents nonspecific abnormalities, such changes may be beneficial in determining sites for muscle biopsy.7 Ultimately, definitive diagnosis requires biopsy of affected muscles for microscopic evaluation. Histopathology reveals a mixed mononuclear inflammatory cell infiltrate with an endomysial or perimysial distribution or both, typical of polymyositis. Most of the muscle biopsies in the dogs reported here were of the temporalis muscle; however, inflammatory infiltrates were found in other sites, consistent with a more widespread distribution. The presence of CD4+ T cells in excess of CD8+ T cells suggests that the pathogenesis of Vizsla polymyositis differs from that of other reported cases of canine polymyositis in which CD8+ T cells predominated.5,8 In canine masticatory muscle myositis, a focal inflammatory myopathy with distinct differences from polymyositis, CD4+ T cells are also in excess of CD8+ T cells.j As shown in two of our cases, increased expression of MHC Class I and Class II antigens in muscle biopsies, even in areas remote from inflammation, aids in the diagnosis of polymyositis when the diagnosis is suspected.8–10
The underlying etiopathogenesis for polymyositis remains undetermined. In humans, genetic predispositions for the development of polymyositis and other autoimmune diseases are suspected based on familial clusters of affected individuals.1,11 Genes coding for the MHC are thought to be involved.1 While certain alleles seem to increase an individual's genetic risk, it is likely that multiple genetic and environmental factors play a role in the development of disease.1 Muscle fibers do not normally express MHC Class I and Class II antigens, so the widespread overexpression of these antigens suggests muscle plays an active role in the immune response. Affected families are useful for the study of autoimmune disease as individuals within the family have similar genetic make-ups and environmental exposures. Dogs of the same breed have similar genetic makeup but usually have disparate environmental exposures.
Overlap syndromes in which individuals are affected by more than one autoimmune disease have been described in humans12 and dogs.2,13,14 This is of interest in the Hungarian Vizsla breed-associated polymyositis as dogs had inflammatory disease suggestive of an autoimmune pathogenesis identified in tissue other than skeletal muscle. One dog (Dog 1) had inflammatory infiltrates within the cardiac musculature. Another dog (Dog 3) was diagnosed with inflammatory gastrointestinal disease 6 months before diagnosis with polymyositis. While not verified histopathologically, an immune-mediated dermatopathy was diagnosed by a dermatologist in 1 dog and improvement was noted concurrent with the initiation of immunosuppressive prednisone therapy. These findings suggest that the autoimmune inflammatory response in affected Vizsla dogs may not be limited to just skeletal muscle.
aLevothyroxine, generic, Vet A Mix, Shenandoah, IA
bMetoclopramide, generic, Barr Labs, Pomona, NY
cMetronidazole, generic, Pliva Krakow Pharmaceutical Company, SA Krakow, Poland; distributed by Barr Labs
dVitamin Research Products, Boynton Beach, FL
eBaytril, Bayer, Shawnee Mission, KS
fCerenia, Pfizer, New York, NY
hAtopica, Novartis, Greensboro, NC
iChloramphenicol, generic, Osborn (Bimedia, a division of Cross Vepharm Group Ltd) Bimedia Inc, Le Sueur, MN
jPeter Moore, University of California, Davis, Davis, CA
kMHC, Class I from VRMD, Pullman, WA and Class II from Peter Moore
lAll antibodies obtained from Novocastra, Newcastle-upon-Tyne, with the exception of α-sarcoglycan and laminin α-2 which were obtained from Dr Eva Engvall, Sanford-Burnham Institute for Medical Research, La Jolla, CA
mPrednisone, generic, Qualitest Pharmaceuticals, Huntsville, AL
nFamotidine, generic, Wockhardt LTD Mumbai, India, distributed by Wockhardt USA LLC, Parsippany, NJ
oSucralfate, generic, Teva Pharmaceutical USA, Sellersville, PA
pOmeprazole, generic, Mylan Pharmaceuticals Inc, Morgantown, WV
qImuran, Faro Pharmaceuticals, Bedminster, NJ
The authors acknowledge the Muscular Dystrophy Association for funding in the laboratory of one of the authors (GDS).