Background: The diagnosis of equine protozoal myeloencephalitis (EPM) relies heavily on the clinical examination. The accurate identification of neurologic signs during a clinical examination is critical to the interpretation of laboratory results. Objective: To investigate the level of agreement between board-certified veterinary internists when performing neurologic examinations in horses. Animals: Ninety-seven horses admitted to the Veterinary Teaching Hospital at The Ohio State University from December 1997 to June 1998. Methods: A prospective epidemiologic research design was used. Horses enrolled in the study were examined by the internist responsible for care of the horse, and later by an internist who was not aware of the presenting complaint or other patient history. Data were analyzed by descriptive statistics, and kappa (K) statistics were calculated to assess interobserver agreement. Results: Ninety-seven horses were enrolled in the study. Overall, examiners, also referred to as observers, agreed that 60/97 (61.9%) were clinically abnormal, 21/97 (21.6%) were clinically normal, and the status of 16/97 (16.5%) of horses was contested. There was complete agreement among the examiners with regard to cranial nerve signs and involuntary movements. Disagreement involving severity of clinical signs occurred in 31 horses, and 25 of those horses (80.6%) were considered either normal or mildly affected by the primary observer. When examining the results of all paired clinical examinations for 11 different categories, there was wide variability in the results. When examiners rated the presence or absence of any neurologic abnormalities, lameness, or ataxia, the agreement among observers was either good or excellent for 80% of horses. When assessing truncal sway, the agreement among observers was good or excellent for 60% of the horses. When examining the horses for asymmetry of deficits, agreement was either good or excellent for 40% of the horses. Agreement among observers was excellent or good for only 20% of the horses when assessing muscle atrophy, spasticity (hypermetria), and overall assessment of the severity of neurologic abnormalities. Conclusions and Clinical Importance: This study underscores the subjectivity of the neurologic examination and demonstrates a reasonable level of agreement that may be achieved when different clinicians examine the same horse.

Triazine-based antiprotozoal agents are known for their lipophylic characteristics and may therefore be expected to be well absorbed following oral administration. However, although an increase in lipid solubility generally increases the absorption of chemicals, extremely lipid-soluble chemicals may dissolve poorly in gastrointestinal (GI) fluids, and their corresponding absorption and bioavailability would be low. Also, if the compound is administered in solid form and is relatively insoluble in GI fluids, it is likely to have limited contact with the GI mucosa, and therefore, its rate of absorption will be low. Based on the above considerations, we sought a solvent with low or no toxicity that would maintain triazine agents in solution. As the oral route is most preferred for daily drug therapy, such a solvent would allow an increased rate of absorption following oral administration. In present study, it was demonstrated that dimethylsulfoxide (DMSO) increased the oral bioavailability of toltrazuril sulfone (Ponazuril) threefold, relative to oral administrations of toltrazuril sulfone suspended in water. The cross-over study of toltrazuril sulfone formulated in DMSO indicated that the absolute oral bioavailability of toltrazuril sulfone in DMSO is 71%. The high bioavailability of the DMSO-preparation suggests that its daily oral administration will routinely yield effective plasma and cerebral spinal fluid (CSF) concentrations in all horses treated. Also, this improved formulation would allow clinicians to administer loading doses of toltrazuril sulfone in acute cases of Equine Protozoal Myeloencephalitis. Another option would involve administration of toltrazuril sulfone in DMSO mixed with feed (1.23 kg daily dose) meeting the US Food and Drug Administration (FDA) recommendations for the levels of DMSO permissible in pharmaceutical preparations.

Diclazuril is a triazine-based antiprotozoal agent which may have clinical application in the treatment of equine protozoal myeloencephalomyelitis (EPM). In this study, the use of the sodium salt diclazuril to increase the apparent bioavailability of diclazuril for the treatment and prophylaxis of EPM and various other Apicomplexan mediated diseases is described. In this study, diclazuril sodium salt was synthesized and administered to horses as diclazuril sodium salt formulations. The absorption, distribution, and clearance of diclazuril sodium salt in the horse are described. Diclazuril was rapidly absorbed, with peak plasma concentrations occurring at 8-24 hours following an oral mucosal administration of diclazuril sodium salt. The mean oral bioavailability of diclazuril as Clinacox was 9.5% relative to oral mucosal administration of diclazuril sodium salt. Additionally, diclazuril in DMSO administered orally was 50% less bioavailable than diclazuril sodium salt following an oral mucosal administration. It was also shown that diclazuril sodium salt has the potential to be used as a feed additive for the treatment and prophylaxis of EPM and various other Apicomplexan mediated diseases.

Sarcocystis neurona is the primary causative agent of equine protozoal myeloencephalitis (EPM), a common neurologic disease of horses in the Americas. We have developed a set of enzyme-linked immunosorbent assays (ELISAs) based on the four major surface antigens of S. neurona (SnSAGs) to analyze the equine antibody response to S. neurona. The SnSAG ELISAs were optimized and standardized with a sample set of 36 equine sera that had been characterized by Western blotting against total S. neurona parasite antigen, the current gold standard for S. neurona serology. The recombinant SnSAG2 (rSnSAG2) ELISA showed the highest sensitivity and specificity at 95.5% and 92.9%, respectively. In contrast, only 68.2% sensitivity and 71.4% specificity were achieved with the rSnSAG1 ELISA, indicating that this antigen may not be a reliable serological marker for analyzing antibodies against S. neurona in horses. Importantly, the ELISA antigens did not show cross-reactivity with antisera to Sarcocystis fayeri or Neospora hughesi, two other equine parasites. The accuracy and reliability exhibited by the SnSAG ELISAs suggest that these assays will be valuable tools for examining the equine immune response against S. neurona infection, which may help in understanding the pathobiology of this accidental parasite-host interaction. Moreover, with modification and further investigation, the SnSAG ELISAs have potential for use as immunodiagnostic tests to aid in the identification of horses affected by EPM.

Neurologic disease in horses caused by Sarcocystis neurona is difficult to diagnose, treat, or prevent, due to the lack of knowledge about the pathogenesis of the disease. This in turn is confounded by the lack of a reliable equine model of equine protozoal myeloencephalitis (EPM). Epidemiologic studies have implicated stress as a risk factor for this disease, thus, the role of transport stress was evaluated for incorporation into an equine model for EPM. Sporocysts from feral opossums were bioassayed in interferon-gamma gene knockout (KO) mice to determine minimum number of viable S. neurona sporocysts in the inoculum. A minimum of 80,000 viable S. neurona sporocysts were fed to each of the nine horses. A total of 12 S. neurona antibody negative horses were divided into four groups (1–4). Three horses (group 1) were fed sporocysts on the day of arrival at the study site, three horses were fed sporocysts 14 days after acclimatization (group 2), three horses were given sporocysts and dexamethasone 14 days after acclimatization (group 3) and three horses were controls (group 4). All horses fed sporocysts in the study developed antibodies to S. neurona in serum and cerebrospinal fluid (CSF) and developed clinical signs of neurologic disease. The most severe clinical signs were in horses in group 1 subjected to transport stress. The least severe neurologic signs were in horses treated with dexamethasone (group 3). Clinical signs improved in four horses from two treatment groups by the time of euthanasia (group 1, day 44; group 3, day 47). Post-mortem examinations, and tissues that were collected for light microscopy, immunohistochemistry, tissue cultures, and bioassay in KO mice, revealed no direct evidence of S. neurona infection. However, there were lesions compatible with S. neurona infection in horses. The results of this investigation suggest that stress can play a role in the pathogenesis of EPM. There is also evidence to suggest that horses in nature may clear the organism routinely, which may explain the relatively high number of normal horses with CSF antibodies to S. neurona compared to the prevalence of EPM.

Equine protozoal myeloencephalitis (EPM) is a serious neurological disease of horses in the Americas. The protozoan most commonly associated with EPM is Sarcocystis neurona. The complete life cycle of S. neurona is unknown, including its natural intermediate host that harbors its sarcocyst. Opossums (Didelphis virginiana, Didelphis albiventris) are its definitive hosts. Horses are considered its aberrant hosts because only schizonts and merozoites (no sarcocysts) are found in horses. EPM-like disease occurs in a variety of mammals including cats, mink, raccoons, skunks, Pacific harbor seals, ponies, and Southern sea otters. Cats can act as an experimental intermediate host harboring the sarcocyst stage after ingesting sporocysts. This paper reviews information on the history, structure, life cycle, biology, pathogenesis, induction of disease in animals, clinical signs, diagnosis, pathology, epidemiology, and treatment of EPM caused by S. neurona.

Equine Veterinary EducationVolume 12, Issue 4 p. 195-200 Diclazuril and equine protozoal myeloencephalitis (EPM): a clinical report B. G. Bentz, B. G. Bentz Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA. 121 Mockingbird Hill Drive, Richmond, Kentucky 40475, USA.Search for more papers by this authorL. Dirikolu, L. Dirikolu Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. G. Carter, W. G. Carter Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. Saville, W. Saville The Ohio State University, College of Veterinary Medicine, Columbus, Ohio, USA.Search for more papers by this authorN. M. Williams, N. M. Williams Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. V. Bernard, W. V. Bernard Rood and Riddle Equine Hospital, Lexington, Kentucky, USA.Search for more papers by this authorC. Wulff-Strobel, C. Wulff-Strobel Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorC. B. Baker, C. B. Baker Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorS. McCrillis, S. McCrillis Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorS. Reed, S. Reed The Ohio State University, College of Veterinary Medicine, Columbus, Ohio, USA.Search for more papers by this authorJ. D. Harkins, J. D. Harkins Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorD. E. Granstrom, D. E. Granstrom Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorT. Tobin, T. Tobin Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this author B. G. Bentz, B. G. Bentz Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA. 121 Mockingbird Hill Drive, Richmond, Kentucky 40475, USA.Search for more papers by this authorL. Dirikolu, L. Dirikolu Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. G. Carter, W. G. Carter Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. Saville, W. Saville The Ohio State University, College of Veterinary Medicine, Columbus, Ohio, USA.Search for more papers by this authorN. M. Williams, N. M. Williams Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorW. V. Bernard, W. V. Bernard Rood and Riddle Equine Hospital, Lexington, Kentucky, USA.Search for more papers by this authorC. Wulff-Strobel, C. Wulff-Strobel Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorC. B. Baker, C. B. Baker Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorS. McCrillis, S. McCrillis Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorS. Reed, S. Reed The Ohio State University, College of Veterinary Medicine, Columbus, Ohio, USA.Search for more papers by this authorJ. D. Harkins, J. D. Harkins Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorD. E. Granstrom, D. E. Granstrom Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this authorT. Tobin, T. Tobin Department of Veterinary Science, The Maxwell H. Gluck Equine Research Center, University of Kentucky, Lexington, Kentucky, USA.Search for more papers by this author First published: 05 January 2010 https://doi.org/10.1111/j.2042-3292.2000.tb00040.xCitations: 1 AboutPDF ToolsExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinkedInRedditWechat Citing Literature Volume12, Issue4June 2000Pages 195-200 RelatedInformation

Objective:To determine the seroprevalence of antibodies to Sarcocystis neurona in horses residing in northern Colorado during 1995 and 1996.Design: Prevalence survey.Sample Population:Aliquots of serum were collected from 608 equids from samples submitted to the Veterinary Diagnostic Laboratory at Colorado State University for testing for antibodies to equine infectious anemia (EIA).Procedure:Sera were analyzed for the presence of antibodies to S. neurona using Western blot analyses. Information regarding age, gender, breed, county of origin and quarter of the year when the sample was collected was recorded for each animal from the EIA form. Data were analyzed using chi-square analysis and multiple logistic regression,Results:Seroprevalence was 33.6%. Gender and county of origin were not associated with seroprevalence. Variables that were associated with seropositivity included age, breed, and quarter of the year. Seroprevalence increased with age. The highest seroprevalence, 66.6%, was found in the group that represented ponies and non-horse equids. Stock breeds had a seroprevalence of 32.6% and hot-blooded breeds had a seroprevalence of 27.9%. Seroprevalence was lowest during the coldest months (20.1%).Clinical Implication:Data from this sample population indicate that exposure of horses to S. neurona in northern Colorado is less than that reported for eastern regions of the United States, The overall seroprevalence is similar to that reported in horses from eastern Oregon. Although additional data are necessary, this finding suggests that areas of lower seroprevalence in the Rocky Mountain states corresponds to areas of lower opossum density. The results of this survey further support the conclusion that although a negative serum antibody test for S. neurona in a horse with neurologic signs may help rule out equine protozoal myeloencephalitis (EPM) as a diagnosis, a positive serum test result alone, especially in a clinically normal horse, does not lead to a definitive diagnosis of EPM.

OBJECTIVE:To determine serologic prevalence of Sarcocystis neurona, Toxoplasma gondii, and Neospora caninum in horses in Brazil.DESIGN:Prevalence survey.ANIMALS:101 Thoroughbreds in Brazil.PROCEDURE:Blood samples were obtained from horses and tested for serum antibodies against S neurona by use of an immunoblot procedure with culture-derived S neurona merozoites as antigen, and for serum antibodies against T gondii and N caninum by use of a modified agglutination test with formalin-preserved tachyzoites and mercaptoethanol.RESULTS:Antibodies against S neurona and T gondii were detected in 36 and 16 of 101 horses, respectively. Cross-reactivity between antibodies against T gondii and S neurona was not detected. Antibodies against N caninum were not detected in any samples.CONCLUSIONS AND CLINICAL RELEVANCE:The high prevalence of antibodies against S neurona detected in clinically normal horses emphasizes the importance of examining CSF for antibodies when establishing a diagnosis of equine protozoal myeloencephalitis.

Diclazuril (4-chlorophenyl [2,6-dichloro-4-(4,5-dihydro-3H-3,5-dioxo-1,2,4-triazin-2-yl)pheny l] acetonitrile), is a benzeneacetonitrile antiprotozoal agent (Janssen Research Compound R 64433) marketed as Clinacox . Diclazuril may have clinical application in the treatment of Equine Protozoal Myeloencephalitis (EPM). To evaluate its bioavailability and preliminary pharmacokinetics in the horse we developed a sensitive quantitative high-pressure liquid chromatography (HPLC) method for diclazuril in equine biological fluids. MS/MS analysis of diclazuril in our HPLC solvent yielded mass spectral data consistent with the presence of diclazuril. After a single oral dose of diclazuril at 2.5 g/450 kg (as 500 g Clinacox), plasma samples from four horses showed good plasma concentrations of diclazuril which peaked at 1.077 +/- 0.174 microg/mL (mean +/- SEM) with an apparent plasma half-life of about 43 h. When this dose of Clinacox was administered daily for 21 days to two horses, mean steady state plasma concentrations of 7-9 microg/mL were attained. Steady-state levels in the CSF ranged between 100 and 250 ng/mL. There was no detectable parent diclazuril in the urine samples of dosed horses by HPLC or by routine postrace thin layer chromatography (TLC). These results show that diclazuril is absorbed after oral administration and attains steady-state concentrations in plasma and CSF. The steady state concentrations attained in CSF are more than sufficient to interfere with Sarcocystis neurona, whose proliferation is reportedly 95% inhibited by concentrations of diclazuril as low as 1 ng/mL. These results are therefore entirely consistent with and support the reported clinical efficacy of diclazuril in the treatment of clinical cases of EPM.

Seven members of a 15-man U.S. military team that had operated in rural Malaysia developed an acute illness consisting of fever, myalgias, bronchospasm, fleeting pruritic rashes, transient lymphadenopathy, and subcutaneous nodules associated with eosinophilia, elevated erythrocyte sedimentation rate, and elevated levels of muscle creatinine kinase. Sarcocysts of an unidentified Sarcocystis species were found in skeletal muscle biopsies of the index case. Albendazole ameliorated symptoms in the index case; however, his symptoms persisted for more than 5 years. Symptoms in 5 other men were mild to moderate and self-limited, and 1 team member with laboratory abnormalities was asymptomatic. Of 8 team members tested for antibody to Sarcocystis, 6 were positive; of 4 with the eosinophilic myositis syndrome who were tested, all were positive. We attribute this outbreak of eosinophilic myositis to accidental tissue parasitism by Sarcocystis.

Sarcocystis neurona is the etiologic agent of equine protozoal myeloencephalitis (EPM). Based on an analysis of 25,000 equine serum and cerebrospinal fluid (CSF) samples, including samples from horses with neurologic signs typical of EPM or with histologically or parasitologically confirmed EPM, four major immunoblot band patterns have been identified. Twenty-three serum and CSF samples representing each of the four immunoblot patterns were selected from 220 samples from horses with neurologic signs resembling EPM and examined for inhibitory effects on the infectivity of S. neurona by an in vitro neutralization assay. A high correlation between immunoblot band pattern and neutralizing activity was detected. Two proteins, Sn14 and Sn16 (14 and 16 kDa, respectively), appeared to be important for in vitro infection. A combination of the results of surface protein labeling, immunoprecipitation, Western blotting, and trypsin digestion suggests that these molecules are surface proteins and may be useful components of a vaccine against S. neurona infection. Although S. neurona is an obligate intracellular parasite, it is potentially a target for specific antibodies which may lyse merozoites via complement or inhibit their attachment and penetration to host cells.

OBJECTIVE:To determine the clinical findings, course of treatment, and long-term outcome of horses on a farm in central Kentucky during an epizootic of equine protozoal myeloencephalitis (EPM).DESIGN:Cohort study.ANIMALS:21 horses on a farm in central Kentucky, 12 of which developed clinical signs of EPM.PROCEDURE:Horses on the farm were serially examined for signs of neurologic disease and serum and CSF antibodies to Sarcocystis neurona. Horses were considered to have EPM if they had neurologic signs and positive test results for antibodies to S neurona in CSF. Blood values were monitored for evidence of abnormalities resulting from long-term pyrimethamine and trimethoprim-sulfamethoxazole administration Physical, neurologic, and fetal necropsy examinations were performed as needed. Horses were treated for EPM until they had negative test results for CSF antibodies to S neurona.RESULTS:Of 21 horses on the farm, 12 had EPM over the course of 6 months. The duration of treatment ranged from 45 to 211 days, excluding 1 horse that persistently had CSF antibodies to S neurona. Adverse effects from pyrimethamine and trimethoprim-sulfamethoxazole administration included transient fever, anorexia, and depression (n = 2); acute worsening of ataxia (2); mild anemia (4); and abortions (3).CLINICAL IMPLICATIONS:EPM may develop as an epizootic. In the horses of this report subtle clinical signs that were originally considered unimportant ultimately progressed to obvious neurologic signs. Adverse effects associated with EPM treatment included worsening of neurologic signs, anemia, abortion, and leukopenic and febrile episodes.

OBJECTIVE:To determine seroprevalence of antibodies to Sarcocystis neurona in neurologically normal horses residing in 4 regions of Oregon and to describe the effects of age, gender, breed, and housing on seroprevalence within each region.DESIGN:Prevalence survey.SAMPLE POPULATION:Serum samples from 334 horses systematically selected by practicing veterinarians.PROCEDURE:Antibodies to S neurona were measured in sera, using a western blot. Information including age, gender, breed, housing, geographic location, and duration of residence was obtained for each horse. Data were analyzed, using descriptive statistics.RESULTS:45% (149/334) of horses evaluated were seropositive for antibodies to S neurona with significant differences in the percentage of seropositive horses from different regions of the state. Seroprevalances of antibodies to S neurona in horses in regions I and II, west of the Cascade Range, were 65 and 60%, respectively; whereas seroprevalances in central and eastern Oregon, regions III and IV, were 43 and 22%, respectively. Seroprevalence consistently increased with age of horse for each region. Gender, breed, and housing were not associated with significant differences in seroprevalence of antibodies to S neurona in the overall sample population, or in comparisons of samples obtained from horses within a particular region, or among samples obtained from horses residing in different regions.CLINICAL IMPLICATIONS:The high seroprevalence of antibodies to S neurona in neurologically normal horses indicates that analysis of serum alone would not be useful for definitive diagnosis of equine protozoal myeloencephalitis in horses in Oregon.

We report a simple, economical, and efficient protocol for protein purification from cells. First, proteins of cell lysates were separated by standard sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and electroblotted to protein-blotting membrane. The blots were stained with Coomassie blue or developed by immunoblotting to visualize specific proteins. The bands corresponding to those visible by immunoblotting were excised from the dye-stained blots and subjected to isoelectric focusing. The focused gel was stained with Coomassie blue. Finally, the stained bands were excised and subjected to another SDS-PAGE separation and electrotransferred back to protein-blotting membrane. At this stage, the purified proteins were suitable for microsequencing. We have tested the feasibility of this novel technique by purifying proteins with molecular weights ranging from 19 to 100 kDa from a lysate of Sarcocystis neurona, the etiologic agent of equine protozoal myeloencephalitis. The purity of proteins was demonstrated by reverse-phase high-performance liquid chromatography. Partial sequences of these purified proteins were obtained by N-terminal or digestive sequencing.

The protein concentration in biological samples is a crucial limiting factor for a successful analysis by electrophoresis. Many techniques have been adopted to increase protein concentrations, however, they are often insufficient and require special equipment or poisonous chemicals. Herein, we report a simple and efficient technique for concentrating dilute protein samples by absorbing proteins onto protein-blotting membrane strips. In this technique, blotting membrane strips were incubated in dilute protein solutions to capture proteins. For either sodium dodecyl sulphate-polyacrylamide gel electrophoresis or isoelectric focusing, the protein-absorbed membrane strips were directly loaded to the sample wells which contained a strong protein elution buffer and electrophoresis was performed under standard conditions.

Data are presented on the last 3 years of a 7-year study (1989-1995) on transmission of natural infections of internal parasites in horse foals (n = 27) born in 1993, 1994, and 1995 on the same pasture on a farm in central Kentucky. The foals were in a closed breeding herd of horses. Research on the first 4 years (1989-1992) of the study was published earlier (Lyons et al., 1991, 1994). Thirty-five species of endoparasites were identified, including 24 species of small strongyles. Monthly, seasonal, and host-age transmission patterns were elucidated for the parasites. Comparison of data between the first 4 years and last 3 years of the study indicates similarities, but also differences, including an increase in prevalence and numbers of Thelazia lacrymalis and Anoplocephala perfoliata. (C) 1997 Elsevier Science B.V.

Sarcocystis sp. sporocysts isolated from eight feral opossums (Didelphis virginiana) were pooled and fed to 18 commercially reared budgerigars (Melopsittacus undulatus), 14 wild-caught sparrows (Passer domesticus), one wild-caught slate-colored Junco (Junco hyemalis) and five weanling horses (Equus caballus). All budgerigars died within 5 weeks post inoculation (wpi). Histologic examination revealed meronts within the pulmonary epithelia and typical Sarcocystis falcatula sarcocysts developing in the leg muscles. Sparrows were euthanized 13 and 17 wpi and their carcasses were fed to four laboratory raised opossums. Sporocysts were detected in the feces of two opossums on 15 days post inoculation (dpi) and in a third opossum on 40 dpi. Fecal samples from the fourth opossum remained negative; however, sporocysts were found in intestinal digests from all four opossums. Sporocysts were not found in the feces or intestinal digest of an additional opossum that was fed three uninoculated sparrows. Five foals were fed sporocysts (Foals 2, 3, 4, 5, and 7) and two foals were maintained as uninoculated controls (Foals 1 and 6). Sporocysts from two additional feral opossums also were fed to foals. Foal 5 was given 0.05 mg kg−1 dexamethasone sodium phosphate daily beginning 2 days before inoculation for a total of 2 weeks. Horse sera were tested three times per week, and cerebrospinal fluid (CSF) samples were tested biweekly for anti-Sarcocystis neurona antibodies by Western blot analysis. No foals had any S. neurona-specific antibodies by Western blot analysis prior to sporocysts ingestion. Seroconversion occurred in Foals 3, 5, and 7 by 24 dpi, followed by positive CSF tests on 28 dpi. Foals 2 and 4 seroconverted by 40 dpi. Cerebrospinal fluid from Foal 2 tested positive by 42 dpi, but Foal 4 remained seronegative throughout the study. Sera and CSF from control Foals 1 and 6 remained seronegative. All foals with positive CSF developed neurologic clinical signs. Neurologic disease was evident in Foals 2 and 3 by 42 dpi and in Foal 7 by 28 dpi. The severity of clinical signs progressed to marked spasticity, hypermetria and ataxia in Foal 7 by the end of the trial. Necropsy examination of inoculated foals did not reveal gross lesions; however, microscopic lesions consistent with equine protozoal myeloencephalitis (EPM) were found in Foals 2, 3, and 7. Protozoa were not observed in the tissue sections. Microscopic lesions consistent with EPM were not found in Foals 4 and 5 or in uninoculated control Foals 1 and 6. Foal 5 had unilateral non-inflammatory lesions in the cervical and thoracic spinal cord consistent with cord compression. These data indicate that the opossum is a definitive host of S. neurona.