editorial board
past issues
contact us


printer friendly page    email page

2011, Vol. 6 No. 2, Article 89


Equine Viral Arteritis – A Review

S. Nandi* and Manoj Kumar


Centre for Animal Disease Research and Diagnosis,
Indian Veterinary Research Institute, Izatnagar, U. P.


*Corresponding Author; e-mail address: snandi03@yahoo.com



Equine viral arteritis (EVA) is an infectious disease characterized by panvasculitis inducing edema, hemorrhage, and abortion in pregnant mares. The disease is caused by equine arteritis virus (EAV) which belongs to the genus Arterivirus and family Arteriviridae. Only one serotype of EAV has been identified so far but antigenic variation among the virus isolates has been reported which vary in pathogenicity. The severity of clinical disease is high in horses infected with the Bucyrus strain followed by Kentucky strain and Penn strain. These variants also have different abortogenic potential. The transmission commonly occurs through aerosol and venereal route. Venereal transmission plays a significant role in the widespread dissemination of the virus by carrier stallions as the EAV has been found constantly in the semen but the mares did not establish the carrier state. Control of disease can be achieved by vaccination against EVA in horses by a cell culture based attenuated vaccine using the Bucyrus strain and proper and early diagnosis of carrier stallions.


Equine Viral Arteritis.


Equine arteritis virus (EAV) was first isolated from lung tissue of an aborted fetus during outbreak of equine arteritis among horses on a farm in Bucyrus, Ohio in 1953 (10) and in Indiana, California, Pennsylvania and Kentucky later (19,32). In spite of the apparent widespread prevalence around the world, infrequent epizooties have generally been reported from various parts of the United States and some of the European countries such as Poland (29), Switzerland (2) and Austria (24). Although no clinical case of equine arteritis has been reported in Germany, a serological survey indicated the presence of high anti-EAV antibody titres in 3.8% of horse serum samples tested. Similar observations have also been reported from various surveys in France, Argentina, Canada, Morroco, India and Netherlands (8,30). Equine viral arteritis (EVA) is a global infectious disease of horses and is characterized by panvasculitis inducing edema, hemorrhage and abortion in pregnant mares (20). Natural outbreaks of EVA are characterized by transient clinical signs and abortion in pregnant mares contrast with experimental disease, which features high mortality and prominent systemic vascular necrosis. Blood vessel cells are the major, but not exclusive, target of EAV. Lung, intestine, kidney, the reproductive tract, and occasionally the placenta are important viral replication sites, which favor the spread of the virus. Considerable importance has been expressed over the economic severity and veterinary significance of the disease after the epizootic of EVA in Kentucky in 1984 (49). The threat of outbreaks of abortion and potential for establishment of the carrier state in stallion has become a major concern. The name ‘Viral arteritis’ has been given for this hitherto unrecognized disease because the main pathological change is a hyaline necrosis in the tunica media of small muscular arteries.


The causal agent of EVA has been classified as the member of the genus Arterivirus and family Arteriviridae. EAV is an enveloped, spherical and positive-stranded RNA virus with a diameter of 50–70 nm and molecular weight of 4×106 (54). The virion is comprised of an isometric core surrounded by a lipid-containing envelope containing delicate spikes of 12 to 15 nm in diameter (21,41). The viral genomic RNA, which is encapsulated by a single nucleocapsid protein, is contained within the core particle. The arterivirus replication strategy resembles that of the family Coronaviridae (44). The genome of EAV is 12.7 kb and consists of seven open reading frames (ORFs). ORFs 2, 5, 6 and 7 encode the four structural proteins of the virus, of which a nucleocapsid protein (Molecular weight 12 Kd), a non-glycosylated envelope protein (Molecular weight 14 Kd) and a glycosylated envelope protein having molecular weight of 21Kd and a minor protein (21,55,61), whereas ORFs 1a and 1b encode the viral polymerase. EAV is an ether sensitive RNA virus which is resistant to trypsin but inactivated by MgCl2 at 50˚C (48). The buoyant density of infectious virus is 1.17 to 1.24 g/ml (22).


So far only one serotype of EAV has been identified but evidence of limited antigenic variation among the virus isolates have been reported (13). The pathogenicity of strains of EAV vary considerably and clinical disease is often severe. The severity of disease is high (often resulting in death) in horses infected experimentally with the Bucyrus strain than those infected naturally or by transmission of infection by carrier stallions. A strain isolated from the spleen of a naturally infected horse in a relatively mild epizootic in Penn (Penn strain) caused a milder disease than Bucyrus strain in experimental infection of horses or Kentucky strain isolated in 1984 (38). Vaccination of horses with isolates from different epizooties showed about equally protective ability suggesting the prevalence of a single serotype with some antigenic variation among different isolates. Antigenic variation has also been evident after analyzing EAV strains using monoclonal antibodies (59). The EAV variants having different abortogenic potential are also prevalent in the field. However, other factors, such as differences in susceptibility of various breeds of horses to EAV infection should also be taken into account. It is still unclear how the virus is maintained between the outbreaks and what factors are essential to cause an epizootics. Perhaps the generation and selection of virus having increased virulence play the pivotal role. The incidence of EVA has not been observed in U.K., Japan and Netherlands though it is widespread in endemic areas (20).


The primary mode of transmission has been suggested to be aerosol via the respiratory route. This had been proved in both natural and experimental infections. Subsequently, it became apparent that transmission of EAV via venereal route also played a significant role in the widespread dissemination of the virus (42,50). The virus may persist for a long period of time in the kidney of infected horse and may be spread directly or indirectly by contaminating the surrounding with urine. The importance of indirect contact through medium of virus contaminated fomites varied from outbreak to outbreak cannot be overlooked. The extent of lateral spread of EAV infection although not a notable feature but possibly plays a significant role in transmission of the disease among the horses having close association at a race track or horse show and green pasture. The virus also passes across the placenta from an acutely infected mare to her unborn foal (36,38).


Within two days of aerosol infection, EAV is rapidly spread within the lung and bronchial lymph nodes, and then disseminated throughout the body via the circulation (38). After experimental intranasal infection with the Bucyrus strain of EAV, the initial replication of the virus takes place in macrophages of the lung. The virus then spread to bronchial lymph nodes followed by the dissemination throughout the body via circulating system. Viral antigen can be detected practically in all the tissues, except the brain. The primary cells supporting EAV replication are macrophages present in all tissues. Secondary sites of replication have been identified as medial cells, mesothelium, endothelial cells and the epithelium of certain organs like adrenals (7,43).
The sites of paramount importance to the vascular lesion development were the endothelium and medial cells. Endothelial damage and infiltration of neutrophils was followed by damage (six days after infection) to the internal elastic laminar of the small arteries. In horses, which survived infection and recovered clinically, lesions in the vascular and lymphatic systems had largely subsided by 10 days after infection while arterial damage persisted for several weeks (6). Infectious EAV persisted in the kidney for up to 19 days and was isolated from the urine 16 days after infection. This correlates with histopathological evidence of prolonged glomerulonephritis in recovering animals and evidence from immunofluorescence studies for prolonged viral replication in the kidney, up to 14 days after infection. An immunological component to the pathogenesis has been postulated but is as yet unsubstantiated (7). The mechanism of abortion following EAV infection is not properly understood. It has been suggested that although fetal death may occur in uterus during acute viral infection (5), abortion is probably caused by lesions in the uterus of the mares (40,58). The presence of EAV in the reproductive tract of the infected mare has not been as well characterized as in the stallion.


The development of a clinical or inapparent infection in horse depends on the size of the viral challenge and the strain of the EAV. Outbreaks in Switzerland (1,16) and Austria (24) were of a milder form than that originally described (10). In the outbreak which occurred in a riding stable in Switzerland in 1975 (16), respiratory signs were absent apart from the occasional individual with elevated respiratory rates.
In 1977, a mild form of the disease was identified at two Kentucky race courses. Affected horses developed fever, scrotal, palpebral and leg edema, leucopenia and mild anemia (37). In natural cases of EVA, clinical signs vary considerably in range and clinical severity. Typical signs include pyrexia for 4-9 days with temperature ranging between 39-41˚C following an incubation period of 5-10 days (usually 6-8 days after venereal exposure). This is usually accompanied by anorexia, depression, nasal and ocular discharges, conjunctivitis, rhinitis, congested nasal mucosa, respiratory distress and leucopenia. Other frequently observed signs are photophobia, opacity of the cornea, coughing, diarrhea, limb edema, palpebral edema, edema of the mammary gland or scrotum and the prepuce and skin rash most commonly on the neck but sometimes generalized depression, muscular weakness, prostration, ataxia, enteritis, colic and abortion in pregnant mares. It is emphasized that abortion induced by arteritis virus occurs 12-30 days after infection. Signs were generally less severe in barren mares and weanling fillies. Again, clinical signs in adults were mild and characterized by slight fever, apathy, occasional colic and respiratory signs. More pronounced signs noticed in foals were anorexia, respiratory signs, pneumonia, colic and diarrhea preceded death. Regardless of the severity of the illness, naturally infected horses invariably make uneventful clinical recoveries. Mortality has only been observed in horses experimentally exposed to unattenuated Bucyrus strain of the virus and varies rarely in sporadic cases of naturally acquired infection in foals of few days to few weeks of age (10,17,46).


Pathological lesions may consist of edema, congestion and hemorrhages of subcutaneous tissues, lymph nodes and viscera of the peritoneal and pleural cavities (10). In addition, congestion and hemorrhages of the upper respiratory tract, bronchopneumonia, edema of larynx and mediastinal tissues, gross effusion of fluid into the peritoneal and pleural cavities, catarrhal enteritis, degeneration of the liver and kidney and hemorrhages and infarction of the spleen. Phlebitis, adrenal necrosis and endothelial changes were described by Estes and Cheville (1970) (12). Multifocal necrotizing myometritis has been observed in the uterus of the pregnant mares aborted 11 days post infection having 5-6 months pregnancy (56). Sub mucosal edema of the uterus and the edema of the broad ligament are significant in pregnant mares. Vascular lesions are not usually apparent in aborted fetuses (33). Aborted fetuses are usually partially autolyzed; the only visible abnormalities may be excessive fluid in the body cavities and interlobular interstitial pneumonia.


Common microscopic lesions in the maternal reproductive organs indicated myometritis with a degeneration of the myocytes and infiltration of the mononuclear cells. Epithelial cells of endometrial glands showed sporadic degeneration. Lesions in the fetal tissue include atrophy of lymphoid follicles in the spleen and lymph nodes with degenerated lymphocytes. The placenta was edematous in the sub villous layers (56,58).
In terminal stages of experimentally infected horses, widespread necrotizing arteritis affecting the media of the muscle arteries is apparent. Affected arterial musculature loses its nuclei and becomes hyaline and acidophilic in nature. Edema of the adventitia of the artery and lymphocytic infiltration in adventitia and media are evident (25).


The most complicating factor in the dissemination of EAV in any horse population is the existence of the carrier state in the stallion. It has been documented that a percentage of the stallions persistently infected with the EAV, were constant shedders of the virus in the semen (52,53). Both long term and short term carriers were shown to exist and frequency of the former group varies considerably among the different groups of stallions. The duration of carrier state can vary from a period of several weeks to years and perhaps for the rest of the life time in some stallions (20).
In carrier stallions, the EAV has been found constantly in the semen but it has not been detected either in nasopharyngeal secretions, urine or in the buffy coat. Unlike the stallion, the mares did not establish the carrier state. The possibility of transplacental transmission of EAV to the foal in uterus with the development of congenitally acquired carrier state and EAV induced teratogenic abnormalities born of mares pregnant at the time of infection or vaccination against EVA has not been demonstrated (51,52,53). However, it is now evident that carrier stallion probably plays a significant role in the perpetuation of the virus from year to year. Establishment of clinical or unapparent infection in the mare after getting infection from a carrier stallion via venereal route produced a state of profuse virus shedding from respiratory tract, which in turn becomes a source of secondary horizontal transmission to susceptible animals (14). The development of carrier state in colts and the castrated animals have not been successful indicating the essential role of testosterone for the induction of carrier state (26,39).


EAV was shown to replicate in primary horse, rabbit and hamster kidney cells. The development of cytopathic effect is initiated 3 days after incubation period and by the end of 6 days, most of the cells in the infected cultures show cytopathic shrinkage, pycnosis and detachment from the glass (18). EAV is found to replicate in a variety of cell lines e.g. hamster kidney cells (BHK21), rabbit kidney cells (RH-13, LLC-RK-1), African green monkey cells (Vero, B-SC-1), rhesus monkey cells (LLC- MK2) and a diploid line of equine dermal origin (NBL-6) with the production of plaques 2-4 mm in diameter. The maximum production of viral protein and RNA are observed at 35-37˚C between 6-8 hours and virus release is completed by 10-20 hours post infection. Electron microscopic studies indicated that EAV is released from infected cell is by a process of budding from cytoplasmic matrix into cisternae of the endoplasmic reticulum (26,35,57).


An assumptive diagnosis of EVA can be established based on the presence of the characteristic clinical signs of disease that include fever, depression, edema, conjunctivitis, nasal discharges, and abortions (43). The virus can be isolated from nasopharyngeal and conjunctival swabs, citrated blood and semen samples and confirmatory diagnosis is only available from serological tests viz., CFT and ELISA. The isolation of virus has been possible from a variety of necropsy material especially the lymph nodes associated with alimentary tract and related organs. The placental, lymphoreticular tissue and placental and fetal fluid can be productive source of virus in putative cases of EAV abortion.
Virus Isolation
Virus can be cultured in a variety of cells (equine kidney, rabbit kidney, monkey kidney) producing a CPE usually within 6 days of inoculation of cultures (36). Virus was transmitted to horses experimentally by intra peritoneal inoculation of blood from febrile horses to susceptible animals and subsequent isolation of virus achieved from nasal swabs and blood of inoculated horses. The isolation of EAV in equine kidney cells (1) and rabbit kidney and Vero cells (17,35) was successful by several blind passage of material from foals and aborted fetuses.
Antigen Detection
Immunofluorescence has been used to detect viral antigen in the tissues of experimental infected horses (7) using gamma-globulin conjugated to fluorescein (60). An indirect method using a rabbit antiserum prepared against EAV has been used to detect antigen in tissue culture (23). Immunohistochemistry is a reliable, powerful and rapid method to diagnose EAV infection in tissues and occasionally in skin biopsies (27,60). An avidin-biotin complex (ABC) immunoperoxidase staining using monoclonal antibodies to individual EAV proteins has been successfully used to detect viral antigens in formalin-fixed, paraffin-embedded tissues, as well as in frozen tissues sections. (9).
A wide variety of serological tests such as immunofluorescent (Wada et al., 1996), immunoperoxidase (28) and virus neutralization tests (14,47) has been used for the detection of antibody to EAV. Up to present time virus neutralizing test (by plaque reduction or in micro titre systems) in the presence of complement have been found to be the most sensitive than complement fixation, immunodiffusion and immunofluorescence tests (40,60). The complement fixation tests, although less sensitive may be more useful for diagnosing recent infections because complement fixation antibody titre decline faster than virus neutralizing antibody titre (13,14,15). It has been reported that high titres of neutralizing antibody can persist for several years after initial infection. More recently, ELISA has been applied to EAV. The test is as sensitive as VNT and provides results more quickly (4).
Molecular Biology Techniques
Several reverse transcription polymerase chain reaction (RT-PCR) assays, including, nested RT-PCR (RT-nPCR) and real-time RT-PCR assays have been developed for detection of the EAV nucleic acids in cell culture supernatants and clinical specimens. In these latest techniques, the single stranded viral RNA is converted to cDNA using reverse transcriptase enzyme followed by the amplification of the cDNA employing polymerase chain reaction. It is extremely sensitive and can detect the disease with high degree specificity even in the presence of a single copy of undamaged RNA in the clinical sample (3,55).


The differential diagnosis includes equine influenza, equine herpes virus 1 and 4 infections, equine infectious anemia, African horse sickness, Getah virus infection, purpura hemorrhagica and other streptococcal infections. Abortions caused by equine viral arteritis must be distin-guished from abortions caused by equine herpes virus 1 and 4 infections.


The immunoprophylaxis against EVA in horses could be achieved by an attenuated vaccine developed by propagating the virus in cell culture (30,31). The idea of developing the attenuated vaccine derived from the protective activity of anti-EAV antibodies. The Bucyrus strain was first passaged repeatedly through cultures of horse kidney cells and later through cultures of rabbit kidney cells and equine dermis cells. It was found that there is reduced virulence of the virus for horse revealed by the development of protective antibodies and no transmission from vaccinated animals to in contact susceptible animals. Additional cell passages further reduced the virulence of the virus and latest attenuated strains have been passaged 131 times in horse kidney cell cultures, 110 times in rabbit kidney cell cultures, and 19 or 25 times in equine dermal cell cultures (HK131-RK-110-AK19 or 25) (18,32,34). These strains conferred complete protection from clinical disease in response to challenge infection. The infectivity and immunizing potency of the lyophilized attenuated live virus vaccine containing 2-5% fetal bovine or horse serum were completely stable at -20˚C for one year and rapidly lost stability at room temperature or higher (11,31,32). Vaccine virus administered intramuscularly was not recovered from naso-pharyngeal secretion nor was vaccine virus transmitted to in-contact susceptible animals. Vaccine stimulated little or no hematological changes and only mild and irregular febrile reactions. Pathogenicity of vaccine virus has been reduced such that infection by the respiratory tract was unreliable and back passages of the attenuated virus did not restore pathogenicity. The dose of vaccine virus between 3×102 TCID50 and 5×107.5 TCID50 was used to immunize horses without causing unacceptable reactions (33).
Vaccination of pregnant mares late in gestation was undesirable because vaccine virus was recovered from a weak foal delivered by a vaccinated mare. Immunity provided by vaccine was examined in challenge infection experiments using 20% suspension of spleen tissue containing prototype Bucyrus strain and protection was assessed in terms of reduction in clinical signs, virus excretions and sero-conversions were monitored. The fact that vaccine allowed the viral replication in the absence of clinical signs was confirmed by the development of significant virus neutralization antibody responses following challenge even in individuals with high levels (VN titre >512) of antibody at the time of challenge (18). The killed or subunit vaccine has been developed as the exposure of the live attenuated vaccine immunized horse to wild type virus results in reinfection and transient secretion of virus. Subunit or killed vaccine induced serum neutralizing antibodies comparable to live attenuated vaccine and there were no abortions in immunized pregnant mares (15).


Neutralizing anti-EAV antibodies appear in horses within a week after natural or artificial infection. Early antibodies are mainly IgM types followed by IgG. IgG antibodies neutralizes EAV infectivity for cultured cells although inefficiently but efficiency can be greatly enhanced by the presence of complement (45). Neutralization of EAV by anti-EAV antibodies generated in a variety of other species is greatly enhanced indicating that the effect of complement is a property of the virions rather than horse IgG. It became evident that anti-EAV antibodies acquired by either natural or artificial infection protect the horses against clinical disease not from reinfection after challenge virus infection.


There is no systemic program to be available for the prevention and control of EVA. A mandatory serological survey of the horse population throughout the world to determine the prevalence of EAV infection and identification of long term carriers were the pre-requisite steps before launching any control strategy. It is of prime importance to prevent the further spread of EAV, a strategic vaccination of the sero-negative stallion with modified vaccine and halt in increase in the number of carrier stallions should be achieved. This would check the dissemination of the disease and minimizing the risk of getting infection by mares (20). Currently, it has been felt that effective means of preventing the EAV infection and possibly the carrier state establishment can be achieved by compulsory annual vaccination (19).
One of the most contentious issues is whether the confirmed carriers and semen shedders of EAV should be used subsequently for breeding purposes. According to some scientists, carrier stallions should be kept separately and bred only to mares vaccinated not less than three weeks or seropositive from natural exposure to EAV. After breeding, mares should be kept isolated from other non-vaccinated or sero-negative horses for a period of three weeks, whereas another group opined that carrier stallions should be slaughtered instead of using in breeding purposes (52,53). In addition, it is suggested to restrict the import of sero-positive studs and mares and semen from countries where the disease is prevalent by a country free from the disease.


Since the discovery of EAV in 1953, lot of information has been gathered about the disease and its devastating consequences on equines has been felt. An exhaustive research carried out over the years has provided a safe and effective vaccine and the retrospective epidemiological analysis has helped to provide additional insight about the modes of natural transmission of the virus and the establishment of carrier state. Although much has been learned about EVA, there is still no way to detect the carrier animals and a possible risk of disseminating the disease from a carrier or inapparently infected stallion to mares. Another controversial issue is whether the live modified vaccine should be used in large scale when vaccine is unable to check re-infection but only clinical disease. The answer of all questions still lies in the development of a potent, safe and efficacious vaccine and the early detection of carrier stallions. Constant research on EVA by scientists with a creative thinking and novel insight will find a possible breakthrough to defeat such a notorious and notifiable disease causing severe economic losses in near future.


    1. Burki F. Eigenschaften des virus der Equinen Arteritis. Pathol Microbiol. 1965;28: 939-949.

    2. Burki F. The virology of equine arteritis. Proc. 2nd Int. Conf. Equine Infect. Dis. Paris, 1969 pp: 125-129.

    3. Chirnside ED, Spann WJM. Reverse transcription and cDNA amplification by the polymerase chain reaction of equine arteritis virus. J Virol Meth. 1990;30:133-140.

    4. Chirnside ED, Francis PM, Vries De AAF, Sinclair R, Mumford, JA. Development and evaluation of ELISA using recombinant fusion protein to detect the presence of host antibody to EAV. J Virol Meth. 1995;54:1-13.

    5. Conignoul FL, Cheville NF. Pathology of the maternal genital tract, placenta and fetus in equine viral arteritis. Vet Pathol. 1984;21:333-340.

    6. Crawford TB, Hanson JB. Viral arteritis of horses. Adv Exp Med Biol. 1971;22: 175-183.

    7. Crawford TB, Henson JB. Immunofluorescent light microscopic and immunologic studies of equine viral arteritis. Proc. 3rd Int. Conf. equine Infec. Dis. Paris 1972, Karger, Basel, 1973, pp: 282-302.

    8. De Boer GF, Osterhaus A , Wemmenhove R. Prevalence of antibodies to equine viruses in the Netherland. 4th Int. Conf. Equine Infec. Dis., Lyon 1978. Vet. Publications Inc., pp: 487-492.

    9. Del Piero F. Equine viral arteritis. Vet Pathol. 2000;37:287–96.

    10. Doll ER, Bryans JT, McCollum WH, Crowe M. Isolation of a filterable agent causing arteritis of horses and abortion by mares. Its differentiation from the equine abortion (influenza) virus. Cornell Vet. 1957;47: 3-41.

    11. Doll ER, Bryans JT, Wilson JC, McCollum WH. Immunization against equine viral arteritis using modified live virus propagated in cell culture of rabbit kidney. Cornell Vet. 1968;58: 497-524.

    12. Estes PC, Cheville NF. The ultrastructure of vascular lesions in equine viral arteritis. Am J Pathol. 1970;58: 235-253.

    13. Fukunaga Y, McCollum WH. Complement fixation reactions in equine viral arteritis. Am J Vet Res. 1977;38:2043-2046.

    14. Fukunaga Y, Matsumura T, Sugiura T, Wada R, Imagawa H, Kanemaru T, Kamada, M. Use of serum neutralization test for equine viral arteritis with different viral strains. Vet Rec. 1994;134:574-576.

    15. Fukunaga Y, Wada R, Kanemaru T, Imagawa H, Kamada M, Samejima T. Immune potency of lyophilized killed vaccine for EVA and its protection against abortion in pregnant mares. J Equi Vet Sci. 1996;16:217-221.

    16. Gerber H, Steck F, Hofer B, Walter L, Freidli U. Serological investigations on equine viral arteritis. 4th Int. Conf. Equine Infec. Dis. Lyon 1978. Veterinary Publications Inc. pp: 461-465.

    17. Golnik W, Michalska Z, Michalak T. Natural equine viral arteritis in foals. Schweiz Arch Tierheilk. 1981;123:523-533.

    18. Harry TO, McCollum WH. Stability of viability and immunizing potency of lyophilized, modified equine arteritis live virus vaccine. Am J Vet Res. 1981;42: 1501-1505.

    19. Herbert K. Controlling arteritis. The Blood-Horse June, 1984;9th pp: 3972-3973.

    20. Holyoak GR, Balasuriya UBR, Broaddus CC, Timoney PJ. Equine viral arteritis: Current status and prevention. Theriogenology 2008;70:403–414.

    21. Horzinek MC. Nonarthropod borne togavirus. 1981;Academic Press. New York.

    22. Hyllseth B. Buoyant density studies on equine arteritis virus. Arch Ges Virus Forch. 1970;30: 97-104.

    23. Inoue T, Yanagawa R, Shinagawa M. Immunofluorescent studies on the multiplication of equine arteritis virus in Vero and E. Derm. (NBL6) cells. Jap J Vet Sci. 1975;37:569-575.

    24. Jaksch Von W, Sibalin M, Taussig E, Pichler L, Burki F. Naturliche Falle und experimentelle Ubertragungen Equiner Virus- Arteritis in Osterreich. Dt Tierarztl Wschr. 1973;80:374-380.

    25. Jones TC, Doll ER, Bryans JT. The lesions of equine viral arteritis. Corn Vet. 1957;47:52-68.

    26. Little TV, Holyoak GR, Timoney PJ, McCollum WH. Output of equine arteritis virus from persistently infected carrier stallions is testosterone dependent. Proc. 6th Int. Conf. Equine Infec. Dis. Cambridge, 1992, pp: 225-229.

    27. Little TV, Deregt D, McCollum WH, Timoney PJ. Evaluation of an immunocytochemical method for the rapid detection of EAV in natural cases of infection. In: Equine Infect. Dis. VII Proc. 7th Int. Conf., Tokyo, 8-11th June, 1994.

    28. Lopez JW, Piero F Del, Glaser A, Finazzi M. Immunoperoxidase fixed histochemistry as a diagnostic tool for detection of EAV in formalin fixed tissues. Equi Vet J. 1996;28:77-79.

    29. Magdalena L, Jerzy R. Molecular epizootiology of equine arteritis virus isolates from Poland. Vet Microbiol. 2008;127:392–398.

    30. Matumoto M, Shimizu T, Ishizaki R. Constant anticorps contre le virus de’arterite equine dans le serum de juments indiennes. C R Soc Biol. 1965;159:1262-1264.

    31. McCollum WH. Development of a modified virus strain and vaccine for equine viral arteritis. J Am Vet Med Assoc. 1970a;155:318-322.

    32. McCollum WH. Vaccination for equine viral arteritis. Proc. 2nd Int. Conf. Infect. Dis. Paris, 1970b; Karger, Basel, pp: 143-151.

    33. McCollum WH. Pathological features of horses given avirulent virus intramuscularly. Am J Vet Res. 1981;42:1218-1220.

    34. McCollum WH, Doll ER, Wilson JC, Johnson CB. Propagation of equine arteritis virus in monolayer cultures of equine kidney. Am J Med Res. 1961;22:731-735.

    35. McCollum WH, Doll ER, Wilson JC, Cheatham J. Isolation and propagation of equine arteritis virus in monolayer cell culture of rabbit kidney. Corn Vet. 1962;52:452-458.

    36. McCollum WH, Prickett ME, Bryans JT. Temporal distribution of EAV in respiratory mucosa, tissues and body fluids of horses infected by inhalation. Res Vet Sci. 1971;2:459-464.

    37. McCollum WH, Swerczek TW. Studies on an epizootic of equine viral arteritis in racehorses. Equi Vet J. 1978;2: 293-299.

    38. McCollum WH, Timoney PJ. The pathogenic qualities of the 1984 strain of equine arteritis virus. Proc. Grayson Found Int. Conf. of Throughbred Breeders Organization of Equine Viral Arteritis, Ireland, Grayson Found Inc. Lexington, Kentucky, 1984: 34-47.

    39. McCollum WH, Little TV, Timoney PJ, Swerczek TW. Resistance of castrated male horses to attempted establishment of the carrier state with EAV. J Comp Path. 1994;111:383-388.

    40. McCollum WH, Timoney PJ, Tengelsen LA. Clinical, virological and serological responses of donkeys to I/N inoculation with KY-84 strain of EAV. J Comp Path. 1995;112:207-211.

    41. Murphy FA. Togavirus morphology and morphogenesis. The Togaviruses: Biology structure and Replication. Ed R.W. Schlesinger. Academic Press, New York. 1995; Pp: 241-316.

    42. Paweska JT, Volkman DH, Barnard BJH, Chirnside ED. Sexual and incontact transmission of Asinine strain of EAV among donkeys. J Clin Microbiol. 1995;33:3296-3299.

    43. Prickett ME, McCollum WH, Bryans JT. The gross and microscopic pathology observed in horses experimentally infected with equine arteritis virus. Proc. 3rd Int. Conf. equine Infect. Dis. Paris, 1973. Pp: 265-272.

    44. Posthuma CC, Pedersen KW, Zhengchun L, Joosten RG, Norbert R, Jessika C, Eric JS. Formation of the Arterivirus Replication/Transcription Complex: a key role for nonstructural protein 3 in the Remodeling of Intracellular Membranes. J Virol. 2008;82(9):4480–4491.

    45. Radwin AI, Burger D. The complement requiring neutralization of equine arteritis virus by late antisera. Virology 1973;51:71-77.

    46. Rusvai M, Hornyyak A, Abonyi T, Fehervari T, Medveczky I, Miklos R, Akos H, Tamas A, Tamas F, Istvan M. Newer observations in the epizootiology of the equine viral arteritis. Maga Allatorv Lapja. 1995;50:16-19.

    47. Senne DA, Pearson JE, Carbrey EA. Equine viral arteritis: a standard procedure for the virus neutralization test and comparison of results of a proficiency test performed at five laboratories. Proc. U. S. Animal Health Association Meeting, Louisville, KY, 1985; pp: 29-33.

    48. Shirai J, Kanno T, Tsuchiya Y, Mitsubayashi S, Seki R. Effects of chlorine, iodine, and quaternary ammonium compound disinfectants on several exotic disease viruses. J Vet Med Sci. 2000;62:85–92.

    49. Timoney P. Equine viral arteritis in Kentucky. Proc. Int. Equine Viral Arteritis. Sem. Ireland. 1984; Grayson Foundation.

    50. Timoney PJ, McCollum WH. The epidemiology of equine viral arteritis. Proc Am Assoc Equi Prac. 1985;30:545-551.

    51. Timoney PJ, McCollum WH, Roberts AW. Detection of the carrier state in stallion persistently infected with equine arteritis virus. In: Proc. 32nd Annual Meeting American Equine Practioners, Nuxville, 1986; pp: 57-65.

    52. Timoney PJ, McCollum WH, Murphy TW, Roberts AW, Willard JG, Carswell GD. The carrier state in equine arteritis virus infection in the stallion with specific emphasis on the venereal mode of virus transmission. J Repro Fertil. 1987a;35:95-102.

    53. Timoney PJ, McCollum WH, Roberts AW. Detection of the carrier state in stallions persistently infected with equine arteritis virus. Proc Am Assoc Equi Prac. 1987b;32:57-65.

    54. Van Der Zeijst BAM, Horzinek MC, Moening V. The genome of equine arteritis virus. Virology 1975;68:418-425.

    55. Vries AAF. The molecular biology of EAV. Procfschrift Faculteit Diergeneeskunde, Universiteit Utrecht, Netherlands. 1994; Pp: 171.

    56. Wada R, Kondo T, Fukunaga Y, Kanemaru T. Histopathological and immunofluorescent studies on the uterus of aborted mares experimentally infected with EAV. J Vet Med. 1994;5: 41-43.

    57. Wada R, Fukunaga Y, Kondo T, Kanemaru T. Ultrastructure and immunochemistry of BHK21 cells infected with a modified Bucyrus strain of EAV. Arch Virol. 1995;40:1173-1180.

    58. Wada R, Fukunaga Y, Kanemaru T, Kondo T. Histopathological and immunofluorescent studies on transplacental infection in experimentally induced abortion by EAV. J Vet Med. 1996;43:65-74.

    59. Wescott D, Lucas M, Paton D. EAV: antigenic analysis of strain variation. In: Immunobiology of viral infections. Proc. 3rd Cong. Eur. Soc. Vet. Virol. Interlaken, Switzerland.1995.

    60. Yun YG, Wong SJ, Branscum AJ, Timoney PJ, Udeni BRB. Development of a fluorescent-microsphere immunoassay for detection of antibodies specific to equine arteritis virus and comparison with the virus neutralization test. Clin Vac Immunol. 2008;15(1):76–87.

    61. Zeggers JJW, Van Der Zeijst BAM, Horzinek MC. The structural proteins of equine arteritis virus. Virology 1976;73:200-205.


Copyright © Vet Scan 2005-

All Right Reserved with VetScan
www.vetscan.co.in and www.kashvet.org
ISSN 0973-6980


Home | e-Learning |Resources | Alumni | Forum | Picture blog | Disclaimer




powered by eMedia Services