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SUMMARY
Equine viral arteritis (EVA) is a contagious viral disease of equids caused by equine arteritis virus (EAV), an RNA virus classified in the family Arteriviridae. The virus is found in horse populations in many countries world-wide. Although infrequently reported in the past, confirmed outbreaks of EVA appear to be on the increase.
The majority of naturally acquired infections with EAV are subclinical. Where present, clinical signs of EVA can vary in range and severity. The disease is characterised principally by fever, depression, anorexia, dependent oedema, especially of the limbs, scrotum and prepuce in the stallion, conjunctivitis, an urticarial-type skin reaction, abortion and, rarely, a fulminating pneumonia or pneumo-enteritis in young foals. Apart from mortality in young foals, the case-fatality rate in outbreaks of EVA is very low. Affected horses almost invariably make complete clinical recoveries. A long-term carrier state can occur in a high percentage of infected stallions, but not in mares or nonbreeding horses.
Identification of the agent: EVA cannot be differentiated clinically from a number of other respiratory and systemic equine diseases. Diagnosis of EAV infection is based on virus isolation, detection of viral antigen or nucleic acid, or demonstration of a specific antibody response. Virus isolation should be attempted from appropriate clinical or post-mortem specimens in rabbit, equine, or monkey kidney cell culture. The identity of isolates of EAV should be confirmed by neutralisation test, reverse-transcription polymerase chain reaction (RT-PCR) assay, or by immunocytochemical methods, namely indirect immunofluorescence or avidin-biotin-peroxidase techniques.
Detection and identification of EAV in suspect cases of the disease can also be attempted using the RT-PCR assay and appropriate viral RNA primers.
Where mortality is associated with a suspected outbreak of EVA, a wide range of tissues should be examined for histological evidence of panvasculitis that is especially pronounced in the small arteries throughout the body. The characteristic vascular lesions present in the mature animal are not a notable feature in EVA-related abortions.
Serological tests: A variety of serological tests, including virus neutralisation (VN), complement fixation (CF), indirect fluorescent antibody, agar gel immunodiffusion and the enzyme-linked immunosorbent assay (ELISA), have been used for the detection of antibody to EAV. The tests currently in widest use are the complement-enhanced VN test and the ELISA. The VN test is a very sensitive and highly specific assay of proven value in diagnosing acute infection and in seroprevalence studies. Several ELISAs have been developed, none of which have been as extensively validated as the VN test though some appear to offer comparable specificity and sensitivity. The CF test is less sensitive than either procedure, but it can be used for diagnosing recent infection.
Requirements for vaccines and diagnostic biologicals: Two commercial tissue culture vaccines are currently available against EVA. One is a modified live virus (MLV) vaccine prepared from virus that has been attenuated for horses by multiple serial transfers in primary equine and rabbit cell cultures. It has been shown to be safe and protective for stallions and nonpregnant mares. Vaccination of foals under 6 weeks of age and of pregnant mares in the final 2 months of gestation is contraindicated. There is no evidence of back reversion to virulence of the vaccine virus following its use in the field over a significant number of years. The second vaccine is an inactivated, adjuvanted product prepared from virus grown in equine cell culture that can be used in nonbreeding and breeding horses. In the absence of appropriate safety data, the vaccine is not currently recommended for use in pregnant mares.
A. INTRODUCTION
Equine viral arteritis (EVA) is a contagious viral disease of equids caused by equine arteritis virus (EAV), a positive-sense, single-stranded RNA virus, and the prototype member of the genus Arterivirus, family Arteriviridae, order Nidovirales (6). Epizootic lymphangitis pinkeye, fièvre typhoide and rotlaufseuche are some of the descriptive terms used in the past to refer to a disease that clinically resembled EVA. The natural host range of EAV would appear to be restricted to equids and the virus does not present a human health hazard (40). EAV is present in the horse population of many countries world-wide (40). There has been an increase in the incidence of EVA in recent years associated with the greater frequency of movement of horses and use of transported semen (2, 40).
While the majority of cases of acute infection with EAV are subclinical, certain strains of the virus can cause disease of varying severity (40). Typical cases of EVA can present with any combination of the following clinical signs: fever, depression, anorexia, leukopenia, dependent oedema, especially of the limbs, scrotum and prepuce of the stallion, conjunctivitis, ocular discharge, supra or periorbital oedema, rhinitis, nasal discharge, a local or generalised urticarial skin reaction, abortion and, rarely, a fulminating pneumonia or pneumo-enteritis in young foals. Regardless of the severity of clinical signs, affected horses almost invariably make complete recoveries. The case-fatality rate in outbreaks of EVA is very low; mortality is usually only seen in very young foals, especially those congenitally infected with the virus (29, 40).
EVA cannot be differentiated clinically from a number of other respiratory and systemic equine diseases, the most common of which are equine influenza, equine herpesvirus 1 and 4 infections, infection with equine rhinitis A and B viruses, equine adenoviruses and streptococcal infections, with particular reference to purpura haemorrhagica. The disease also has clinical similarities to equine infectious anaemia, cases of infection with Getah virus, African horse sickness fever, and toxicosis caused by hoary alyssum (Berteroa incana). EAV replicates and is shed in large quantities from the respiratory tract of acutely infected animals (39). A variable percentage of acutely infected stallions become long-term carriers in the reproductive tract and constant semen shedders of the virus (40, 41). The carrier state has only been found in the stallion, not in the mare, gelding or sexually immature colt (40).
B. DIAGNOSTIC TECHNIQUES
| 1. | Identification of the agent
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| | a) | Virus isolation
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| | | Where an outbreak of EVA is suspected, or when attempting to confirm a case of subclinical EAV infection, virus isolation should be attempted from nasopharyngeal and conjunctival swabs, unclotted blood samples, and semen from stallions considered to be possible carriers of the virus (40). To optimise the chances of virus isolation, the relevant specimens should be obtained as soon as possible after the onset of fever in affected horses. There is evidence that Hheparin can inhibit the growth of EAV in RK-13 cells (1), and therefore, its use as an is contraindicated as an anticoagulant may interfere with isolation of the virus from whole bloodwhen attempting virus isolation from blood, as there is evidence that it inhibits the growth of EAV in RK-13 cells (1). Where EVA is suspected in cases of mortality in young foals or older animals, isolation of EAV can be attempted from a variety of tissues, especially the lymphatic glands associated with the alimentary tract and related organs, and also the lungs, liver and spleen (31). In outbreaks of EVA-related abortion, placental and fetal fluids and a wide range of placental, lymphoreticular and other fetal tissues can be productive sources of virus (40).
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| | | Swabs for attempted isolation should be immersed in a suitable viral transport medium and these, together with any fluids or tissues collected for either virus isolation or reverse-transcription polymerase chain reaction (RT-PCR) testing should be shipped either refrigerated or frozen in an insulated container to the laboratory, preferably using an overnight delivery service. Unclotted blood samples must be transported refrigerated but not frozen.
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| | | Although reportedly not always successful in natural cases of EAV infection (32, 40), virus isolation should be attempted from clinical specimens or necropsied tissues using rabbit, equine or monkey kidney cell culture (32, 40). Selected cell lines, e.g. RK-13 (ATCC CCL-37), LLC-MK2 (ATCC CCL-7), and African green monkey kidney (Vero) (ATCC CCL-81) cells or primary horse or rabbit kidney cell culture can be used, with early passage RK-13 cells being the cell system of choice. Several factors have been shown to influence primary isolation of EAV from semen in RK-13 cells (37). Higher isolation rates have been obtained using 3-5-day-old monolayers, a large inoculum size in relation to the cell surface area in the inoculated flasks or multiwell plates, and most importantly, the incorporation of carboxymethyl cellulose in the overlay medium. It should be noted that most RK-13 cells, including ATCC CCL-37, are contaminated with bovine viral diarrhoea virus, the presence of which appears to enhance sensitivity of this cell system for the primary isolation of EAV. There is some evidence to suggest that primary isolation rates of EAV, particularly from semen, may be increased in RK-13 cells of high passage history.
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| | | Inoculated cultures are examined daily for the appearance of viral cytopathic effect (CPE), which is usually evident within 2-6 days. In the absence of visible CPE, culture supernatants should be subinoculated on to fresh cell monolayers after 5-7 days. The majority of isolations of EAV are made in the first or second passage in cell culture (40, 41). The identity of isolates of EAV can be confirmed in a one-way neutralisation test, by RT-PCR assay (2) or by an immunocytochemical method, namely indirect immunofluorescence (11) or the avidin-biotin-peroxidase (ABC) technique (28). A polyclonal rabbit antiserum has been used to identify EAV in infected cell cultures. Mouse monoclonal antibodies (MAbs) to the nucleocapsid (N) protein and envelope GL protein of EAV (12) and a monospecific polyclonal rabbit antiserum to the envelope (M) protein have also been developed and these can detect various strains of the virus in RK-13 cells as early as 12-24 hours after infection (2, 28).
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Virus isolation from semen (a prescribed test for international trade)
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There is considerable evidence that short- and long-term carrier stallions shed EAV constantly in the semen, but not in respiratory secretions or urine; nor has it been demonstrated in the buffy coat of the blood of such animals (40, 41). Stallions should first be blood tested using the virus neutralisation (VN) test or an appropriately validated enzyme-linked immunosorbent assay (ELISA). Virus isolation should be attempted from the semen of stallions serologically positive for antibodies to EAV that do not have a certified history of vaccination against EVA. Virus isolation is also indicated in the case of shipped semen where a blood sample from the donor stallion is not available. It is recommended that virus isolation from semen be attempted from two samples, which can be collected on the same day, on consecutive days or after an interval of several days or weeks. There is no evidence that the outcome of attempted virus isolation from particular stallions is influenced by the interval between collections or time of the year. Isolation of EAV should be carried out preferably on a portion of an entire ejaculate collected using an artificial vagina or a condom and a teaser or phantom mare. When it is not possible to obtain semen by this means, a less preferable alternative is to collect a dismount sample at the time of breeding. Care should be taken to ensure that no antiseptics/disinfectants are used in the cleansing of the external genitalia of the stallion prior to collection. Samples should contain the sperm-rich fraction of the ejaculate with which EAV is associated. The virus is not present in the pre-sperm fraction of semen (40, 41). Immediately following collection, the semen should be refrigerated on crushed ice or on freezer packs for transport to the laboratory with a minimum of delay. Where there is likely to be a delay in submitting a specimen for testing, the semen can be frozen at or below -20°C for a varying period of days or weeks before being dispatched to the laboratory. Freezing a sample has not been found to militate against isolation of EAV from the semen of a carrier stallion.
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Test procedure
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Semen samples are pretreated before inoculation into cell culture by short-term sonication (for three 15-second cycles) followed by centrifugation at 1000 g for 10 minutes at 4°C to sediment the spermatozoa.
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After removal of culture medium, 3-5-day-old confluent monolayer cultures of RK-13 cells, either in 25 cm2 tissue culture flasks or multiwell plates, are inoculated with serial decimal dilutions (10-1-10-3) of seminal plasma in tissue culture maintenance medium containing 2% fetal bovine serum and antibiotics. An inoculum of 1 ml per 25 cm2 flask is used and no fewer than two flasks per dilution of seminal plasma are inoculated. Inoculum size and number of wells inoculated per dilution of a specimen should be pro-rated where multiwell plates are used. Appropriate dilutions of a virus positive control semen sample or virus control of known titre diluted in culture medium should be included in each test.
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The flasks or cover plates are closed and gently rotated to disperse the inoculum over the cell monolayers.
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Inoculated cultures are then incubated for 1 hour at 37°C either in an aerobic incubator or an incubator containing a humidified atmosphere of 5% CO2 in air, depending on whether flasks or multiwell plates are used.
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Without removing any of the inoculum or washing the cell monolayers, the latter are overlaid with 0.75% carboxymethyl cellulose containing medium with antibiotics.
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The flasks or plates are reincubated at 37°C and checked microscopically for viral CPE, which is usually evident within 2-6 days.
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In the absence of visible CPE, culture supernatants are subinoculated on to fresh cell monolayers after 5-7 days. After removal of the overlay medium, monolayers are stained with 0.1% formalin-buffered crystal violet solution.
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The identity of any virus isolates should be confirmed by VN, immunofluorescence or ABC technique, using a monospecific polyvalent antiserum to EAV or MAbs to the N or GL proteins of the virus (11, 28, 40, 41), or by RT-PCR assay and appropriate viral RNA primers (2).
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In the one-way neutralisation test, serial decimal dilutions of the virus isolate are tested against an MAb or monospecific antiserum prepared against the prototype Bucyrus strain of EAV (ATCC VR 796) and also a serum negative for neutralising antibodies to the virus. Corresponding titrations of the prototype Bucyrus virus with the same reference antibody reagents are included as test controls. The test is performed in either 25 cm2 tissue culture flasks or multiwell plates. Appropriate quantities of the known EAV positive and negative antibody reagents are inactivated for 30 minutes in a water bath at 56°C and diluted 1/4 in phosphate buffered saline, pH 7.2; then 0.3 ml of diluted antibody reagent is dispensed into five tubes for each virus isolate to be tested. Serial decimal dilutions (10-1-10-5) of each virus are made in Eagles Minimal Essential Medium containing 10% fetal bovine serum, antibiotics and 10% freshly diluted guinea-pig complement. Then, 0.3 ml of each virus dilution is added to the tubes containing positive and negative antibody reagents. The tubes are shaken and the virus/antibody mixtures are incubated for 1 hour at 37°C. The mixtures are then inoculated on to 3-5-day-old confluent monolayer cultures of RK-13 cells, either in 25 cm2 flasks or multiwell plates, using two flasks or wells per virus dilution. Each flask is inoculated with 0.25 ml of virus/antibody mixture; the inoculum size is pro-rated where multiwell plates are used. Inoculated flasks or plates are incubated for 2 hours at 37°C, gently rocking after 1 hour to disperse the inoculum over the cell monolayers. Without removing any of the inoculum or washing the cell monolayers, the latter are overlaid with 0.75% carboxymethyl cellulose containing medium and incubated for 4-5 days at 37°C, either in an aerobic incubator or an incubator containing a humidified atmosphere of 5% CO2 in air. After removal of the medium, monolayers are stained with 0.1% formalin-buffered crystal violet solution. Plaques are counted and the virus infectivity titre is determined both in the presence and absence of EAV antibodies using the Spearman-Kärber method. Identification of a virus isolate is based on the extent of plaque reduction compared with that of the prototype Bucyrus strain of EAV.
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Most EAV isolates from carrier stallions are made in the first passage in cell culture using the described test procedure (39, 40, 41). The occurrence of nonviral cytotoxicity or bacterial contamination of specimens are not significant problems when attempting isolation of this virus from stallion semen. Nonviral cytotoxicity, if observed, usually affects monolayers inoculated with the 10-1 and, much less frequently, the 10-2 dilution of seminal plasma. Treatment of seminal plasma with polyethylene glycol (Mol. wt 6000) prior to inoculation has been used with some success in overcoming this problem (19). The method described involves the addition of polyethylene glycol to the 10-1-10-3 dilutions of seminal plasma to give a final concentration of 10% in each dilution. The mixtures are held overnight at 4°C with gentle stirring, after which they are centrifuged at 2000 g for 30 minutes and the supernatants are discarded. The precipitates are suspended in cell culture maintenance medium to one-tenth the volume of the original dilutions and the mixtures are homogenised. They are then centrifuged at 2000 g for 30 minutes and the supernatants are taken off and used for inoculation. There is no evidence to indicate that pretreatment of seminal plasma in this manner reduces sensitivity of the virus isolation procedure.
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The presence of anti-EAV antibody in the seminal plasma of certain virus-shedding stallions has not been found to prevent detection of the carrier state in these animals.
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| | b) | Nucleic acid recognition methods
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| | | The RT-PCR assay is an additional method (to virus isolation in cell culture) for EAV detection. Single, nested, and one-tube real-time TaqMan® RT-PCR assays have been developed and evaluated for detection of various strains of the virus in cell culture, semen and nasal secretions (3, 5, 21). Single-step extraction of RNA followed by reverse transcription and amplification has been described (5, 21). Detection of amplified products can be performed either by agarose gel electrophoresis or by an ELISA-PCR assay using commercial reagents. The RT-PCR assay provides a means of identifying virus-specific RNA in clinical specimens, namely nasopharyngeal swab filtrates, buffy coats, semen and urine and in post-mortem tissue samples (2, 3, 5, 21). Comparable results to virus isolation have been obtained with both a nested RT-PCR assay that takes only 2 days to complete and a real-time TaqMan® assay (2, 3, 5, 21). The assay has the advantage of not requiring viable virus for performance of the test. Because of the high sensitivity of the procedure, however, there is the potential for cross-contamination between samples in the laboratory, giving rise to a false-positive result. The risk of cross-contamination is greater in conventional RT-PCR because of the separate reverse transcription step and the second PCR step in nested PCR assays (3). To minimise the risk of this occurring, considerable care needs to be taken, especially during the steps of RNA extraction and amplification, and relevant EAV positive and negative controls and, where appropriate, RNA extracted from the tissue culture fluid of uninfected cells, need to be included in each PCR assay.
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| | | Primer selection is critical to the sensitivity of the RT-PCR assay with primers preferably designed from the most conserved regions of the viral genome. Sequences chosen from the viral polymerase, nucleocapsid (N) or envelope (M) protein genes are reportedly effective in a single, nested or real-time TaqMan® RT-PCR assay (3, 5, 21). There is growing evidence that open reading frame (ORF) 7 is the most conserved gene among different strains of EAV (4) and ORF-7-specific primers have detected a diversity of strains of the virus of European and North American origin (3). However, until such time as a complete consensus or universal primer set for EAV has been agreed upon, where feasible, it is advisable to use the RT-PCR assay in conjunction with, and not as an alternative to, virus isolation for the identification of virus in clinical or post-mortem specimens. When virus isolation is attempted in full accordance with OIE recommended procedures, it has been found to be comparable to the RT-PCR assay for the detection of EAV (2, 3, 5, 21).
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| | | It should be emphasised that as the RT-PCR assay will not distinguish between infectious and noninfectious or incomplete virus and, consequently, it cannot establish the current infectivity status of a particular animal or sample of semen, a factor of considerable importance with respect to the international trade in horses and equine semen.
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| | | Strains of EAV isolated from different regions of the world have been classified into different phylogenetic groups by sequence analysis of the envelope (GL, GS, M) protein genes and the nucleocapsid (N) protein gene (4, 8, 39). The relationships between strains demonstrated by nucleotide sequencing is a useful molecular epidemiological tool for tracing the origin of outbreaks of EVA (2).
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| | c) | Histopathological and immunohistochemical methods
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| | | Where mortality is associated with a suspected outbreak of EVA, a wide range of tissues should be examined for histological evidence of panvasculitis that is especially pronounced in the small arteries throughout the body, particularly in the caecum, colon, spleen, associated lymphatic glands and adrenal cortex (11, 26). The presence of a disseminated necrotising arteritis involving endothelial and medial cells of affected vessels is considered to be pathognomonic of EVA. The characteristic vascular lesions present in the mature animal are not, however, as prominent a feature in EAV-related abortions (25).
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| | | EAV antigen can be identified in various tissues of EVA-affected animals either in the presence or absence of lesions (11). It can be detected within the cytoplasm of infected cells by immunofluorescence using conjugated equine polyclonal anti-EAV serum (11), or by the ABC technique using mouse MAbs to the N or GL proteins of the virus (29).
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| 2. | Serological tests
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| | A variety of serological tests including neutralisation (microneutralisation [38] and plaque reduction [30]), the complement fixation (CF) test (16), the indirect fluorescent antibody test (11), the agar gel immunodiffusion (11), and the ELISA (7, 9, 23, 24, 27, 36) have been used to detect antibody to EAV.
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| | The test currently in widest use to diagnose infection, carry out seroprevalence studies, and test horses for export, is a microneutralisation test in the presence of complement. Apart from the VN test, the CF test has been used for diagnosing recent EAV infection sinceas scomplement-fixing antibodies are relatively short-lived in duration (16). In contrast, neutralising antibody titres to EAV can persist for several years after natural infection (40). Although a number of ELISAs have been developed (7, 9, 27), none has as yet been as extensively validated as the VN, though some appear to offer comparable sensitivity and specificity (7, 23, 24, 36). Unlike the VN test, a positive reaction in the ELISA is not necessarily reflective of the protective immune status of an individual horse to EAV as both non-neutralising and neutralising antibodies are involved.
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| | Antiserum to unpurified EAV has been prepared in horses and in rabbits using conventional immunisation protocols. Also, mouse monoclonal and monospecific rabbit polyclonal antibodies have been developed to the nucleocapsid (N) protein envelope (GL) protein, envelope (M and GS) proteins and other proteins of EAV (12).
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| | OIE Standard Sera for EAV are available (from Dr P.J. Timoney, Director, Maxwell H. Gluck Equine Research Center, Dept of Veterinary Science, University of Kentucky, Lexington, Kentucky 40546-0099, United States of America) and these can facilitate international standardisation of the microneutralisation test and ELISA.
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| | Only one major serotype of EAV has been recognised so far (30, 40). This is represented by the prototype Bucyrus strain (ATCC VR 796), from which the reference virus used in all EAV serological assays has been derived. Virus stock is grown in the RK-13 cell line, clarified of cellular debris by low-speed centrifugation and stored in aliquots at -70°C. Several frozen aliquots are thawed and the infectivity of the stock virus is determined by titration in RK-13 cells.
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Virus neutralisation (a prescribed test for international trade)
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| | | The VN test is used to screen stallions for evidence of EAV infection and to determine whether there is a need to attempt virus detection in semen using cell culture or RT-PCR assay. It can also be used for diagnostic purposes to confirm infection in suspect cases of EVA. The VN test procedure in current widest use is that developed by the National Veterinary Service Laboratories of the United States Department of Agriculture (38). It is recommended that the test be carried out in RK-13 cells using the approved CVL-Bucyrus (Weybridge) strain of EAV as reference virus (available from: Virology Department, Veterinary Laboratories Agency, Weybridge, New Haw, Addlestone KT15 3NB, United Kingdom). Originally derived from the prototype Bucyrus virus, the passage history of the CVL (Weybridge) strain is not fully documented. The sensitivity of the VN test for detection of antibodies to EAV can be significantly influenced by several factors, especially the source and passage history of the strain of virus used (14, 15). The CVL-Bucyrus strain and the highly attenuated vaccine strain of EAV are of comparable sensitivity for detecting low-titred positive sera, especially from EVA-vaccinated horses. For this test, it is important to obtain a sterile blood sample as contamination of serum can interfere with the result. Efforts are continuing to bring about greater uniformity in the testing protocol and comparability in serological results among laboratories providing the VN test.
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| | | . | Test procedure
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| | | i) | Sera are inactivated for 30 minutes in a water bath at 56°C (control sera, only once).
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| | | ii) | Serial twofold dilutions of the inactivated test sera in serum-free cell culture medium (25 µl volumes) are made in a 96-well, flat-bottomed, cell-culture grade microtitre plate starting at a 1/2 serum dilution and using duplicate rows of wells for each serum to be tested. Most sera are screened initially at a 1/4 and 1/8 serum dilution (i.e. final serum dilution after addition of an equal volume of the appropriate dilution of stock virus to each well). Positive samples at the 1/8 dilution can, if desired, be retested and titrated out for end-point determination. Individual serum controls, together with negative and known low- and high-titred positive control sera must also be included in each test.
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| | | iii) | A dilution of stock virus to contain from 100 to 300 TCID50 (50% tissue culture infective dose) per 25 µl is prepared using as diluent, serum-free cell culture medium containing antibiotics and fresh guinea-pig or rabbit complement at a final concentration of 10%.
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| | | iv) | 25 µl of the appropriate dilution of stock virus is added to every well containing 25 µl of each serum dilution, except the test serum control wells.
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| | | v) | A virus back titration of the working dilution of stock virus is included, using four wells per tenfold dilution, to confirm the validity of the test results.
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| | | vi) | The plates are covered and shaken gently to facilitate mixing of the serum/virus mixtures.
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| | | vii) | The plates are incubated for 1 hour at 37°C in a humid atmosphere of 5% CO2 in air.
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A suspension of cells from 3-5-day-old cultures of RK-13 cells are prepared using a concentration that will ensure confluent monolayers in the microtitre plate wells within 18-24 hours after seeding.
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| | | ix) | 100 µl of cell suspension is added to every well, the plates covered with plate lids or sealed with tape and shaken gently.
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| | | x) | The plates are incubated at 37°C in a humid atmosphere of 5% CO2 in air.
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| | | xi) | The plates are read microscopically for non-viral CPE after 12-18 hours and again for viral CPE after 48-72 hours' incubation. The validity of the test is confirmed by establishing that the working dilution of stock virus contained 30-300 TCID50 virus and that the titres of the positive serum controls are within 0.3 log10 units of their predetermined titres.
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| | | A serum dilution is considered to be positive if there is an estimated 75% or perferably preferably 100% reduction in the amount of viral CPE in the serum test wells compared with that present in the wells of the lowest virus control dilution. End-points are then calculated using the Spearman-Kärber method. A titre of 1/4 or greater is considered to be positive. A negative serum should only have a trace (less than 25%) or no virus neutralisation at the lowest dilution tested. Antibody titres may, on occasion, be difficult to define as partial neutralisation may be observed over a range of several serum dilutions. Infrequently, sera will be encountered that cause toxic changes in the lower dilutions tested. In such cases it may not be possible to establish whether the sample is negative or a low-titred positive. The problem may be overcome by testing another serum sample from the animal in question or by retesting the toxic sample using microtitre plates with confluent monolayers of RK-13 cells that had been seeded the previous day. It has been reported that the toxicity of the serum can be reduced or eliminated if adsorbed with packed RK-13 cells. Vaccination status for equine herpesviruses should be considered when evaluating sera causing non-viral cytotoxicity. One of the equine herpesvirus vaccines currently available in Europe can stimulate antibodies to rabbit kidney cells which, in turn, can interfere with interpretation of the test results (20).
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| | b) | Enzyme-linked immunosorbent assay
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| | | A number of direct or indirect ELISAs have been developed for the detection of antibodies to EAV (7, 9, 23, 24, 27, 36). These have been based on the use of purified virus or recombinant-derived viral antigens. The usefulness of earlier assays was compromised by the frequency of false-positive reactions (10). The latter were associated with the presence of antibodies to various tissue culture antigens in the sera of horses that had been vaccinated with tissue-culture-derived antigens (10). Identification of the importance of the viral GL protein in stimulation of the humoral antibody response to EAV led to the development of several ELISAs that employ a portion of, or the entire recombinant protein produced in a bacterial or baculovirus expression system (9, 12, 23). Most recently, an ovalbumin-conjugated synthetic peptide representing amino acids 81-106 of the GL protein has been used (36). Some of these assays appear to offer comparable sensitivity and specificity to the VN test and may detect EAV-specific antibodies prior to a positive reaction being obtainable in the VN test (7). False-negative reactions can occur, however, with some of these assays. Screening a random peptide-phage library with polyclonal sera from EAV-infected horses led to the identification of ligands, which were purified and used as antigen in an ELISA for EAV (24). No correlation was found, however, between absorbency values obtained with this assay and neutralising antibody titres, indicating that the antibodies being detected were largely against nonsurface epitopes of the virus. An ELISA based on the use of a combination of the GL, M or N structural proteins of EAV expressed from recombinant baculoviruses successfully detected viral antibody in naturally or experimentally infected horses but not in EVA-vaccinated animals (23). Of major importance with respect to any GL protein-based ELISA for EAV is the fact that test sensitivity will vary depending on the ectodomain sequence(s) of this viral protein used in the assay. Considerable amino acid sequence variation within this domain has been found between isolates of EAV (4). To maximise sensitivity of a GL-based ELISA, it may be necessary to include multiple ectodomain sequences representative of known phylogenetically different isolates of EAV rather than depend on a single ectodomain sequence. Two more recently described ELISAs appear to offer most promise as reliable serodiagnostic tests for EAV infection (9, 36). A blocking ELISA involving MAb produced against the GL protein was reported to have a sensitivity of 99.4% and a specificity of 97.7% compared with the VN test (9). Another assay, a GL ovalbumin-conjugated synthetic peptide ELISA was shown to have a sensitivity and specificity of 96.75% and 95.6%, respectively, using a panel of 400 VN positive sera and 400 VN negative samples (36). It is expected that an ELISA will be available soon having very similar if not equivalent sensitivity and specificity to the VN test.
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C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS
A number of experimental and commercial vaccines have been developed against EVA. Currently, there are two commercially available vaccines, both tissue-culture derived. The first is a modified live virus (MLV) vaccine prepared from virus that has been attenuated for horses by multiple serial transfers in equine and rabbit cell cultures (13, 30). This vaccine is licensed for use in stallions, nonpregnant mares and in nonbreeding horses. Whereas nonbreeding horses can be vaccinated at any time, stallions and mares should be vaccinated not less than 3 weeks prior to breeding. The vaccine is not recommended for use in pregnant mares, especially in the last 2 months of gestation, nor in foals under 6 weeks of age unless in the face of significant risk of exposure to natural infection. The vaccine is commercially available in the USA and Canada. It has also been used in New Zealand, subject to ministerial controls, to aid in that country's EVA eradication programme.
The second commercially available vaccine against EVA is an inactivated product prepared from virus grown in equine cell culture, which is filtered, chemically inactivated and then combined with a metabolisable adjuvant. This vaccine is licensed for use in nonbreeding and breeding horses. In the absence of appropriate safety data, the vaccine is currently not recommended for use in pregnant mares. The initial vaccination regimen involves two doses of vaccine administered intramuscularly 3-6 weeks apart. Booster vaccination at 6-month intervals is recommended by the manufacturer. The inactivated vaccine is licensed for commercial use in certain European countries, including Denmark, France, Germany, Ireland, Sweden and the United Kingdom.
An additional inactivated vaccine against EVA has been developed in Japan for use should an outbreak of EVA occur in that country (18). It is an aqueous formalin-inactivated vaccine that has been shown to be safe and effective for use in nonbreeding and breeding horses. For optimal immunisation with this vaccine, horses require a primary course of two injections given at an interval of 4 weeks, with a booster dose administered every 6-12 months. As the vaccine is currently not commercially available, no details will be provided on its production.
Guidelines for the production of veterinary vaccines are given in Chapter I.1.7. Principles of veterinary vaccine production. The guidelines given here and in Chapter I.1.7 are intended to be general in nature and may be supplemented by national and regional requirements.
| 1. | Seed management
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| | a) | Characteristics of the seed
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| | | Both MLV and inactivated commercial vaccines are derived from the prototype Bucyrus strain of EAV (ATCC VR 796). Available evidence points to the existence of only one major serotype of the virus, and strain variation is not considered to be of significance in relation to vaccine efficacy (30, 40).
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| | | In the case of the MLV vaccine, the prototype virus was attenuated by serial passage in primary cultures of horse kidney (HK-131), rabbit kidney (RK-111), and a diploid equine dermal cell line, ATCC CCL57 (ECID-24) (13, 22, 30). The indications from the use of this vaccine are that the virus is safe and immunogenic between its 80th and 111th passage in primary rabbit kidney (13, 22, 30, 33, 34, 42).
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| | | The inactivated adjuvanted vaccine is prepared from the unattenuated prototype Bucyrus strain of EAV (ATCC VR 796) that has been plaque purified and in its fourth serial passage in the diploid equine dermal cell line (ECID-4). After growth in cell culture, the virus is then purified by filtration before being chemically inactivated and adjuvanted.
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| | | Suitable lots of master seed virus for each vaccine should be maintained in liquid nitrogen or its equivalent.
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| | b) | Method of culture
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| | | The virus for both MLV and inactivated vaccines should be grown in a stable cell culture system, such as equine dermal cells, using an appropriate medium supplemented with sterile bovine serum or bovine serum albumin as replacement for bovine serum in the growth medium. Cell monolayers should be washed prior to virus inoculation to remove traces of bovine serum. Extensive virus growth as evidenced by the appearance of cytopathic changes in 80-100% of the cells should be obtained within 2-3 days.
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| | c) | Validation as a vaccine
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| | | In the case of both MLV and inactivated vaccines, the respective virus strains should be grown in an appropriate cell culture system that has been officially approved for vaccine production and confirmed to be free from extraneous bacteria, fungi, mycoplasmas and viruses (35). The identity of the vaccine virus in the master seed should be confirmed by neutralisation with homologous anti-EAV serum. Incomplete neutralisation of EAV by homologous horse or rabbit antisera has been scientifically documented (38) and is a problem when screening master seed virus for extraneous viruses and when attempting to confirm the identity of the vaccine virus. The problem has been circumvented by reducing the infectivity titre of the master seed virus below that required for seed virus production before conducting a neutralisation test on the diluted virus. Virus/serum mixtures are tested for residual live virus by serial passage in cell culture. No evidence of cytopathic viruses, haemadsorbing viruses, or noncytopathic strains of bovine virus diarrhoea virus should be found, based on attempted virus isolation in cell culture. If cells of equine origin are used, they should be confirmed to be free from equine infectious anaemia virus. The newer technologies of PCR and antigen-capture ELISA may be used in the future as adjuncts to virus isolation in screening for adventitious agents.
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| | | The MLV vaccine has been shown to be both safe and effective for use in stallions and nonpregnant mares (33, 34, 42). Vaccination confers a high level of protective immunity that persists for at least several years (22, 30, 40). Based on experimental studies and extensive field use of the vaccine since 1985, there is no evidence of back reversion to virulence of the vaccine virus, nor of recombination of the vaccine virus with naturally occurring strains of EAV. Furthermore, there are ample data to confirm that the attenuated strain of EAV in the current vaccine is not shed in the semen of stallions after vaccination nor does it localise and set up the carrier state in the reproductive tract of the stallion (34, 40, 42).
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| | | The commercial inactivated vaccine has been shown to be nonreactive and safe for use in healthy nonbreeding and breeding horses. Transient local reactions may be observed in less than 10% of horses vaccinated with the inactivated vaccine. Limited field studies of this vaccine indicate that it is immunogenic, stimulating a satisfactory degree of immunity, the duration of which has yet to be reported.
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| | | Although there are no published reports on the efficacy of either commercial vaccine in preventing establishment of the carrier state in the stallion, an aqueous formalin inactivated vaccine against EVA has been shown to prevent virus persistence in the reproductive tract of vaccinated stallions following subsequent experimental challenge with EAV (17).
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| 2. | Method of manufacture
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| | Both the MLV and inactivated vaccines are produced by cultivation of the respective seed viruses in an equine dermal cell system. Cell monolayers should be washed prior to inoculation with seed virus to remove traces of bovine serum in the growth medium. Inoculated cultures should be maintained on an appropriate maintenance medium. Harvesting of infected cultures should take place when almost the entire cell sheet shows the characteristic CPE. Supernatant fluid and cells are harvested and clarified of cellular debris and unwanted material by filtration. In the case of the inactivated vaccine, the purified virus is then chemically inactivated and adjuvanted with a metabolisable adjuvant.
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| 3. | In-process control
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| | The MLV and inactivated vaccines should be produced in a stable cell line that has been tested for identity and confirmed to be free from contamination by bacteria, fungi, mycoplasmas or other adventitious agents. In addition to the preproduction testing of the master seed virus for each vaccine and the cell line for adventitious contaminants, the cell cultures infected with the respective vaccine viruses should be examined macroscopically for evidence of microbial growth or other extraneous contamination during the incubation period. If growth in a culture vessel cannot be reliably determined by visual examination, subculture, microscopic examination, or both should be carried out.
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| 4. | Batch control
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| | a) | Sterility
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| | | Tests for sterility and freedom from contamination of biological materials may be found in Chapter I.1.5.
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| | b) | Safety
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| | | In the case of both MLV and inactivated vaccines, each production lot of vaccine should be checked for extraneous bacterial, fungal and mycoplasmal contaminants. The vaccine should be safety tested by the intramuscular inoculation of at least two horses seronegative for neutralising antibodies to EAV with one vaccine dose of lyophilised virus each (35). None of the inoculated horses should develop any clinical signs of disease other than mild pyrexia during the ensuing 2-week observation period. In addition, nasopharyngeal swabs should be collected daily from each horse for attempted virus isolation; white blood cell counts and body temperatures should also be determined on a daily basis. No significant febrile or haematological changes should supervene following vaccination (34, 40, 42). Limited shedding of vaccine virus by the respiratory route for at most 7 days may be demonstrated in the occasional vaccinated horse (42). There is no evidence of virus shedding in the semen of stallions following vaccination (34, 42).
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| | | To ensure complete inactivation of the vaccine virus, each serial lot of the inactivated vaccine should be checked for viable virus by three serial passages in equine dermal cells and by direct fluorescent antibody staining with specific EAV conjugate before being combined with adjuvant. This should be followed by a safety test in guinea-pigs and mice.
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| | c) | Potency
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| | | Potency of the vaccine in the final containers is determined by plaque infectivity assay in monolayer cultures of equine dermal cells and by a vaccination challenge test in horses (35). The vaccine must be tested in triplicate in cell culture, the mean infectivity titre calculated and the dose rate determined on the basis that each dose of vaccine shall contain not less than 3 x 104 plaque-forming units of attenuated EAV. The in-vivo potency of the MLV and inactivated vaccines is evaluated in a single vaccination challenge test using 17-20 vaccinated and 5-7 control horses or in two tests each comprising ten vaccinates and five controls.
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| | | The viral antigen concentration in the inactivated vaccine is over one-thousand times the concentration of viral antigen present in the MLV vaccine.
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| | d) | Duration of immunity
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| | | Detectable neutralising antibody titres to EAV should develop in the majority of horses within 3 weeks of vaccination with the MLV vaccine (33, 34, 40, 42). Responses to primary vaccination are characterised by a rapid fall in antibody titres with a significant number of animals reverting to seronegativity 1-3 months after vaccination (42). Revaccination with this vaccine results in an excellent anamnestic response, however, with the development of high antibody titres that remain relatively undiminished for 12 months or longer (40).
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| | | Experimental studies have shown that most horses vaccinated with the inactivated vaccine develop low to moderate neutralising antibody titres to EAV by day 14 after the second vaccination. There is no published information on the duration of immunity conferred by this vaccine.
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| | e) | Stability
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| | | The lyophilised MLV vaccine can be stored for at least 3-4 years at 2-7°C without loss in infectivity, provided it is kept in the dark (22). Infectivity is preserved for much longer periods if vaccine is frozen at -20°C or below. Once rehydrated, however, the vaccine should be used within 1 hour or else destroyed. The inactivated vaccine is stored as a liquid suspension at 2-8°C, with no loss of potency for at least 1 year, provided it is protected from light.
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| | f) | Preservatives
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| | | The preservatives added to the MLV and inactivated vaccines are neomycin, polymyxin B and amphotericin B.
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| | g) | Precautions (hazards)
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| | | Pregnant mares should not be vaccinated with the MLV vaccine during the last 2 months of gestation, as there is a risk, albeit minimal, of fetal invasion by the vaccine virus. The possibility of a vaccinally induced anaphylactic reaction, though very rare, could result from the administration of either the MLV or inactivated vaccine. In the absence of appropriate safety data, the inactivated vaccine is currently not recommended for use in pregnant mares.
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| 5. | Tests on the final product
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| | a) | Safety
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| | | With the exception of the inactivated vaccine, which needs to be sterility tested a second time to ensure freedom from contamination, no further safety tests are required on the inactivated or MLV vaccines.
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| | b) | Potency
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| | | No potency tests additional to those conducted on each production lot of the MLV or inactivated vaccines are required on either final product.
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| 2. | Balasuriya U.B.R., Evermann J.F., Hedges J.F., McKeirnan A.J., Mitten J.Q., Beyer J.C., McCollum W.H., Timoney P.J. & MacLachlan N.J. (1998). Serologic and molecular characterization of an abortigenic strain of equine arteritis virus isolated from infective frozen semen and an aborted equine fetus. J. Am. Vet. Med. Assoc., 213, 1586-1589.
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| 4. | Balasuriya U.B.R., Timoney P.J., McCollum W.H. & MacLachlan N.J. (1995). Phylogenetic analysis of open reading frame 5 of field isolates of equine arteritis virus and identification of conserved and nonconserved regions in the GL envelope glycoprotein. Virology, 214, 690-697.
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| 5. | Belak S., Ballagi-Pordany A., Timoney P., McCollum W.H, Little T.V., Hyllseth B. & Klingeborn B. (1995). Evaluation of a nested PCR assay for the detection of equine arteritis virus infection. Proceedings of the 7th International Conference on Equine Infectious Diseases, Tokyo, Japan, 1994, 33-38.
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| 16. | Fukunaga Y. & McCollum W.H. (1977). Complement fixation reactions in equine viral arteritis. Am. J. Vet. Res., 38, 2043-2046.
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| 18. | Fukunaga Y., Wada R., Matsumura T., Sugiura T. & Imagawa H. (1990). Induction of immune response and protection from equine viral arteritis (EVA) by formalin inactivated-virus vaccine for EVA in horses. J. Vet. Med. (B), 37, 135-141.
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| 19. | Fukunaga Y., Wada R., Sugita S., Fujita Y., Nambo Y., Imagawa H., Kanemaru T., Kamada M., Komatsu N. & Akashi H. (2000). In vitro detection of equine arteritis virus from seminal plasma for identification of carrier stallions. J. Vet. Med. Sci., 62, 643-646.
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| 20. | Gerahty R.J., Newton J.R., Castillo-Olivares J., Cardwell J.M. & Mumford J.A. (2003). Testing for equine arteritis virus. Vet. Rec., 152, 478.
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| 21. | Gilbert S.A., Timoney P.J., McCollum W.H. & Deregt D. (1997). Detection of equine arteritis virus in the semen of carrier stallions using a sensitive nested PCR assay. J. Clin. Microbiol., 35, 2181-2183.
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| 22. | Harry T.O. & McCollum W.H. (1981). Stability of viability and immunising potency of lyophilised, modified equine arteritis live-virus vaccine. Am. J. Vet. Res., 42, 1501-1505.
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| 23. | Hedges J.F., Balasuriya U.B.R., Shabbir A., Timoney P.J., McCollum W.H., Yilma T. & MacLachlan N.J. (1998). Detection of antibodies to equine arteritis virus by enzyme linked immunosorbant assays utilizing GL, M and N proteins expressed from recombinant baculoviruses. J. Virol. Methods, 76,127-137.
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| 25. | Johnson B., Baldwin C., Timoney P. & Ely R. (1991). Arteritis in equine fetuses aborted due to equine viral arteritis. Vet. Pathol., 28, 248-250.
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| 26. | Jones T.C., Doll E.R. & Bryans J.T. (1957). The lesions of equine viral arteritis. Cornell Vet., 47, 52-68.
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| 28. | Little T.V., Deregt D., McCollum W.H., & Timoney P.J. (1995). Evaluation of an immunocytochemical method for rapid detection and identification of equine arteritis virus in natural cases of infection. Proceedings of the Seventh International Conference on Equine Infectious Diseases, Tokyo, Japan, 1994, 27-31.
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| 31. | McCollum W.H., Prickett M.E. & Bryans J.T. (1971). Temporal distribution of equine arteritis virus in respiratory mucosa, tissues and body fluids of horses infected by inhalation. Res. Vet. Sci., 2, 459-464.
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| 32. | McCollum W.H. & Swerczek T.W. (1978). Studies of an epizootic of equine viral arteritis in racehorses. J. Equine Med. Surg., 2, 293-299.
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| 33. | McCollum W.H. Timoney P.J., Roberts A.W., Willard J.E. & Carswell G.D. (1988). Response of vaccinated and non-vaccinated mares to artificial insemination with semen from stallions persistently infected with equine arteritis virus. Proceedings of the Fifth International Conference on Equine Infectious Diseases, Lexington, 1987, University Press of Kentucky, Lexington, Kentucky, USA, 13-18.
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| 33. | McKinnon A.O., Colbern G.C., Collins J.K., Bowen R.A., Voss J.L. & Umphenour N.W. (1986). Vaccination of stallions with modified live equine viral arteritis virus. J. Equine Vet. Sci., 6, 66-69.
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| 34. | Moore B.O. (1986). Development and evaluation of three equine vaccines. Irish Vet. J., 40, 105-107.
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| 35. | Nugent J., Sinclair R., deVries A.A.F., Eberhardt R.Y., Castillo-Olivares J., Davis Poynter N., Rottier P.J.M. & Mumford J.A. (2000). Development and evaluation of ELISA procedures to detect antibodies against the major envelope protein (GL) of equine arteritis virus. J. Virol. Methods, 90, 167-183.
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| 37. | Ostlund E.N., Peters J.C, Stoker A.M, McCollum, W.H & Timoney P.J. (1997). Enhancement of cell culture growth of two arteriviruses by carboxymethyl cellulose overlay. Abstract. Proceedings of the Annual Meeting of the American Association of Veterinary Laboratory Diagnosticians, Louisville, Kentucky, USA, p. 33.
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| 38. | Senne D.A., Pearson J.E. & Cabrey E.A. (1985). Equine viral arteritis: A standard procedure for the virus neutralisation test and comparison of results of a proficiency test performed at five laboratories. Proc. U.S. Anim. Health Assoc., 89, 29-34.
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| 39. | Stadejek T., Bjorklund H., Bascunana C.R., Ciabatti I.M., Scicluna M.T., Amaddeo D., McCollum W.H., Autorino G.L., Timoney P.J., Paton D.J., Klingeborn B. & Belak S. (1999). Genetic diversity of equine arteritis virus. J. Gen. Virol., 80, 691-699.
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| 40. | Timoney P.J. & McCollum W.H. (1993). Equine viral arteritis. Vet. Clin. North Am. Equine Pract., 9, 295-309.
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| 41. | Timoney P.J. & McCollum W.H. (2000). Equine viral arteritis: Further characterization of the carrier state in the stallion. J. Reprod. Fertil. (Suppl.), 56, 3-11.
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| 42. | Timoney P.J., Umphenour N.W. & McCollum W.H. (1988). Safety evaluation of a commercial modified live equine arteritis virus vaccine for use in stallions. Proceedings of the Fifth International Conference on Equine Infectious Diseases, Lexington, 1987, University Press of Kentucky, Lexington, Kentucky, USA, 19-27.
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*
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NB: There are OIE Reference Laboratories for Equine viral arteritis (please consult the OIE Web site at: http://www.oie.int/eng/OIE/organisation/en_LR.htm).
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