Return to warmwell.com

Taken from http://oie.int/eng/normes/mmanual/A_00032.htm

Diagnostic Techniques

Introduction to Bluetongue

REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS

SUMMARY

Bluetongue (BT) is an infectious, noncontagious, insect-borne viral disease of sheep and domestic and wild ruminants, such as goats, cattle deer, bighorn sheep, most species of African antelope and various other Artiodactyla. The outcome of infection ranges from inapparent in the vast majority of infected animals to fatal in a proportion of infected sheep, goats, deer and wild ruminants. Clinical signs of disease, when they appear in domestic and wild ruminants, include a febrile response characterised by inflammation and congestion, facial oedema and haemorrhages, and ulceration of the mucous membranes. In severe cases the tongue may show intense hyperaemia, and become oedematous and protrude from the mouth. Hyperaemia may extend to other parts of the body particularly the groin, axilla and perineum. There is often severe muscle degeneration. Dermatitis may cause wool breaks. Sheep may become lame as a result of coronitis, inflammation of the coronary band of the hoof, or skeletal myopathy. A similar severe disease of wild ruminants is caused by epizootic haemorrhagic disease virus (EHDV), which, like BT virus (BTV) , is a member of the Orbivirus genus, but is classified in a separate serogroup. EHD may occasionally cause clinical signs in cattle that appear to be similar to bluetongue.
 
Identification of the agent: BTV is a member of the Orbivirus genus of the family Reoviridae. Within the genus there are 14 serogroups. The BT serogroup contains 24 serotypes. The former are differentiated by immunological tests that detect viral proteins that are conserved within each serogroup. Most serogroups appear to be immunologically distinct, but there is considerable cross-reaction between members of the BT and EHD serogroups. The serotype of individual viruses in each serogroup is identified on the basis of neutralisation tests. Complete BTV particles are double-shelled and the outer layer contains two proteins, one of which is the major determinant of serotype specificity. The inner icosahedral core contains two major and three minor proteins and ten species of double-stranded RNA. VP7 is a major core protein possessing the serogroup-specific antigens. Virus identification traditionally requires isolation and amplification of the virus in tissue culture, and the subsequent application of serogroup- and serotype-specific tests. Recently, the application of polymerase chain reaction (PCR) technology has permitted very rapid amplification of BTV RNA in clinical samples, and PCR-based procedures are now available to provide information on virus serogroup and serotype.
 
Serological tests: Serological responses in ruminants appear some 7-14 days after BTV infection and are generally long-lasting. Until recently, tests such as agar gel immunodiffusion and indirect enzyme-linked immunosorbent assay (ELISA) were used to detect BT serogroup-specific antibody, although these tests had the major drawback of being unable to consistently distinguish between antibodies to viruses in the BT and EHD serogroups. A monoclonal antibody-based competitive ELISA has solved this problem and competitive ELISAs to specifically detect anti-BTV antibodies are recommended. Current procedures to determine the serotype-specificity of antibodies in sera are cumbersome because they require determination of the capacity of test sera to inhibit the infectivity of panels of known virus serotypes in time-consuming neutralisation tests.
 
Requirements for vaccines and diagnostic biologicals: Live, attenuated vaccines that are serotype-specific are used in several countries of the world, such as South Africa, where nonvaccination may lead to outbreaks of disease. Attenuated viruses are prepared by serial passage of field virus in embryonated chicken eggs or cultured cells. Following serial passage, virulence is attenuated and, concomitantly, viruses replicate to lower titres in sheep. Attenuated virus vaccines are teratogenic and should not be administered to pregnant sheep during the first half of pregnancy as this may cause fetal death and abnormalities. In determining an appropriate degree of attenuation for vaccine purposes, a compromise is sought between a level of replication sufficient to reduce virulence but stimulate protective immunity in sheep, and a need to reduce the titre of virus in the blood in an attempt to prevent infection of feeding insects. Procedures to test vaccine efficacy and teratogenic potential in sheep are easily performed. In contrast, few studies have been carried out to determine whether or not attenuated virus can be transmitted by insects from vaccinated sheep to other animals. The fact that attenuated viruses are teratogenic makes determination of transmissibility very important especially if live virus vaccines are used in countries for the first time.
 

A. INTRODUCTION

Midges of the genus Culicoides transmit bluetongue virus (BTV) to and from susceptible animals, having become infected by feeding on viraemic vertebrates. After a replication period of 6-8 days, and following its appearance in the salivary gland, the virus can be transmitted to a vertebrate host during a blood meal. Infected midges remain infective for life. The central role of the insect in BT epidemiology ensures that prevalence of the disease is governed by ecological factors, such as high rainfall, temperature, humidity and soil characteristics, which favour insect survival (6). In many parts of the world therefore, the disease has a seasonal occurrence (13).
 
BT is an infectious, noncontagious disease of sheep and other domestic and wild ruminants, such as goats, cattle, deer, bighorn sheep, most species of African antelope and other Artiodactyla. The outcome of infection ranges from inapparent in the vast majority of infected animals to fatal in a proportion of infected sheep, deer and some wild ruminants. Although the frequency of BTV infection of cattle is generally higher than in sheep, overt disease in cattle is rare and the signs, when they occur, are much milder than those observed in sheep. In nondomestic ruminants, the disease can vary from an acute haemorrhagic disease with high mortality, as observed in white-tailed deer (Odocoilus virginianus), to an inapparent disease as seen in the North American elk (Cervus canadensis). Epizootic haemorrhagic disease virus (EHDV) can produce a disease in wild ruminants with clinical manifestations identical to those observed in response to BTV infection.
 
Clinical signs of disease in domestic and wild ruminants range from subclinical in the vast majority of cases to an acute febrile response characterised by inflammation and congestion, leading to oedema of the face, eyelids and ears, and haemorrhages and ulceration of the mucous membranes. Extensive erosions can develop in the cheeks and on the tongue opposite molar teeth. The tongue may show intense hyperaemia and become oedematous, protrude from the mouth and, in severe cases become cyanotic. Hyperaemia may extend to other parts of the body particularly the groin, axilla and perineum. There is often severe muscle degeneration. Dermatitis may cause wool breaks. Coronitis with haemorrhage of the coronary band of the hoof is common and may cause lameness. When sheep die as a result of acute BT disease, the lungs may show interalveolar hyperaemia, severe alveolar oedema and the bronchial tree may be filled with froth. The thoracic cavity may contain several litres of plasma-like fluid and the pericardial sac may show many petechial haemorrhages. Most cases show a distinctive haemorrhage near the base of the pulmonary artery (11).
 
BTV is a member of the Orbivirus genus, currently one of nine genera classified in the family Reoviridae. Within the Orbivirus genus, 14 groups are differentiated on serological grounds. The best-studied Orbiviruses are in the BT, EHD and African horse sickness (AHS) serogroups. Within the serogroups, individual members are differentiated on the basis of neutralisation tests, and 24 serotypes of BTV have been described to date. There is significant immunological cross-reactivity between members of the BT and EHD serogroups (16). Details of EHD-specific tests will not be provided in this chapter.
 
BTV particles are composed of three protein layers. The outer layer contains two proteins, VP2 and VP5. VP2 is the major neutralising antigen and determinant of serotype specificity. It is also responsible for haemagglutination and the binding of BTV to mammalian cells. The ability of a MAb to VP5 to neutralise AHS virus (AHSV) and react with the equivalent protein of BTV and EHDV confirms a role for VP5 in neutralisation of Orbiviruses and highlights the extent of immunological cross-reactivity between members of the different Orbivirus serogroups (24). Removal of the outer VP2/VP5 layer leaves a bi-layered icosahedral core particle that is composed of two major proteins, VP7 and VP3, three minor proteins and the ten species of double-stranded RNA. VP7 is a major determinant of serogroup specificity and the site of epitopes used in competitive enzyme-linked immunosorbent assay (C-ELISA) to detect anti-BTV antibody. VP7 can also mediate attachment of BTV to insect cells (39). VP7 subunits consist of two domains.
 

B. DIAGNOSTIC TECHNIQUES

1.    Identification of the agent (a prescribed test for international trade)
 
      a)    Virus isolation
 
            The same diagnostic procedures are used for domestic and wild ruminants. A number of virus isolation systems are in common use, but two of the most efficient are embryonated chicken eggs (ECE) and sheep. Identification of BTV following inoculation of sheep may be a useful approach if the titre of virus in the sample blood is very low, as may be the case several weeks after virus infection. Attempts to isolate virus in cultured cells in vitro may be more convenient, but the success rate is frequently much lower than that achieved with in-vivo systems. Within a virus population not all BTV particles are identical at the genetic and amino acid level and only a small, perhaps minute, proportion of viruses present in the blood of infected animals may have appropriate amino acid sequences in key viral proteins to bind to and replicate in cells in culture. This may be the reason why direct inoculation on to cultured cells of viraemic blood that contains a relatively small number of virus particles is an inefficient way to isolate BTV. A high-titre virus preparation, and one more likely to contain virus that has the ability to replicate in tissue culture, is most readily generated by one or at most two passages in ECE. Cell culture has proven to be a more sensitive technique for isolation of EHDV.
 
            .    Isolation in embryonated chicken eggs
 
            i)    Blood is collected from febrile animals into an anticoagulant such as heparin; EDTA (ethylamine diamine tetra-acetic acid) or sodium citrate, and the blood cells are washed three times with sterile phosphate buffered saline (PBS). Washed cells are resuspended in PBS or isotonic sodium chloride and either stored at 4C or used immediately for attempted virus isolation.
 
            ii)    For long-term storage where refrigeration is not possible blood samples are collected in oxalate-phenol-glycerin (10). If samples can be frozen, they should be collected in buffered lactose peptone or 10% dimethyl sulphoxide (36) and stored at -70C or colder. The virus is not stable for long periods at -20C.
 
            iii)    In fatal cases, spleen and lymph nodes are the preferred organs for virus isolation attempts. Organs and tissues should be kept and transported at 4C to a laboratory where they are homogenised in PBS or isotonic saline, and used as described below, for blood cells.
 
            iv)    Washed blood cells are resuspended in distilled water or sonicated in PBS and 0.1 ml amounts inoculated intravascularly into 6-12 ECE that are 9-12 days old. This procedure is difficult to perform and requires practise. Details are provided by Clavijo et al. (8).
 
            v)    The eggs are incubated in a humid chamber at 33.5C and candled daily. Any embryo deaths within the first 24 hours post-inoculation are regarded as nonspecific.
 
            vi)    Embryos that die between days 2 and 7 are retained at 4C and embryos remaining alive at 7 days are killed. Infected embryos often have a haemorrhagic appearance. Dead embryos and those that live to 7 days are homogenised as two separate pools. Whole embryos, after removal of their heads, or specific organs such as the liver, are homogenised and the debris is removed by centrifugation.
 
            vii)    Virus in the supernatant may be identified either directly by antigen-capture ELISA (18), or indirectly by antigen-detection methods, such as immunofluorescence or immunoperoxidase, after further amplification in cell culture, as described in the next section.
 
            viii)    If no embryos are killed following inoculation of sample material, an inoculum made from the first egg passage material may be repassaged in ECE or in cell culture.
 
            .    Isolation in cell culture
 
            Virus may also be added to mouse L, baby hamster kidney (BHK)-21, African green monkey kidney (Vero) or Aedes albopictus (AA) cells in culture. The efficiency of isolation is often significantly lower following direct addition to cultured cells compared with that achieved in ECE. Greatest efficiency of isolation in cell culture is achieved by first passaging ECE homogenates in AA cells, followed by either antigen detection procedures or additional passages in mammalian cell lines, such as BHK-21 or Vero. A cytopathic effect (CPE) is not necessarily observed in AA cells. Cell monolayers are monitored for the appearance of a CPE for 5 days at 37C in 5% CO2 with humidity. If no CPE appears, a second passage is made in cell culture.
 
            The identity of BTV in the culture medium of cells manifesting a CPE may be confirmed by a number of serological methods described below, including antigen-capture ELISA, immunofluorescence, immunoperoxidase, or virus neutralisation (VN) tests.
 
            .    Isolation in sheep
 
            i)    Sheep are inoculated with washed cells from 10 ml up to approximately 500 ml of blood, or 10-50 ml tissue suspension. Inocula are administered subcutaneously in 10-20 ml aliquots. Large volumes may aid in the virus isolation attempts and should be administered intravenously.
 
            ii)    The sheep are held for 28 days and checked for antibody using the agar immunodiffusion (1) test or C-ELISA as described below.
 
      b)    Immunological methods
 
            .    Serogrouping of viruses
 
            Orbivirus isolates are typically serogrouped on the basis of their reactivity with specific standard antisera that detect proteins, such as VP7 that are conserved within each serogroup. The cross-reactivity between members of the BT and EHD serogroups raises the possibility that an isolate of EHDV could be mistaken for BTV on the basis of a weak immunofluorescence reaction with a polyclonal anti-BTV antiserum. For this reason, a BT serogroup-specific MAb can be used. A number of laboratories have generated such serogroup-specific reagents (3, 22). In contrast to serogrouping, the usual method of serotyping is by VN testing using methods described later. Commonly used methods for the identification of viruses to serogroup level are as follows.
 
            i)    Immunofluorescence
 
                  Monolayers of BHK or Vero cells on glass cover-slips are infected with either tissue culture-adapted virus or virus in ECE lysates. After 24-48 hours at 37C, or after the appearance of a mild CPE, infected cells are fixed with agents such as paraformaldehyde, acetone or methanol, dried and viral antigen detected using anti-BTV antiserum and standard immunofluorescent procedures.
 
            ii)    Antigen capture enzyme-linked immunosorbent assay (27)
 
                  Virus in ECE lysates, culture medium and infected insects may be detected directly. In this technique, virus and/or core particles are captured by antibody adsorbed to an ELISA plate and bound virus is detected using a second antibody. The capture antibody may be polyclonal or a serogroup-specific MAb. Serogroup-specific MAbs and polyconal antibody raised to baculovirus-expressed core particles have been used successfully to detect captured virus (18).
 
            iii)    Immunospot test (14)
 
                  Small volumes (2 l) of infected cell culture supernatant or lysed or sonicated infected cells are adsorbed to nitrocellulose and air-dried. Nonspecific binding sites are blocked by incubation in a solution containing skim milk protein. After incubation with a BT serogroup-reactive MAb, bound antibody is detected using horseradish peroxidase-conjugated anti-mouse IgG.
 
            .    Serotyping by virus neutralisation
 
            Neutralisation tests are type specific for the currently recognised 24 BTV serotypes and can be used to serotype a virus isolate, or can be modified to determine the specificity of antibody in sera. In the case of an untyped isolate, the characteristic regional localisation of BTV serotypes should generally obviate the need to attempt neutralisation by all 24 antisera, particularly when endemic serotypes have been identified.
 
            There is a variety of tissue culture-based methods available to detect the presence of neutralising anti-BTV antibody. Cell lines commonly used are BHK, Vero and L929. Four methods to serotype BTV are outlined briefly below. BTV serotype-specific antisera generated in guinea-pigs or rabbits have been reported to have less serotype cross-reactivity than those made in cattle or sheep. It is important that antiserum controls be included to ensure that an effective level of reference antiserum is used against comparable and standardised titres of reference and untyped virus.
 
            i)    Plaque reduction
 
                  The virus to be serotyped is diluted to contain approximately 100 plaque-forming units (PFU), and incubated with either no antiserum or with individual standard antisera to a panel of BTV serotypes. Virus/antiserum mixtures are added to monolayers of cells and the virus titre is determined by plaque assay. The unidentified virus is considered serologically identical to a standard serotype if the latter is run in parallel with the untyped virus in the test, and is similarly neutralised.
 
            ii)    Plaque inhibition
 
                  Tests are performed in 90 mm diameter Petri dishes containing confluent cell monolayers that are infected with approximately 5 x 104 PFU standard or untyped virus. After adsorption and removal of inoculum, monolayers are overlaid with agarose. Standard anti-BTV antisera are added to individual filter paper discs and placed on the agarose surface. Dishes are incubated for at least 4 days. A zone of virus neutralisation, with concomitant survival of the cell monolayer, will surround the disc containing the homologous antiserum.
 
            iii)    Microtitre neutralisation
 
                  Approximately 100 TCID50 (50% tissue culture infective dose) of the standard or untyped virus is added in 50 l volumes to test wells of a flat-bottomed microtitre plate and mixed with an equal volume of standard antiserum diluted in tissue culture medium. Approximately 104 cells are added per well in a volume of 100 l, and after incubation for 4-6 days, the test is read using an inverted microscope. Wells are scored for the degree of CPE observed. Those wells that contain cells only or cells and antiserum, should show no CPE. In contrast, wells containing cells and virus should show convincing CPE. The unidentified virus is considered to be serologically identical to a standard BTV serotype if both are neutralised in the test to a similar extent.
 
            iv)    Fluorescence inhibition test (5)
 
                  This rapid and simple neutralisation assay requires varying concentrations of an unknown virus and standard concentrations of reference antisera. Virus isolates grown in cell culture are serially diluted starting and mixed with individual reference antisera in the wells of a Lab-Tek slide for 1 hour prior to addition of cells. After incubation for 16 hours, cells are fixed and probed by an immunofluorescent procedure using a BT serogroup-specific MAb. The serotype of the virus is indicated by the specificity of the antiserum causing the largest reduction in the number of fluorescent cells.
 
      c)    Polymerase chain reaction (a prescribed test for international trade)
 
            Primer-directed amplification of viral nucleic acid has revolutionised BT diagnosis (8, 26, 37). Polymerase chain reaction (PCR) techniques have allowed the rapid identification of BT viral nucleic acid in blood and other tissues of infected animals. Regarding international trade, PCR has allowed the identification of BT antibody-positive animals that are negative for viral nucleic acid, permitting their importation. PCR can also be used to 'serogroup' Orbiviruses and may ultimately be used to 'serotype' BTV within a few days of receipt of a clinical sample, such as infected sheep blood. Traditional approaches, which rely on virus isolation followed by virus identification serologically, may require at least 3-4 weeks to generate information on serogroup and serotype.
 
            Oligonucleotide primers used so far have been derived from RNA 7 (VP7 gene) (37), RNA 6 (NS1 gene) (9), RNA 3 (VP3 gene) (30), RNA 10 (NS3 gene) (4) and RNA 2 (VP2 gene) (26). The size of the amplified transcripts is usually small - in the order of several hundred nucleotides - but can also be a full-length gene. In the procedure described in detail below, a 101-nucleotide stretch of RNA 6 is amplified. Primers derived from the more highly conserved genes, such as VP3, VP7 and NS1, may be used for serogrouping (i.e. will react with all members of the BT serogroup), while primers for which the sequence was determined from VP2 gene sequences provide information on virus serotype. A multiplex PCR assay that depends on the size of the amplified products has been used to identify the five North American BTV serotypes, both alone and in mixtures, in a single reaction (19).
 
            The nucleic acid sequence of cognate BTV genes may differ with the geographical area of virus isolation (17). This has provided a unique opportunity to complement studies of BTV epidemiology by providing information on the potential geographical origin of virus isolates, a process termed genotyping or topotyping. Thus, determination of the nucleic acid sequence of portions of RNA 3 and RNA 6 may provide information on whether the virus came form Australia, North America or South Africa. It appears likely that sequencing of BTV isolates from other parts of the world may permit finer discrimination of geographical origin. However, the relationship between sequence and geographical origin may not be straightforward. Genotypes specific to geographical l cations were not as clearly defined by PCR analyses of RNA genome segment 7 (38) as they appeared to be using RNA genome segment 3 (17). The development of topotyping as an epidemiological tool thus depends on the acquisition of sequence data for BTV isolates from many and diverse regions of the world and availability of the data in readily accessible data banks. In principle, given a large enough RNA 2 sequence database, it should ultimately be possible to determine rapidly virus serotype by PCR amplification of RNA 2. To facilitate this process new sequence data derived from both characterised and uncharacterised BTV isolates should be made widely available by submitting the data to web sites such as:
 
            http://www.iah.bbsrc.ac.uk/dsRNA_virus_proteins/ and http://www.iah.bbsrc.ac.uk/dsRNA_virus_proteins/btv_sequences.htm.
 
            The web site http://www.iah.bbsrc.ac.uk/dsRNA_virus_proteins/btv2-segment-2-tree.htm provides phylo-genetic tree analyses of BTV isolates based on the sequence of RNA2. These compiled data will provide a resource for epidemiological studies, the identification of new isolates and the design of new primers for further reverse transcription (RT) PCR and possibly serotype-specific assays for BTV.
 
            It has been observed that BTV nucleic acid can be detected by PCR from the blood of infected calves and sheep at least 30 days, and sometimes over 90 days, after the virus can be isolated. When blood that was positive for virus isolation (infectious) and blood that was negative for virus isolation but positive by PCR (PCR-detectable only) were inoculated into or fed to the vector, Culicoides sonorensis, it was shown that the virus was amplified and transmitted only by vectors exposed to infectious blood. Vectors exposed to PCR-detectable only blood did not amplify or transmit the BTV (23). Because of this, PCR-based diagnostics should be interpreted with caution. The PCR procedure will detect virus-specific nucleic acid, but this does not necessarily indicate the presence of infectious virus.
 
            The capacity of PCR assays to detect very small numbers of nucleic acid molecules means that such tests are exquisitely sensitive to contamination by extraneous nucleic acids. The latter may include any primers in use in the laboratory or previously amplified polynucleotides. It is critical therefore to have a 'clean' area containing all equipment necessary for reagent and test preparation and a separate area with its own equipment for amplification. Latex gloves should be worn and changed frequently at all stages of the procedure, particularly after working with sample RNA or amplified DNA. This will help protect reagents and samples from contamination by ubiquitous RNases and other agents and from cross-contamination by DNA. The possibility of false positives, due to sample contamination, highlights the importance of sequencing PCR products to determine, for example, if the amplified sequence is identical to or different from that of the positive control. False negatives, due for example to poor sample quality or inappropriate primers, may be identified following the failure to amplify both BTV and a host gene, such as globin, from extracts of infected cells.
 
            The PCR assay described here involves three separate procedures. In the first, BTV RNA is extracted from blood using a chaotropic agent such as guanidine thiocyanate (GuSCN) to denature protein and release viral RNA. A number of commercial kits are available for this purpose and the protocol below describes the use of one such kit, IsoQuick (Orca Research, Bothell, Washington, United States of America [USA]). The reagents provided with the kit are numbered and their use is indicated in the protocol below. Other kits are available and one, TRIZOL (Life Technologies, Grand Island, New York, USA), is particularly useful for the extraction of viral nucleic acid from spleen or blood clots. Operators should follow the procedures specified in each kit and use reagent solutions either provided or recommended for the kit of their choice. The second procedure is the denaturation of viral double-stranded RNA and reverse transcription to generate cDNA, which is amplified by PCR. In the procedure described below, the SuperscriptTM Preamplification System (Life Technologies) is used to transcribe viral RNA, and reagents from Perkin-Elmer are used for the PCR. Equivalent kits and reagents are available from other sources. The final step of the process is the analysis of the PCR product by electrophoresis. Procedures used to determine the sequence of the amplified product are not described here.
 
            .    Extraction of viral RNA
 
            i)    Whole blood is collected from test and uninfected control animals in EDTA tubes and centrifuged at 800-1000 g for 10 minutes. The plasma is aspirated and the red blood cells (RBCs) are gently resuspended in sterile PBS. RBCs are pelleted by centrifugation at 1000 g for 10 minutes and the supernatant is removed.
 
            ii)    Next, 400 l of test RBCs is added to each of four 1.7 ml microcentrifuge tubes, and 400 l of control RBCs is added to each of two microcentrifuge tubes. An equal volume of RNase-free water is added to each tube and the tubes are vortexed briefly to mix and lyse the cells. Two tubes containing test RBCs are frozen at -70C for repository purposes and the extraction is continued in duplicate.
 
            iii)    Lysed test and control RBCs are centrifuged at 12,000-16,000 g for 10 minutes and the supernatant is discarded. Next, 800 l RNase-free water is added and the tubes are vortexed and centrifuged again at the same speed for 10 minutes. The supernatant is removed and the RBC pellet is drained.
 
            iv)    A small volume of BTV (e.g. 5 l containing from 103 to 107 PFU) is added to one of two control RBC pellets. This is the positive control. The other control RBC pellet remains as the negative control.
 
            v)    Next, 75 l of sample buffer (IsoQuick reagent A) is added to each pellet, and the pellets are then vortexed vigorously, followed by the addition of 125 l of the GuSCN-containing lysis solution (IsoQuick reagent 1). The mixture is vortexed vigorously for 30 seconds.
 
            vi)    Before use the extraction matrix provided with the kit (IsoQuick reagent 2 plus dye 2A) is shaken vigorously and 500 l is added to the sample lysates. Then, 400 l extraction buffer (IsoQuick reagent 3) is added and the tubes are vortexed for 10 seconds.
 
            vii)    The tubes are incubated at 65C for 10 minutes, vortexed briefly after 5 minutes and centrifuged at 12,000 g for 5 minutes.
 
            viii)    The aqueous phase (500 l) is transferred to a new microcentrifuge tube and an equal volume of extraction matrix (IsoQuick reagent 2) is added. The tubes are vortexed for 10 seconds and centrifuged at 12,000 g for 5 minutes.
 
            ix)    The aqueous phase (330 l) is transferred to a new microcentrifuge tube and a 10% volume (33 l) of sodium acetate (IsoQuick reagent 4) and 365 l isopropanol are added. After gentle mixing, the tubes are placed at -20C for from 20 minutes to 1 hour.
 
            x)    The RNA is pelleted by centrifugation at 12,000 g for 10 minutes. The supernatant is decanted and 1.0 ml 70% ethanol is added and mixed gently. After centrifugation at 12,000 g for 5 minutes, the supernatant is decanted and 1.0 ml 100% ethanol is added. The tubes are stored at -70C until ready for use in the RT-PCR.
 
            .    Reverse-transcription polymerase chain reaction
 
            i)    RNA in ethanol is centrifuged at 12,000 g for 5 minutes. The ethanol is decanted and the tubes are inverted and allowed to drain. The pellet, which may not be obvious, must not be allowed to dry out because this makes resuspension difficult. A dry pellet is also likely to fall out of the inverted tube.
 
            ii)    Next, 12 l RNase-free water is added to each tube, mixed and heated at 65C for 5-10 minutes. The samples are placed in ice.
 
            iii)    In a 'clean' biohazard hood, stock solutions containing 200 pmol/l of primers A, B, C and D are prepared in RNase-free water and stored at -70C.
 
                  First stage PCR primers (to amplify RNA 6 from nucleotide 11 to 284)
                  Primer A: 5'-GTT-CTC-TAG-TTG-GCA-ACC-ACC-3'
                  Primer B: 5'-AAG-CCA-GAC-TGT-TTC-CCG-AT-3'
 
                  Nested PCR primers (to amplify RNA 6 from nucleotide 170 to 270)
                  Primer C: 5'-GCA-GCA-TTT-TGA-GAG-AGC-GA-3'
                  Primer D: 5'-CCC-GAT-CAT-ACA-TTG-CTT-CCT-3'
 
            iv)    Primer stock solutions are diluted to a concentration of 15-20 pmol/l. Primers for the first stage PCR reaction are prepared by mixing equal volumes of A and B. Primers for the nested PCR reaction are prepared by mixing equal volumes of C and D. Small aliquots of pooled primer mixes are frozen at -20C.
 
            v)    PCR reaction tubes are labelled and, for first stage synthesis, 4.0 l of primer (A + B) mix is added to each tube. The tubes are held on ice.
 
            vi)    In a 'clean' fume hood methylmercuric hydroxide is diluted to 50 mM (1/20 dilution) and 2-mercaptoethanol is diluted to 350 mM (1/40 dilution) in RNase-free water. Methylmercuric hydroxide and 2-mercaptoethanol are considered to be extremely and highly toxic, respectively. Use both chemicals with extreme care and dispose of them and pipette tips as required by safety regulations.
 
            vii)    Next, 4 l of test and positive and negative control RNA samples (step ii) are added to 4 l of the primer mix in PCR tubes (38).
 
            viii)    To each PCR tube 2.0 l of the 1/20 dilution of methylmercuric hydroxide is added with gentle mixing and allowed to sit at room temperature for 10 minutes prior to adding 2.0 l of the 1/40 dilution of 2-mercaptoethanol. For safety reasons, some laboratories use formamide instead of methylmercuric hydroxide for double-stranded RNA denaturation. However, for optimum sensitivity, methylmercuric hydroxide is preferred.
 
            ix)    In a 'clean' hood a cDNA mix is prepared containing the following reagents in sufficient volume for the number of samples being tested. The amount given is per sample and the reagents are contained in the SuperscriptTM Preamplification System (Life Technologies).
 
                  10 x SuperscriptTM buffer (200 mM Tris/HCl, pH 8.4, and 500 mM KCl)    2.0 l
                  MgCl2 (25 mM)     2.0 l
                  dNTP mix (10 mM each dATP, dCTP, dGTP, dTTP)    1.25 l
                  Dithiothreitol (DTT) (0.1 M)     2.0 l
                  Reverse transcriptase (200 units/l)    0.75 l
 
            x)    Then, 8.0 l of the mix is added to each PCR tube to a final volume of 20.0 l.
 
            xi)    The PCR tubes are placed in a thermal cycler, such as GeneAmpTM PCR System 9600, which is programmed for reverse transcription as follows:
 
                  Hold 44C    50 minutes
                  Hold 4C     Forever
 
            xii)    The tubes are removed from the thermal cycler and 1.0 l RNase H and a wax bead are added to each tube. The cycler is programmed as follows:
 
                  Hold 37C    20 minutes
                  Hold 98C    4 minutes
                  Hold 4C    Forever
 
            xiii)    In a 'clean' hood a first stage amplification mix is prepared containing the following reagents and in a volume sufficient for the number of samples being tested. All these reagents except water are available from Perkin-Elmer. The amount given is per sample.
 
                  RNase-free water     62.0 l
                  10 x PCR Perkin-Elmer buffer (100 mM Tris/HCl, pH 8.3, and 500 mM KCl)    7.0 l
                  MgCl2 (25 mM)     7.0 l
                  dNTP mix (2.5 mM each dATP, dCTP, dGTP, dTTP)     4.0 l
                  Taq DNA polmerase (5 units/l)     0.85 l
 
            xiv)    The first stage mix is removed from the 'clean' area to the thermal cycling area and 80 l is overlaid in each sample tube. The wax layer must not be pierced. Each tube should now contain 101 l.
 
            xv)    The tubes are placed in the thermal cycler, which is programmed as follows (correct for GeneAmp PCR System 9600 - programmes for other thermal cyclers would need to be determined) for first stage amplification:
 

 
One cycle: Hold 95C 3 minutes
Hold 58C 20 seconds
 
 
Hold 72C
 
30 seconds
 
40 cycles: Hold 95C 20 seconds
Hold 58C 20 seconds
 
 
Hold 72C
 
20 seconds
 
One cycle: Hold 95C 20 seconds
Hold 58C 20 seconds
Hold 72C 5 minutes
Hold 4C Forever

 
            xvi)    PCR reaction tubes are prepared for the nested reaction in a 'clean' hood 15 minutes before cycling is complete, and held on ice:
 
                  Rnase-free water    17 l per tube
                  Nested primer mix (C+D)    4.0 l per tube
                  Wax bead
 
            xvii)    When first stage amplification is complete, the tubes are removed from the thermal cycler and placed in a biological safety cabinet (not the 'clean' hood). Then, 1.5 l of the first stage product is transferred to the corresponding nested PCR tube containing primer, water and a wax bead.
 
            xviii)    The tubes are placed in the thermal cycler, which is programmed as follows for wax layer formation:
 
                  Hold 98C    4 minutes
                  Hold 4C    Forever
 
            xix)    In a 'clean' hood the nested mix of the following reagents is prepared in sufficient volume for the number of samples being tested. The reagents used are the same as in the first stage (step xii). The amount given is per sample.
 
                  RNase-free water    17.0 l
                  10 x PCR buffer    5.0 l
                  MgCl2     3.5 l
                  dNTP mix    4.5 l
                  Taq DNA polymerase    0.5 l
 
            xx)    The nested mix is removed from the 'clean' hood to the thermal cycler and 30 l is overlaid into each sample tube. Each tube should now contain 52 l.
 
            xxi)    The tubes are placed in the thermal cycler, which is programmed as follows for nested amplification. After completion, the tubes are held at 4C or at -20C until electrophoresis:
 

 
One cycle: Hold 95C 3 minutes
Hold 58C 20 seconds
 
 
Hold 72C
 
30 seconds
 
40 cycles: Hold 95C 20 seconds
Hold 58C 20 seconds
 
 
Hold 72C
 
20 seconds
 
One cycle: Hold 95C 20 seconds
Hold 58C 20 seconds
Hold 72C 5 minutes
Hold 4C Forever

 
            .    Electrophoretic analysis of PCR product
 
            i)    First, 1 x TBE buffer (0.045 mM Tris/borate, pH 8.6, and 1.5 mM EDTA) is prepared from a x10 stock solution. For the Bio-Rad Wide Mini-Sub cell system, 700 ml buffer is prepared (100 ml for the gel and 600 ml for the tank buffer).
 
            ii)    A 3% solution of NuSieve 3/1 agarose (FMC Bioproducts, Rockland, Maine, USA) or an equivalent is prepared in TBE buffer. The solution is boiled until the agarose is completely dissolved, and then allowed to cool to 40C. Ethidium bromide is added to a concentration of 0.5 g/ml to both the agarose and the tank buffer. Ethidium bromide is a mutagen and is toxic. Gloves, protective clothing, and eye-wear must always be worn.
 
            iii)    The ends of the electrophoresis tray are taped and the agarose solution is poured. The comb is inserted and the agarose is allowed to solidify on a level surface for 30-60 minutes. The comb and the tape are gently removed from the electrophoresis tray.
 
            iv)    Pour the tank buffer into the electrophoresis apparatus and insert the tray with the agarose so that the buffer covers the agarose.
 
            v)    Test and positive and negative control samples are prepared for electrophoresis in 0.65 ml microcentrifuge tubes as follows:
 
                  Gel-loading solution (Cat. G-2526, Sigma, St Louis, Missouri, USA)    5.0 l
                  Amplified DNA from each of the PCR tubes and an extra tube is set up for a DNA ladder:    15.0 l
                  Gel-loading solution (Cat. G-2526, Sigma, St Louis, Missouri, USA)    5.0 l
                  100 base-pair ladder (Cat. 15268-019, Life Technologies, Grand Island, New York, USA)    1.0 l
 
            vi)    Samples are loaded into the appropriate wells in the gel and run at 65-75 volts for 1-1.5 hours or until the dye has travelled about half the length of the gel. The gel is transferred to a transilluminator and photographed for a permanent record. Use protective eye-wear to visualise the gel bands.
 
            vii)    BT-positive samples will have a band of 101 base pairs. For the test to be valid, the positive control must show a band of the correct size, and the negative and 'no RNA' controls show no band. Samples are considered to be positive if there is a band of the same size as the positive control. Duplicate samples should show the same reaction. If there is disparity, the test should be repeated.
 
            viii)    A destaining bag (Ameresco, Solon, Ohio, USA) is placed in the tank buffer overnight to remove the ethidium bromide. The buffer can then be poured down the drain and the destaining bag, after reuse 10-15 times, should be placed in a properly identified ethidium bromide waste container and ultimately incinerated.
 
            Kits and reagents for two prescribed serological tests - the agar gel immunodiffusion (AGID) test and the C-ELISA - are available from three licensed manufacturers in the USA (VMRD, P.O. Box 502, Pullman, Washington 99163, USA; or Veterinary Diagnostic Technology, 4980 Van Gordon Street, Suite 101, Wheat Ridge, Colorado 80033, USA; or Diagxotics, 27 Cannon Road, Wilton, Connecticut 06897, USA). The C-ELISA reagents are available from the European Union 'Community Reference Laboratory' for BTV (Pirbright Laboratory, Ash Road, Pirbright, Woking GU24 0NF, United Kingdom).
 
2.    Serological tests
 
      Anti-BTV antibody generated in infected animals can be detected in a variety of ways that depend on the sensitivity and type of test used. Both serogroup-specific and serotype-specific antibodies are elicited and if the animal was not previously exposed to BTV, the neutralising antibodies generated are specific for the infecting virus. Multiple infections with different BTV serotypes lead to the production of antibodies capable of neutralising serotypes to which the animal has not been exposed. There are two explanations for this phenomenon. First, several serotypes share monoclonal MAb-defined neutralisation epitopes. Secondly, serotypes also share a large number of epitopes that are present in a neutralising conformation in one serotype, but in non-neutralising conformations in other serotypes.
 
      a)    Complement fixation
 
            A complement fixation (7) test to detect BTV antibodies was widely used until 1982, when it was largely replaced by the AGID test although the CF test is still used in some countries.
 
      b)    Agar gel immunodiffusion (a prescribed test for international trade)
 
            The AGID test to detect anti-BTV antibodies is simple to perform and the antigen used in the assay relatively easy to generate. Since 1982, the test has been the standard testing procedure for international movement of ruminants. However, one of the disadvantages of the AGID used for BT is its lack of specificity in that it can detect antibodies to other Orbiviruses, particularly those in the EHD serogroup. Thus AGID positive sera may have to be retested using a BT serogroup-specific assay. The lack of specificity and the subjectivity exercised in reading the results have encouraged the development of ELISA-based procedures for the specific detection of anti-BTV antibodies. The preferred format, a C-ELISA is described in the Section B.2.c.
 
            .    Test procedure
 
            i)    A 2.8 mm thick layer of 0.9% agarose in 0.85% NaCl is prepared and circular wells, 4.0 mm in diameter and 2.4 mm apart, are cut out with six wells arranged around a central well.
 
            ii)    Viral antigen is prepared by generating a crude soluble preparation from BHK or Vero cells infected with a single BTV serotype 24-48 hours previously. Antigen can be concentrated by precipitation or ultrafiltration.
 
            iii)    Three positive and three test sera are placed in alternate wells surrounding antigen in the central well and the plates are incubated at 20-25C in a humid environment for 24 hours.
 
            iv)    A series of precipitin lines form between the antigen and known positive sera and lines generated by strong positive test sera will join up with those of the positive controls. With weak positive samples the control lines bend toward the antigen and away from the test sample well, but may not form a continuous line between the control test wells. With negative samples, the precipitin lines will continue into the sample wells without bending toward the antigen.
 
            v)    All weak positive samples and other samples that produce questionable results should be repeated using wells that are 5.3 mm in diameter placed 2.4 mm apart or retested using the C-ELISA as described below.
 
      c)    Competitive enzyme-linked immunosorbent assay (a prescribed test for international trade)
 
            The BT competitive or blocking ELISA was developed to measure BTV-specific antibody without detecting cross-reacting antibody to other Orbiviruses (1, 3, 22, 28, 31). The specificity is the result of using one of a number of BT serogroup-reactive MAbs, such as MAb 3-17-A3 (3) or MAb 20E9 (22) or MAb 20E9 (21). The antibodies were derived in a number of laboratories, and although different, all appear to bind to the amino-terminal region of the major core protein VP7. In the C-ELISA, antibodies in test sera compete with the MAbs for binding to antigen. The following procedure for the C-ELISA has been standardised after comparative studies in a number of international laboratories.
 
            .    Test procedure
 
            i)    First, 96-well microtitre plates are coated at 4C overnight or 37C for 1 hour with 50-100 l of either tissue culture-derived sonicated cell culture antigen (3) of the major core antigen VP7 expressed in either baculovirus (29) or yeast (25) and diluted in 0.05 M carbonate buffer, pH 9.6.
 
            ii)    The plates are washed five times with PBST (0.01 M PBS containing 0.05% or 0.1% Tween 20, pH 7.2).
 
            iii)    Next, 50 l of test sera is added in duplicate at a single dilution, either 1/5 (1) or 1/10 (22) in PBST containing 3% bovine serum albumin (BSA).
 
            iv)    Immediately, 50 l of a predetermined dilution of MAb diluted in PBST containing 3% BSA is added to each well. MAb control wells contain diluent buffer in place of test serum.
 
            v)    Plates are incubated for 1 hour at 37C or 3 hours at 25C, with continuous shaking.
 
            vi)    After washing as described above, wells are filled with 100 l of an appropriate dilution of horseradish peroxidase-labelled rabbit anti-mouse IgG (H+L) in PBST containing 2% normal bovine serum.
 
            vii)    Following incubation for 1 hour at 37C, the conjugate solution is discarded and plates are washed five times using PBS or PBST. Wells are filled with 100 l substrate solution containing 1.0 mM ABTS (2,2'-azino-bis-[3-ethylbenzothiazoline-6-sulphonic acid]), 4 mM H2O2 in 50 mM sodium citrate, pH 4.0, and the plates are shaken at 25C for 30 minutes. (Other substrates may be used and the reaction continued with shaking for an appropriate length of time to permit colour development.)
 
            viii)    The reaction is stopped by addition of a stopping reagent, such as sodium azide.
 
            ix)    After blanking the ELISA reader on wells containing substrate and stopping reagent, the absorbance values are measured at 414 nm. Results are expressed as per cent inhibition and are derived from the mean absorbance values for each sample by the following formula.
 
                  % inhibition = 100 - [(Mean absorbance test sample)/(Mean absorbance MAb control)] x 100.
 
                  NB: Some laboratories prefer to use a negative control serum that has previously been shown to have a percentage inhibition of zero as an alternative to the MAb control.
 
            x)    Percentage inhibition values >50% are considered to be positive. Inhibition between 40% and 50% is considered to be suspicious. The results of the duplicates of test sera can vary as long as they do not lie either side of the chosen inhibition value.
 
            xi)    Strong and weak positive sera and a negative serum should be included on each plate. The weak positive should give 60-80% inhibition and the negative should give less than 40% inhibition.
 

C. REQUIREMENTS FOR VACCINES AND DIAGNOSTIC BIOLOGICALS

Of several vaccine options available, namely live attenuated, killed or recombinant, only attenuated virus vaccines are in current use in several countries. In South Africa, for example, they have been used for over 40 years and are known to induce an effective and lasting immunity (11). Although the efficacy of inactivated virus vaccines has been investigated in some laboratory studies (15), they do not appear to have been used in the field. There are several options for the development of recombinant BTV vaccines, including live virus delivery of BTV neutralisation antigens and the virus-like particles (VLP) generated in infected insect cells by recombinant baculoviruses expressing the four major BTV coat proteins VP2, 3, 5 and 7. Only the latter has shown significant promise (33). However there is still much to determine, such as the longevity of the neutralising response generated to VLP, the need for multiple VLPs for different serotypes and the commercial scale up of VLP production in a cost effective and efficient process. The following description applies to attenuated virus vaccines.
 
1.    Seed management
 
      a)    Characteristics of seed
 
            The master or primary virus seed is prepared from a single plaque of serially passaged, attenuated BTV. Secondary seed lots, which are used as inocula for vaccine production, are usually not more than three passages beyond the primary seed lot. Primary seed virus must be free of contaminating bacteria, viruses, fungi and mycoplasmas, particularly pestivirus contamination, and must be shown to have the desired serotype specificity. Each primary seed virus lot should also be tested for transmissibility and reversion to virulence prior to vaccine manufacture. Samples of vaccine prepared from secondary seed virus at the maximum permitted passage level should be tested in sheep for avirulence, safety and immunogenicity.
 
      b)    Method of culture
 
            The first BT vaccines were propagated in ECE (2). More recently, several different cells have been used for tissue culture adaptation and serial passage. These include primary bovine embryo, lamb and fetal lamb kidney cells, and the continuous BHK cells. Cells used for attenuation must be thoroughly checked for the presence of contaminating viruses. Not only may continuous cell lines harbour oncogenic viruses, but primary cells may also contain a number of inapparent or latent virus infections, such as pestivirus contamination. For the latter, particular attention should be paid to the fetal bovine serum used in cell cultures, as it may be contaminated. Vaccine viruses have been attenuated by either passage in ECE, tissue culture cells or a combination of both.
 
      c)    Validation as a vaccine
 
            Attenuated BT vaccines must be safe and efficacious, and a brief description of appropriate tests for these parameters is given below. In addition, attenuated viruses should not revert to virulence during replication in vaccinated animals or be transmitted from such animals by insect vectors. The latter criterion is very important because insect-mediated transmission of attenuated virus from vaccinated to nonimmune animals, with the subsequent replicative steps in each host species, increases the possibility of reversion to virulence. Although tests for reversion to virulence and transmissibility are rarely, if ever performed, a brief description of what may be necessary is outlined.
 
            i)    Avirulence
 
                  A number of sheep, seronegative by BT C-ELISA, are inoculated with either the primary seed stock or an equal volume of tissue culture medium. Temperatures are noted twice daily. The animals are monitored at regular intervals over a period of 28 days for clinical signs and any local or systemic reactions to ensure avirulence and innocuity. Blood samples removed at regular intervals can be used to measure viraemia and antibody responses. The test shall be valid if all of the sheep inoculated with vaccine show evidence of virus growth and do not display signs of disease other than mild transient illness. In South Africa, a clinical reaction index (33) is calculated for each animal between days 4 and 14 and must be below a specific standard value.
 
            ii)    Safety
 
                  Safety tests for attenuated vaccines do not address the issue of their teratogenicity (34). Attenuated virus vaccines are teratogenic and should not be administered to pregnant sheep during the first half of pregnancy as this may cause fetal abnormalities and death (20).
 
            iii)    Efficacy
 
                  Vaccinated and unvaccinated sheep are challenged with virulent virus of the same serotype and animals are monitored for clinical signs of BT. Rectal temperatures are taken twice daily. Unvaccinated control sheep should show clinical signs of BT. However, it is difficult to be certain of the appearance of clinical disease following inoculation of sheep with certain BTV serotypes and isolates, and consequently, evidence of infection of unvaccinated control sheep may rest on the appearance of a temperature rise of at least 1.7C over the prechallenge mean. Pre- and post-vaccination sera are checked for the presence of neutralising antibody.
 
            iv)    Transmissibility
 
                  Procedures to determine if attenuated virus can be transmitted by insects that feed on vaccinated, viraemic sheep are difficult to perform and analyse statistically, and consequently, this criterion of vaccine validation is rarely sought. Laboratory data indicate that laboratory-adapted viruses can be transmitted by insect vectors (35). A suitable procedure to determine attenuated virus transmissibility requires that sheep be vaccinated and, during viraemia, that they be exposed to competent, uninfected Culicoides, which are then permitted to feed on uninfected animals that are monitored for the presence of BTV and anti-BTV antibody. Due to the fact that the titre of attenuated virus in the blood of vaccinated sheep is low, very large numbers of Culicoides would be needed and only a small proportion of these would become infected and live long enough to feed on and potentially transmit the virus to other uninfected sheep. It is difficult to design a laboratory experiment that takes account of the large numbers of vaccinated sheep and insects that would be present in field situations. However, in South Africa it is estimated that the minimum titre of virus circulating in the bloodstream of an animal must be at least 103 before feeding Culicoides become infected, although it has also been suggested that a lower titre may sometimes be infective. To select a suitable attenuated virus strain, whole blood is collected between days 4 and 14 after vaccination, and the virus titre is determined. Only attenuated viruses that generate titres under 103 are deemed to be acceptable as vaccines.
 
                  Current data indicate that during viraemia and in contrast to wild-type virus, laboratory-adapted strains of BTV may be found in the semen of bulls and rams (32). The implications of these observations for virus transmissibility are unclear.
 
            v)    Reversion to virulence
 
                  Validation studies confirm that attenuated viruses do not revert to virulence in vaccinated sheep. Consequently, if insects do not transmit attenuated viruses from vaccinated to unvaccinated animals, reversion to virulence becomes a theoretical possibility only. However, if attenuated viruses can be transmitted from vaccinated animals, reversion to virulence during a number of sheep-insect replication cycles becomes a distinct prospect. The only appropriate way to monitor for reversion to virulence under these circumstances is to compare the virulence of the vaccine virus with that which had been subject to several sheep-insect replication cycles as described above. As indicated, this is difficult to achieve. Consequently, the effect of a number of sheep-insect passages on the virulence of attenuated viruses has not been determined. In South Africa, it is accepted that if blood from vaccinated animals during the viraemic stages is serially passaged three times in sheep without reversion to virulence, the chances of reversion in the field will be infinitely small.
 
2.    Method of manufacture
 
      Attenuation of field isolates of BTV was first achieved by serial passage in ECE. More recently, it is clear that passage in cultured cells will also result in attenuation of virulence. No studies have been done to precisely relate passage number and extent of attenuation for individual virus isolates or serotypes. To prepare attenuated virus, field isolates are adapted to cell culture and passaged in vitro up to 40 times or more. Ideally, a number of plaque-purified viruses are picked at this stage and each is examined to determine the level of viraemia they generate and their ability to elicit a protective immune response in vaccinated sheep. The most suitable virus is one that replicates to low titre but generates a protective immune response, and this may represent the source of vaccine primary seed stock virus.
 
3.    In-process control
 
      All ingredients of animal origin, including serum and cells must be checked for the presence of viable bacteria, viruses, fungi or mycoplasmas.
 
4.    Batch control
 
      a)    Sterility
 
            Every batch of vaccine should be tested for the presence of viable bacteria, extraneous viruses, fungi or mycoplasmas, particularly pestivirus contamination. For example, in South Africa a pool of ten randomly selected ampoules are inoculated into soya broth and thioglycollate broth, and incubated at room temperature and 37C, respectively, for 14 days. If contaminated, the batch is disqualified.
 
      b)    Safety
 
            Every batch is safety tested in newborn and adult mice, guinea-pigs and sheep. If any adverse reactions or significant signs are noted, the test is repeated. Any increase in the body temperature of the target animal that is above the level expected for the particular strain of attenuated virus under test should be regarded as symptomatic. If the results are unsatisfactory, the batch is disqualified.
 
      c)    Potency
 
            Each batch is tested by inoculation of susceptible sheep. Prevaccination, and 21- and 28-day post-vaccination sera are tested by VN assay to determine neutralising antibody levels. In order to be passed, the antibody titre must be equal to or higher than a set standard based on international vaccine standards.
 
      d)    Duration of immunity
 
            Studies with live attenuated BTV vaccine have shown that antibodies in sheep may appear before day 10 post-vaccination, reach a maximum approximately 4 weeks later and persist for well over a year. There is a temporal relationship between the increase in neutralising antibody titre and clearance of virus from the peripheral circulation. Live attenuated BTV vaccines have been in use for over 40 years and are known to induce an effective and lasting immunity (13). Many serotypes of BTV are present in endemic areas of South Africa, and polyvalent vaccines are used. The inclusion of 15 serotypes in the vaccine means that an effective immune response is not generated to all serotypes, presumably because of the antigenic mass of individual serotype-specific antigens is small. In an attempt to broaden the response, vaccination is repeated annually (12).
 
      e)    Stability
 
            Stability should be tested over a period of 2 years. Vaccines in liquid and lyophilised forms are deemed to have shelf lives of 1 and 2 years, respectively. Each batch of vaccine is subjected to an accelerated shelf-life test by storing it at 37C for 7 days. It is then titrated and evaluated according to a set standard, as determined in the initial testing of the vaccine.
 
      f)    Precautions (hazards)
 
            The polyvalent vaccine is safe except if used in ewes during the first half of pregnancy. Lambs possessing colostral immunity cannot be effectively vaccinated before 6 months of age.
 
5.    Tests on the final product
 
      a)    Safety
 
            See C.4.b.
 
      b)    Potency
 
            See C.4.c.
 

ACKNOWLEDGEMENT

I wish to thank Dr Peter Mertens for helpful correspondence pertaining to the future of PCR diagnosis, Dr H. Aitchison for providing information relating to BT vaccine production and testing in South Africa, National Veterinary Services Laboratories, Ames, Iowa USA for providing the detailed bluetongue PCR protocol and all the reviewers of the previous chapter published in Terrestrial Manual in 2000.
 
1.    Afshar A., Thomas F.C., Wright P.F., Shapiro J.L. & Anderson J. (1989). Comparison of competitive ELISA, indirect ELISA and standard AGID tests for detecting bluetongue virus antibodies in cattle and sheep. Vet. Rec., 124, 136-141.
 
2.    Alexander R.A. & Haig D.A. (1951). The use of egg attenuated bluetongue in the production of a polyvalent vaccine for sheep. A. Propagation of the virus in sheep. Onderstepoort J. Vet. Res., 25, 3-15.
 
3.    Anderson J. (1984). Use of monoclonal antibody in a blocking ELISA to detect group specific antibodies to bluetongue virus. J. Immunol. Methods, 74, 139-149.
 
4.    Billinis C., Koumbati M., Spyrou V., Nomikou K., Mangana O., Panagiotidis C.A. & Papadopoulos O. (2001). Bluetongue virus diagnosis of clinical cases by a duplex reverse transcription-PCR: a comparison with conventional methods. J. Virol. Methods, 98, 77-89.
 
5.    Blacksell S.D. & Lunt R.A (1996). A simplified fluorescence inhibition test for the serotype determination of Australian bluetongue viruses. Aust. Vet. J., 73, 33-34.
 
6.    Calistri P., Goffredo M., Caporale V. & Meiswinkel R. (2003). The distribution of Culicoides imicola in Italy. Application and evaluation of current Mediterranean models based on climate. J. Vet. Med. B., 40, 132-138.
 
7.    Boulanger P. & Frand F.J. (1975). Serological methods in the diagnosis of bluetongue. Aust. Vet. J., 51, 185-189.
 
8.    Clavijo A., Heckert R.A., Dulac G.C. & Afshar A. (2000). Isolation and identification of bluetongue virus. J. Virol. Methods, 87, 13-23.
 
9.    Dangler C.A., De Mattos C.A., De Mattos C.C. & Osburn B.I. (1990). Identifying bluetongue virus ribonucleic acid sequences by the polymerase chain reaction. J. Virol. Methods, 28, 281-292.
 
10.    Edington A. (1900). South African horse-sickness: its pathology and methods of protective inoculation .J. Comp. Pathol. Therap., 13, 200-231.
 
11.    Erasmus B.J. (1975). Bluetongue in sheep and goats. Aust. Vet. J., 51, 165-170.
 
12.    Erasmus B.J. (1975). The control of bluetongue in an enzootic situation. Aust. Vet. J., 51, 209-210.
 
13.    Erasmus B.J. (1990). Bluetongue virus. In: Virus Infections of Ruminants, Dinter Z. & Morein B., eds. Elsevier, New York, USA, 227-237.
 
14.    Gard G.P. & Kirkland P.D. (1993). Bluetongue virology and serology. In: Australian Standard Diagnostic Techniques for Animal Diseases, Corner L.A. & Bagust T.J., eds. CSIRO Information Services, P.O. Box 89, East Melbourne, Victoria 3002, Australia.
 
15.    Ghalib H.W., Cherrington J.M. & Osburn B.I. (1984). Virological, clinical and serological responses of sheep infected with tissue culture adapted bluetongue virus serotypes 10, 11, 13 and 17.Vet. Microbiol., 10, 179-188.
 
16.    Gouet P., Diprose J.M., Grimes J.M., Malby R., Burroughs J.N., Zientara S., Stuart D.I. & Mertens P.P. (1999). The highly ordered double-stranded RNA genome of bluetongue virus revealed by crystallography. Cell, 97, 481-490.
 
17.    Gould A.R. (1987). The complete nucleotide sequence of bluetongue virus serotype 1 RNA3 and a comparison with other geographic serotypes from Australia, South Africa and the United States of America, and with other orbivirus isolates. Virus Res., 7, 169-183.
 
18.    Hawkes R.A., Kirkland P.D., Sanders D.A., Zhang F., Li Z., Davis R.J. & Zhang N. (2000). Laboratory and field studies of an antigen capture ELISA for bluetongue virus. J. Virol. Methods, 85, 137-149.
 
19.    Johnson D.J., Wilson W.C. & Paul P.S. (2000). Validation of a reverse transcriptase multiplex PCR test for the serotype determination of U.S. isolates of bluetongue virus. Vet. Microbiol., 76, 105-115.
 
20.    Johnson S.J., Polkinghorne I.G., Flanagan M., & Townsend W.L. (1992). The Australian experience: results of a bluetongue vaccination program. In: Bluetongue, African Horsesickness and Related Orbiviruses, Walton T.E. & Osburn B., eds. CRC Press, Boca Raton, Florida, USA, 868-873.
 
21.    Lelli R., Portanti O., Langella V., Luciani M., Di Emidio B. & Conte A.M. (2003). Production of a competitive ELISA kit for the serological disgnosis of bluetongue disease. Veterinaria Italiana, 39, 47.
 
22.    Lunt R.A., White J.R. & Blacksell S.D. (1988). Evaluation of a monoclonal antibody blocking ELISA for the detection of group-specific antibodies to bluetongue virus in experimental and field sera. J. Gen. Virol., 69, 2729-2740.
 
23.    MacLachlan N.J., Nunamaker R.A., Katz J.B., Sawyer M.M., Akita G.Y., Osburn B.I. & Taberchnick W.J. (1994). Detection of bluetongue virus in the blood of inoculated calves: comparison of virus isolation, PCR assay, and in vitro feeding of Culicoides variipennis. Arch. Virol., 136, 1-8.
 
24.    Martinez-Torrecuadrada J.L., Langeveld J.P., Venteo A., Sanz A., Dalsgaard K., Hamilton W.D., Meloen R.H. & Casal J.I. (1999). Antigenic profile of African horse sickness virus serotype 4 VP5 and identification of a neutralizing epitope shared with bluetongue virus and epizootic hemorrhagic disease virus. Virology, 257, 449-459.
 
25.    Martyn C.J., Gould A.R. & Eaton B.T. (1990). High level expression of the major core protein VP7 and the non-structural protein NS3 of bluetongue virus in yeast: use of expressed VP7 as a diagnostic, group-reactive antigen in a blocking ELISA. Virus Res., 18, 165-178.
 
26.    McColl K.A. & Gould A.R. (1991). Detection and characterisation of bluetongue virus using the polymerase chain reaction. Virus Res., 21, 19-34.
 
27.    Mecham J.O., Dean V.C., Wigington J.G. & Nunamaker R.A. (1990). Detection of bluetongue virus in Culicoides variipennis (Diptera: Ceratopognidae) by an antigen capture enzyme-linked immunosorbent assay. J. Med. Entomol., 27, 602-606.
 
28.    Naresh A. & Prasad G. (1995). Relative superiority of c-ELISA for detection of bluetongue virus antibodies. Indian J. Exp. Biol., 33, 880-882.
 
29.    Oldfield S., Adachi A., Urakawa T., Hirasawa T. & Roy P. (1990). Purification and characterization of the major group-specific core antigen VP7 of bluetongue virus synthesized by a recombinant baculovirus. J. Gen. Virol., 71, 2649-2656.
 
30.    Pritchard L.I., Gould A.R., Wilson W.C., Thompson L., Mertens P.P. & Wade-Evans A.M. (1995). Complete nucleotide sequence of RNA segment 3 of bluetongue virus serotype 2 (Ona-A). Phylogenetic analyses reveal the probable origin and relationship with other orbiviruses. Virus Res., 35, 247-261.
 
31.    Reddington J.J., Reddington G.M. & MacLachlan N.J. (1991). A competitive ELISA for detection of antibodies to the group antigen of bluetongue virus. J. Vet. Diagn. Invest., 3, 144-147.
 
32.    Roberts D.H., Lucus M.H. & Bell R.A. (1992). Animal and animal product importation and assessment of risk. In: Bluetongue, African Horsesickness and Related Orbiviruses, Walton T.E. & Osburn B.I., eds. CRC Press, Boca Raton, Florida, USA, 916-923.
 
33.    Roy P., Bishop D.H.L., LeBlois H. & Erasmus B.J. (1994). Long-lasting protection of sheep against bluetongue challenge after vaccination with virus-like particles: evidence for homologous and paretial heterologous protection. Vaccine, 12, 805-811.
 
34.    Shultz G. & Delay P.D. 1955). Losses in newborn lambs associated with bluetongue vaccination of pregnant ewes. J. Am. Vet. Med. Assoc., 128, 224-226.
 
35.    Standfast H.A., Dyce A.L. & Muller M.J. (1985). Vectors of BT in Australia. In: Bluetongue and Related Orbiviruses, Barber T.L. & Jochim M.M., eds. Alan R. Liss, New York, USA, 177-186.
 
36.    Thomas F.C. (1984). Comparison of some storage and isolation methods to recover bluetongue virus from bovine blood. Can. J. Comp. Med., 48, 108-110.
 
37.    Wade-Evans A.M., Mertens P.P.C. & Bostock C.J. (1990). Development of the polymerase chain reaction for the detection of bluetongue virus in tissue samples. J. Virol. Methods, 30, 15-24.
 
38.    Wilson W.C., Ma, H.C., Venter E.H., Van Djik A.A., Seal B.S. & Mecham J.O. (2000). Phylogenetic relationships of bluetongue viruses based on gene S7. Virus Res., 67, 141-151.
 
39.    Xu G., Wilson W., Mecham J., Murphy K., Zhou E.M. & Tabachnick W. (1997). VP7: an attachment protein of bluetongue virus for cellular receptors in Culicoides variipennis. J. Gen. Virol., 78, 1617-1623.
 

*
* *

NB: There are OIE Reference Laboratories for Bluetongue (please consult the OIE Web site at: http://www.oie.int/eng/OIE/organisation/en_LR.htm  ).

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 


 

Archive July and August 2005

Archive Jan 2005 - July 2005

Archive Oct 2004 - Dec 2004

Archive August 2004 -October

OTHER WARMWELL ARCHIVES(opens in new window)