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Journal of Virological Methods 98 (2001) 77 – 89 www.elsevier.com/locate/jviromet Bluetongue virus diagnosis of clinical cases by a duplex reverse transcription-PCR: a comparison with conventional methods Charalambos Billinis a, Maria Koumbati a, Vassiliki Spyrou a, Kyriaki Nomikou b, Olga Mangana b, Christos A. Panagiotidis c, Orestis Papadopoulos a,* a Laboratory of Microbiology and Infectious Diseases, Faculty of Veterinary Medicine, Aristotle Uni6ersity, GR-54006 Thessaloniki, Greece b Institute of Infectious and Parasitic Diseases, GR-15341 Agia Paraske6i, Attiki, Greece c Department of Pharmacy, Aristotle Uni6ersity, GR-54006 Thessaloniki, Greece Received 25 January 2001; received in revised form 21 June 2001; accepted 22 June 2001 Abstract A duplex reverse transcription polymerase chain reaction (RT-PCR) assay for the detection of bluetongue virus (BTV) in clinical samples was developed. This assay, which detects the highly conserved S10 region of BTV, was assessed for sensitivity and application as a rapid and dependable diagnostic tool by comparison with standard assays of virus detection, such as virus isolation in embryonated chicken eggs and cell culture. Simultaneous detection of BTV and host b-actin RNAs minimizes the possibility of false negative results. The sensitivity of the assay was found to be equal to five cell culture infectious dose (CCID50) units and its specificity was confirmed as no RT-PCR product was detected with RNAs from two closely related orbiviruses, i.e. epizootic haemorrhagic disease virus (serotypes 1, 2 and 318) and African horse sickness virus, serotype 9, or RNAs from uninfected BHK-21 cells and blood samples from uninfected sheep or goats. In this study, 36 blood samples from naturally infected mixed flocks of sheep and goats were examined. Seventeen animals were identified as BTV-positive by RT-PCR, whereas only 13 were found positive by virus isolation in embryonated chicken eggs and nine by cell culture assays. These results indicate that the duplex RT-PCR could be a useful technique for monitoring BTV infection in the field. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Bluetongue virus; Reverse transcription duplex polymerase chain reaction; Bluetongue diagnosis; Clinical cases 1. Introduction * Corresponding author. Tel.: + 30-31-999951; fax: +3031-999959. E-mail address: [email protected] (O. Papadopoulos). Bluetongue is an arthropod-transmitted disease of wild and domestic ruminants. It is a virusborne disease that is caused by the bluetongue 0166-0934/01/$ - see front matter © 2001 Elsevier Science B.V. All rights reserved. PII: S 0 1 6 6 - 0 9 3 4 ( 0 1 ) 0 0 3 6 0 - 3 78 C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 virus (BTV), which is the prototype species of the genus Orbi6irus in the family Reo6iridae (MacLachlan, 1994). There are at least 24 serotypes of BTV worldwide (Davies et al., 1992), three of which (4, 9 and 16) have been isolated in Greece (K. Nomikou et al., unpublished data). BTV infection occurs throughout temperate and tropical regions of the world, and infection is dependent on the presence of competent vector midges of Culicoides spp. (Gibbs and Greiner, 1994; Mellor, 1990). These insects become infected persistently with BTV and are infectious to ruminants on which they feed after an extrinsic incubation period of 10– 14 days (Tabachnick, 1996). Bluetongue is an International Office of Epizootics List A disease (Alexander et al., 1994; OIE, 2000) described as an economically devastating affliction of sheep (Alexander et al., 1994). Indeed, the vast economic effects of BTV infection in many parts of the world, due to the high morbidity and mortality rates in the affected animals have been established (Abu Elzein et al., 1992; Eisa et al., 1980; Hourrigan and Klingsporn, 1975; Quist et al., 1997; Barnard et al., 1998; Erasmus, 1975; Gee, 1975; Geering, 1975; Kvasnicka, 1985; Metcalf et al., 1980; Mulhern et al., 1985; Mulhern, 1985; Murphy et al., 1985; Tabachnick, 1996). Early detection of infected animals could reduce drastically the consequences of the disease by reducing the virus pool and by containing the dissemination of the disease through export of potentially infectious animals. The latter can be a very significant problem as infected animals, especially cattle, present viremia for long periods (up to 100 days) with no clinical signs (Erasmus, 1990; Roy, 1996). BTV routine diagnosis is based primarily on serological methods that detect virus-specific antibodies in serum (Pearson et al., 1992). A number of other procedures are also used currently to detect BTV from blood or tissues of infected animals. These include direct inoculation of cultured mammalian or insect cells, or intravenous inoculation into 10– 12 day embryonated chicken eggs, followed by one passage in insect cell culture and up to three passages in mammalian cell cultures (Foster and Leudke, 1968; Gard and Kirkland, 1993; Wechsler and McHolland, 1988). In particular, the inoculation of embryonated chicken eggs and passaging through cell culture is the generally accepted method for testing of animals for export and other regulatory purposes. This is, however, a laborious and time-consuming protocol that may take up to 5 weeks for completion. Consequently, alternative methods of virus detection have been sought. These include antigen capture enzyme linked immunosorbent assay (ELISA), dot immunobinding assay (DIA), immunoelectron microscopy and polymerase chain reaction (PCR) (Hawkes et al., 2000; Katz et al., 1993; McColl and Gould, 1991; Mecham, 1993; Mecham et al., 1990; Mecham and Nunamaker, 1994; Nunamaker et al., 1997a,b; Shad et al., 1997). The use of antigen capture ELISA for the detection of BTV in the blood of infected ruminants has either been unsuccessful (Mecham, 1993), has detected antigen only in animals with high viremias (Stanislawek et al., 1996), or was not consistent enough to allow for the reliable diagnosis of BTV (Hawkes et al., 2000). A major problem in the diagnosis of BTV infection by immunological methods is also the cross-reactivity with proteins from other orbiviruses (Lunt et al., 1988), although this may be circumvented by the use of c-ELISA (Afshar et al., 1989, 1987). To avoid these problems, PCR-based assays were developed and evaluated for the detection of BTV serotypes based on nucleotide sequences of different genome segments (Akita et al., 1992, 1993; Aradaib et al., 1998; Dangler et al., 1990; McColl and Gould, 1991; Parsonson and McColl, 1995; Shad et al., 1997; Wade-Evans et al., 1990; Wilson, 1999; Wilson and Chase, 1993). Although these assays were found to detect BTV in BTV-infected cell cultures and in infected experimentally ruminants, their usefulness and dependability under field conditions have not been demonstrated. The purpose of this study was to develop a sensitive and dependable PCR assay for BTV detection in clinical (field) specimens (blood samples) from sheep and goats infected naturally and evaluate its sensitivity in comparison with conventional methods such as embryonated chicken eggs inoculation and cell culture assays. C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 2. Materials and methods 2.1. Cells and 6iruses Purified preparations of bluetongue virus (BTV) serotypes 1, 2, 3, 4, 9 and 16 were obtained from the Onderstepoort Veterinary Institute, South Africa. Greek isolates of the BTV serotypes 4, 9 and 16, which represent different topotypes than those obtained from South Africa, were also used. The epizootic haemorrhagic disease virus serotypes 1, 2 and 318, and the African horse sickness virus serotype 9, were obtained from the Institute of Animal Health, Pirbright Laboratory, UK. When necessary, the viruses were propagated on confluent BHK-21 cell monolayers, as described previously (Koumbati et al., 1999). In brief, BHK-21 cell monolayers were grown in BHK-21 Glasgow medium (Gibco BRL) supplemented with 10% fetal calf serum (FCS). The cells were infected with 1 ml of treated blood cells and the culture was incubated at 37 °C until the cytopathic effects became apparent. At that time, the infected cell culture fluid was frozen and kept at − 80 °C. 2.2. Clinical samples Blood samples were collected from 24 sheep and 12 goats, from naturally infected mixed flocks, in EDTA-containing tubes (VACUTAINER™), during the bluetongue, serotype 9, epizootic on the island of Rhodes, Greece in 1998. The majority of the sheep in those flocks had shown bluetongue clinical signs during the month prior to sampling, whereas the goats appeared to be healthy. 2.3. Virus isolation in embryonated chicken eggs, cell culture assays and 6irus identification Virus isolation and identification were processed as described previously (Koumbati et al., 1999). Briefly, 0.1 ml of washed, packed and ultrasonic-treated blood cells was inoculated intravenously into each of six 12-day-old embryonated chicken eggs. BTV-positive samples caused embryo haemorrhage that usually resulted in em- 79 bryo death, in most cases. Dead embryos were collected between 2 and 7 days post-inoculation for virus identification. BHK-21 cell monolayers were inoculated with 1.0 ml of treated sample and virus adsorption was allowed to take place for 1 h and 30 min at 37 °C. The monolayers were then washed twice with phosphate-buffered saline (PBS) and BHK-21 Glasgow medium (Gibco BRL, Grand Island, NY) was added. Monolayers were examined daily for BTV cytopathic effects for a maximum of 10 days. If no obvious cytopathic effects were evident for 10 days, the cells were removed and passaged into a new monolayer. A sample was characterized as negative when it had shown no cytopathic effects after three blind passages. For virus identification, Vero cells were cultivated in 8-chambered slides (Lab-tek) and inoculated with either 0.2 ml of infected cell culture medium or 0.2 ml of dispersed heart supernatant from infected chick embryos. Positive and negative controls were included. Forty-eight hours post-inoculation, the cells were fixed in situ with acetone at 4 °C for 15 min and at − 70 °C for another 30 min (Jochim et al., 1974). Virus identification was carried out by indirect immunofluorescence using the Pirbright monoclonal antibody 3-17-A3 (Anderson, 1984). The presence of intracytoplasmic fluorescent inclusions, which are characteristic of bluetongue virus, was observed in the positive cultures. 2.4. RNA extraction procedures The RNA extractions were carried using the single-step method described by Chomczynski and Sacchi (Chomczynski and Sacchi, 1987) with the TRIzol™ LS reagent (Gibco BRL) available commercially. The precautions recommended (Sambrook et al., 1989) to avoid contamination with ribonucleases were observed. 2.4.1. RNA extraction from blood samples and chicken embryos About 0.25 ml of total blood sample (diluted 1:1 with water), or 100 mg of heart tissue from chicken embryos was homogenized in 0.75 ml TRIzol™ LS reagent (Gibco BRL). Following a 5 80 C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 min incubation, 0.2 ml of chloroform was added to each sample and the solution was shaken by hand and then incubated at room temperature for approximately 10 min. The samples were centrifuged at 15,000× g for 15 min at 4 °C and the RNA-containing aqueous phase was transferred to a fresh tube. Following the addition of 0.5 ml ice-cold isopropanol and a 30 min incubation at − 20 °C, the RNAs were precipitated by centrifugation at 15,000×g for 15 min at 4 °C. The resulting RNA pellets were washed with 75% ethanol before air-drying, and each RNA sample was dissolved in 50 ml diethyl pyrocarbonate treated H2O by incubating 10 min at 55 °C. 2.4.2. RNA extraction from infected cell cultures Frozen, infected cell culture fluid, prepared as described above, was thawed and centrifuged at 6000×g for 5 min. The virus was pelleted from 10 ml of the resulting supernatant by centrifugation at 25,000× g for 16 h at 4 °C. Subsequently, the virus pellet was dissolved in 0.75 ml TRIzol™ LS reagent (Gibco BRL). The mixture of virus pellet and the monophasic solution of phenol and guanidine isothiocyanate was incubated for 5 min at room temperature to permit the complete dissociation of nucleoprotein complexes. The RNAs were then isolated as described above. 2.5. Oligonucleotide primers The oligonucleotide primers CB1 and CB2 (Table 1) were designed based on published sequence (Pierce et al., 1998) of the BTV10 segment 10, (EMBL accession no. AF044381). All oligonucleotides were synthesized commercially (MWG, Germany). The control primers BA1 (GAGAAGCTGTGCTACGTCCGC) and BA2 (CCAGACACGCACTGTGTTGGC), whose sequences are based on the conserved bovine b-actin gene, were included in our duplex RT-PCR assay since they provide a positive control for sample quality, the integrity of the extracted RNA and the efficiency of the reactions. BA1 and BA2 have been shown to work well on RNA from all domestic species such as goat, pig, cattle and sheep (Reid et al., 1998). 2.6. Re6erse transcription (RT) and polymerase chain reaction (PCR) amplification Reverse-transcription (RT) reactions were performed by mixing 7.6 ml of RNA extraction product with 12.4 ml of an RT premix to obtain a final concentration of 1× first strand buffer (Gibco BRL), 0.5 mM dNTP mix (Gibco BRL), 10 mM DTT (Gibco BRL), 100 U of Moloney murine leukaemia virus (MMTV) reverse transcriptase (Gibco BRL), and 3.5 pmol/ml of random hexamers (Pharmacia Biotech) per reaction. Following preparation of the RT reaction mixtures on ice, reverse transcription reactions were carried out at 37 °C for 30 min, they were terminated by heating for 5 min at 95 °C. PCR amplifications were carried out observing the guidelines described by Kwok and Higuchi (Kwok and Higuchi, 1989). Each PCR reaction mixture contained 2 ml of RT product, 1× PCR buffer (Gibco BRL), 1 mM MgCl2 (Gibco BRL), 0.2 mM of a dNTP mix (Gibco BRL), 0.15 U of Taq Polymerase (Gibco BRL), and 30 pmol of each primer. The reaction volume was adjusted to 20 ml with DEPC treated H2O. The PCR mixtures were prepared on ice and the reactions were initiated by heating for 3 min at 94 °C, followed by 30 cycles of: 1 min at 94 °C, 2 min at 52 °C and 1 min at 72 °C . The mixtures were then brought to 72 °C for 5 min and then held at 4 °C. The PCR conditions remained the same for both simple and duplex PCR reactions, with the CB1/CB2 and BA1/BA2 primer pairs. Following the amplification, 10 ml of each RT-PCR product was analyzed by electrophoresis on a 2.5% agarose gel and stained with ethidium bromide (0.5 mg/ml). A 100 bp DNA ladder (Gibco, BRL) was analyzed on the same gel to serve as a size marker. The expected sizes of the RT-PCR products were 792 bp for the BTV S10 amplimer and 275 bp for the b-actin amplimer. 2.7. Specificity and sensiti6ity of the RT-PCR procedure The specificity of the assay was determined by applying the above described RT-PCR protocol to RNAs prepared from stocks of either the BTV C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 serotypes 1, 2, 3, 4, 9 and 16, or the closely related orbiviruses epizootic haemorrhagic disease virus, serotypes 1, 2 and 318, and the African horse sickness virus serotype 9, or to RNAs extracted either from cultures of uninfected BHK-21 cells or blood samples from uninfected sheep or goats. The sensitivity of the assay was evaluated by 81 applying it to virus detection using logarithmic dilutions of a titrated BTV, serotype 4, suspension in sterile 0.04 M phosphate buffer, pH 7.2. Each virus dilution was successively subjected to the RNA isolation, reverse transcription and PCR amplification steps, as outlined previously, and the products were analyzed on a 2.5% agarose gel. Table 1 Homology of the BTV primers used to S10 region sequences from field and laboratory strains of bluetongue virus BTV strain/accession numbera CB1 primer (bp 21–41) TGCTATCCGGGCTGATCCAAAb CB2 primer (bp 813–796) TAGCGCCGCGTACCCTCCb VAC10/AF044376 10B80Z/AF044379 10B81X/AF044382 10B81U/AF044381 10B9OZ/AF044384 10O80Z/AF044380 10O90H/AF044385 VAC11/AF044377 11B80Z/AF044702 11B81P/AF044383 11C81Z/AF044703 11O79X/AF044386 11O81X/AF044704 13B80Z/AF044702 13B81K/AF044712 13B89Z/AF044710 13O79Z/AF044713 VAC17/AF044378 17B80Z/AF044705 17B81Y/AF044707 17B90Z/AF044708 17O79Y/AF044706 17O90Y/AF044709 BTV4/AF135226 BTV9 BTV16/AF135229 Austral BTV/D00253 BTV1/AF135223 BTV2/AF135230 BTV3/AF135225 BTV12/AF135227 BTV15/AF135228 + + + + + + + + + + + + + + + + + + + + + + + + (+) + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + (+)c (+) (+) + (+) (+) (+) ?c ? a The primer sequences were compared with all available BTV sequences. The accession number is given next to the virus designation. b Numbers indicate the location of primer sequences within BTV S10 region, and are based on the BTV10 sequence (10B81U) (Pierce et al., 1998). c Plus in parenthesis (+) means that the RT-PCR was found to work well with the isolates tested, despite absence of match between primer sequences and the published sequence of the particular serotype, and (?) that the primer sequence is not present in the published BTV sequence and no experimental verification was done due to unavailability of the particular serotype. 82 C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 Fig. 1. Specificity of the RT-PCR method with the CB1 and CB2 primers. (A) Total RNAs were isolated either from uninfected BHK-21 cells (lanes 2 and 3) or from purified virus preparations of bluetongue virus serotypes 4 (BTV4, lane 4), 9 (BTV9, lane 5) and 16 (BTV16, lane 6), or epizootic haemorrhagic disease virus serotype 1(EHDV1, lane 7), 2 (EHDV2, lane 8) and 318 (EHDV318, lane 9), and African horse sickness virus serotype 9 (AHSV9, lane 10). All BTV viruses used in this panel were isolated in Greece. (B) Total RNAs were isolated either from control uninfected goat blood (Blood, lane 2), or from purified virus preparations of BTV serotypes 1 (BTV1, lane 3), 2 (BTV2, lane 4), 3 (BTV3, lane 5), 4 (BTV4, lane 6), 9 (BTV9, lane 7) and 16 (BTV16, lane 8). All BTV viruses analyzed in this panel were obtained from the Onderstepoort Veterinary Institute, South Africa, and they represent different topotypes than those analyzed in panel A. The RTPCR reactions were carried out, with the CB1/CB2 primer pair, as described in Section 2 and the reaction products were analyzed on a 2.5% agarose gel. The size markers used in both panels (M, lane 1) were a 100-bp DNA ladder (Gibco-BRL). Fig. 1. (Continued) 3. Results 3.1. Specific bluetongue 6irus RNA detection using re6erse transcription PCR with the CB1 and CB2 primers The CB1 and CB2 primers were designed based on the sequence of the highly conserved S10 region, coding for the nonstructural proteins NS3 and NS3A, of the BTV10 strain sequence (10B81U, Accession no. AF044381) (Pierce et al., 1998). The CB1 primer hybridizes to the 5% end of the NS3/NS3A gene (bases 21– 41) (Pierce et al., 1998), and CB2 hybridizes close to the edge of the noncoding 3% end (bases 813–796) of the NS3/ NS3A gene. Care was taken, in primer design, to avoid primer internal stability problems, dimer formation, or homologies with RNAs from other orbiviruses, other regions of the BTV RNA, or cellular RNAs. As shown in Table 1, the sequences of both the CB1 and the CB2 primers were found to exist in the majority of the published BTV S10 region (NS3 and NS3A gene) sequences. The apparent absence of CB2 sequences in the published sequences of some BTV serotypes might be due to incomplete sequencing at the far 3% end of the S10 RNA region. This notion is based on the finding that the CB1/CB2 primers were found to amplify a fragment of the appropriate size (792 bp) from at least five BTV serotypes (1, 2, 3, 4 and 16) whose published S10 region sequences lack homology with the CB2 primer (Fig. 1). As stated above, the RT-PCR method described here was found to detect BTV RNA isolated from purified preparations of the serotypes 1, 2, 3, 4, 9 and 16 (Fig. 1). BTV RNAs for serotypes 4, 9 and 16 isolated from different geographical regions (topotypes), i.e. Greece (Fig. 1A) and South Africa (Fig. 1B) were detected equally well by our method. In addition, two Greek isolates from different geographical regions (topotypes) of bluetongue serotype 4 were also C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 83 detected (data not shown). In contrast, no PCR product was detected when total RNA, isolated either from control uninfected BHK-21 cells (Fig. 1A) or blood samples from uninfected sheep or goats (Fig. 1B), were used. The specificity of the method was further demonstrated by the absence of amplification products when RNAs from three serotypes (1, 2 and 318) of the closely related epizootic haemorrhagic disease virus or the African horse sickness virus, serotype 9 were used as templates (Fig. 1A). The above data argue against the possibility that false positive results will be obtained with the RT-PCR method, due to the presence of either cellular RNAs or RNAs of related viruses. 3.2. Sensiti6ity of the RT-PCR procedure The sensitivity of the assay was evaluated by using serial dilutions of a titrated BTV4 suspension. The virus had been titrated on BHK-21 cells and the titer is expressed as cell culture infectious dose (CCID50) units. It was found that our RTPCR assay could detect amounts of BTV as low as 5 CCID50 (Fig. 2). Fig. 2. Sensitivity of the RT-PCR method. The sensitivity of the assay was evaluated by performing RT-PCR reactions on RNAs prepared from serial dilutions of a BTV4 suspension that had been titrated in BHK-21 cells. The titration units are expressed as CCID50, and the virus load equivalent used in each RT-PCR reaction is 109 (lane 2), 107 (lane 3), 105 (lane 4), 103 (lane 5), 100 (lane 6), 10 (lane 7), 5 (lane 8), 2.5 (lane 9) and 0 (lane 10). The reaction products were analyzed on a 2.5% agarose gel and the size markers used (M, lane 1) were a 100-bp DNA ladder (Gibco-BRL). 3.3. Application of the RT-PCR assay in bluetongue 6irus RNA detection to field specimens To determine the actual value of the above-described RT-PCR assay we applied it to BTV RNA detection using blood samples collected from 24 sheep and 12 goats, from naturally infected mixed flocks, during a bluetongue epizootic on the island of Rhodes, Greece in 1998. The results of this assay were compared with the results of two other standard tests used in routine BTV diagnosis, namely cytopathic effects on BHK-21 cells in culture and inoculation into 12day embryonated chicken eggs. Virus detection in the field diagnostic samples using inoculation of embryonated chicken eggs indicated that 13 out of the total 36 samples were positive for the presence of the virus (Table 2). Embryonic death was noted, in the positive samples, between days 2 and 7 post inoculation and the number of dead embryos varied from one to five out of six eggs that had been inoculated with each field sample. All positive blood samples were derived from sheep, whereas all goat samples were found negative. Further analysis, by indirect immunofluorescence, indicated that all the positive samples were indeed positive for BTV antigens (data not shown). Therefore, 13 out of the 36 samples were confirmed as positive for the presence of the bluetongue virus using embryonated chicken eggs inoculation (Table 2). BTV detection by measuring virus cytopathic effects on cultured BHK-21 cell monolayers was less efficient, since only nine positive samples were detected (Table 2). These nine positive samples, were also found positive by indirect immunofluorescence with an anti-BTV monoclonal antibody, and were a subset of the 13 samples found positive by the embryonated chicken eggs assay (Table 2). C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 84 PCR (Table 2). Inclusion of the b-actin and BTV primers in the same RT-PCR reactions provides an additional control against possible false negative results. Indeed, the presence of the 275 bp b-actin RNA amplification product, using the conserved BA1 and BA2 b-actin primers in the In contrast, the duplex RT-PCR assay was far more sensitive since it detected 17 positive samples (Fig. 3; Table 2). It is noted that the 13 samples found positive by the embryonated chicken eggs inoculation and cell culture assays were among those identified as positive by RT- Table 2 Comparison of the RT-PCR with two conventional BTV diagnosis assays Number S1 S2 S3 S4 S5 S6 S7 S8 S9 S10 S11 S12 S13 S14 S15 S16 S17 S18 S19 S20 S21 S22 S23 S24 G1 G2 G3 G4 G5 G6 G7 G8 G9 G10 G11 G12 RT-PCR + + + + + + + + + + + − − + − − + + + + + − − − − − − − − − − − − − ECEs + + + − − + + + + + + − − + − − − − + + − + − − − − − − − − − − − − − − Cell culture + + + − − + + + + + − − + − − − − − − − − − − − − − − − − − − − − − − RT-PCRa ECEs Cell culture + + + ND ND + + + + + + ND ND + ND ND ND ND + + ND + ND ND ND ND ND ND ND ND ND ND ND ND ND ND + + + + + + + + + + + − − + − − + + + + − + − − − − − − − − − − − − − − The conventional BTV diagnosis assays were (i) inoculation of 12-day embryonated chicken eggs (ECEs) and (ii) inoculation of BHK-21 cell cultures (cell culture). Thirty six clinical samples (blood) were tested. (+)= BTV-positive sample, (−)=negative sample, (ND) = not done. a RT-PCR was performed on RNAs isolated either from positive ECEs (dead embryos) or from cell cultures inoculated with clinical samples. C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 85 Fig. 3. Application of the duplex RT-PCR method in field diagnosis. Total RNAs were isolated either from BHK-21 cells infected with BTV9, as control, or from 24 blood samples from sheep (S1 – S24) and 12 from goats (G1 – G12). RT-PCR reactions were performed and the products (792 bp for the BTV product and 275 bp for that of b-actin, indicated by arrows) were analyzed on 2.5% agarose gels (A and B) and a 100-bp DNA ladder was used as size markers (M). same reactions (Fig. 3), practically nullifies the possibility that the lack of amplification in the negative samples is due to low quality RNA or other technical problems. The increased specificity of the RT-PCR method became more obvious when it was applied on RNAs obtained from BHK-21 cell cultures that had been inoculated with the clinical samples. Positive RT-PCR results, for the presence of BTV, were obtained not only for the nine samples showing cytopathic effects, but for an additional eight samples (Table 2). All of the above samples had been identified as positive by RT-PCR on RNAs isolated from the blood samples (Table 2). RT-PCR assays on RNAs from dead chicken embryos further verified the presence of BTV in these samples (Table 2). 4. Discussion Monitoring and control of BTV infection in cattle, sheep and goats remains a top priority in BTV-endemic and epidemic countries interested in exporting livestock free of this disease or in restricting the introduction of new serotypes into already existing endemic populations (Morley, 1993; Roberts et al., 1993). While serological assays are sensitive and easy to use they only provide evidence indicating earlier animal expo- 86 C. Billinis et al. / Journal of Virological Methods 98 (2001) 77–89 sure to BTV but not necessarily an ongoing infection. Earlier work on cattle and sheep has provided evidence that seropositivity does not correlate with circulating BTV RNA or the presence of infectious virus in the blood of the seropositive animals (Singer et al., 1998). A number of procedures have been developed to detect the presence of BTV antigens or nucleic acids. PCR is a powerful tool in the field of diagnostic medicine and it has been used successfully in identifying several infectious diseases of veterinary importance (Belak and Ballagi-Pordany, 1993). By using PCR-based techniques, researchers may circumvent problems such as serologic cross-reactivity among related orbiviruses (Ristow et al., 1988). Several RT-PCR assays have been described for the detection of BTV (Akita et al., 1992, 1993; Aradaib et al., 1998; Dangler et al., 1990; McColl and Gould, 1991; Parsonson and McColl, 1995; Shad et al., 1997; Wade-Evans et al., 1990; Wilson, 1999; Wilson and Chase, 1993). However, despite the fact that these assays have been tested using virusinfected cultures or experimentally infected animals, their usefulness in clinical diagnosis remains mostly unproven due to a lack of comparative evaluation with standard methods using field samples. The above is underscored by a recent report (Tiwari et al., 2000) which shows that an RT-PCR method that works well with purified BTV RNA fails to detect the virus RNA in clinical samples, unless a second nested-PCR step is included. In an earlier report of a BTV PCR assay (McColl and Gould, 1991), virus nucleic acid was only detected in the leukocyte fraction of blood from infected animals. The aim of the present study, however, was to develop a simpler RT-PCR assay, that could detect the virus in total unfractionated blood. It was also imperative that the assay should detect all known BTV serotypes but no other related viruses (such as epizootic haemorrhagic disease virus), a problem that has plagued many serological assays for this virus. To achieve this primers were designed that could amplify segment 10 of the BTV10 genome, a region that codes for the non structural proteins NS3 and NS3A. Selection of the particular genome segment, as a target for a diagnostic RT-PCR assay was based on the observation that it is one of the most highly conserved segments of BTV genome (Hwang et al., 1992; Pierce et al., 1998). Additional care was taken, in primer design, to avoid homologies between primer sequences and sequences of known virus or mammalian genes. As a result we obtained a set of primers (CB1 and CB2) that could amplify specifically the BTV genome without any obvious background when either uninfected BHK-21 cells (Fig. 1A) or blood samples from uninfected sheep or goats (Fig. 1B) were used. Additionally, no cross-reactivity was observed with two other closely related orbiviruses, such as the epizootic haemorrhagic disease virus and the African horse sickness virus (Fig. 1A). These data minimize the possibility that false positive results will be obtained due to amplification of either cellular or related orbivirus RNAs. Furthermore, the assay proved to be sensitive, as it could detect quantities of BTV as low as 5 CCID50 (Fig. 2). There is always a possibility that a PCR-based assay will yield false negative results due to low quality of some clinical samples, technical problems or experimental error. Therefore, it is crucial to include an internal control for the identification of such false negatives. The presence of the b-actin primers (BA1 and BA2) in our duplex RT-PCR represents such an internal control, since the absence of a b-actin amplification product would indicate that the particular sample is a false negative. The conditions were optimized so that the double BTV/b-actin PCR reactions could take place in the same tubes without lose of specificity or sensitivity (data not shown). The results presented in Fig. 3, where the method was applied in clinical diagnosis using 36 whole blood samples, argue that this method does achieve the goals set at the beginning of the work. Furthermore, the diagnostic sensitivity of the RTPCR assay is greater than the virus inoculation in embryonated chicken eggs or assay of cytopathic effects on cultured cells (Table 2). 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