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Appendix 42 Prospects for improved laboratory diagnosis of FMD using real-time RT-PCR Nigel Ferris*, Scott Reid, Donald King, Geoff Hutchings and Andrew Shaw Pirbright Laboratory, Institute for Animal Health, Ash Road, Woking, Surrey GU24 0NF, UK Abstract: Definitive diagnosis of FMD requires the detection of virus, antigen or genome in clinical material. The aim was to evaluate the performance of an automated real-time RT-PCR procedure for this purpose. Vesicular epithelia from eighteen countries were examined by ELISA, VI and RT-PCR. Retrospective analysis by RT-PCR was also performed on available material of two sample subsets collected from ‘confirmed’ cases during the 2001 UK FMD outbreak : firstly, samples which were negative by both ELISA and VI and secondly, others which were negative by ELISA on epithelial suspension but positive by VI. There was broad agreement between RT-PCR and VI for 79% and VI/ELISA combined for 82% of the overseas epithelial samples tested. There were no false negative results obtained with RT-PCR since all samples assigned negative by RT-PCR were also negative by VI/ELISA. However, the RT-PCR was able to detect FMDV in an additional 18% of the samples tested. Additionally, there was good agreement between the RT-PCR and ELISA/VI for the UK outbreak samples save for a group of related virus isolates from Wales. These viruses had evidently evolved during the epidemic and had a nucleotide substitution in the RT-PCR probe site, which prevented detection by RT-PCR using the routine diagnostic probe. The ELISA and VI are deficient for specimens of poor quality where concentrations of infectious FMDV may either be low or absent. The features that influence sample quality appear to be less important for the RT-PCR as it can detect a small fragment of FMDV genomic RNA, not just live virus. Real-time RT-PCR provides an extremely sensitive and rapid procedure that contributes to improved laboratory diagnosis of FMD. However, the failure to detect the mutant FMDVs from the UK 2001 epidemic illustrates the importance of constant monitoring of representative field FMDV strains by nucleotide sequencing to ensure that the primers/probe set selected for the diagnostic RT-PCR is fit for purpose. Introduction: Control of outbreaks of foot-and-mouth disease (FMD) is dependent upon a system of monitoring and early detection, which requires basic familiarity with clinical signs and the ability to characterise the strain of virus responsible by laboratory tests. Definitive diagnosis of FMD requires the detection of virus, antigen or genome in clinical material. Ideally, the sample of choice should be vesicular epithelium from clinically affected animals since, during the acute stage of the disease, it is rich in virus. The World Reference Laboratory (WRL) for FMD typically receives between 400 and 700 samples annually from overseas countries (Ferris and Donaldson, 1992; Table I), including other sample types besides epithelia, e.g. epithelial suspensions, cell culture antigens, blood, throat swab (probang) and milk samples. For almost twenty years, the WRL for FMD has used an indirect, sandwich enzyme-linked immunosorbent assay (ELISA) (Roeder and Le Blanc Smith, 1987; Ferris and Dawson, 1988; OIE, 2004) to identify FMDV. However, the ELISA is not 100% sensitive. Consequently, suspensions of each specimen are also propagated in sensitive cell cultures (Ferris and Dawson, 1988) and the specificity of any isolated virus confirmed by the ELISA. Whilst such virus isolation (VI) methods are highly sensitive, they require four days before a negative result can be concluded (and reported as ‘no virus detected’ [NVD]). It is evident from Table I that FMDV antigen cannot be detected by ELISA and VI in approximately half the submitted samples. This has given cause for concern as to the efficiency of sample collection and dispatch and also with respect to the adequacy of the laboratory test procedures employed for their examination. In emergencies, speed of diagnosis (clinical and laboratory confirmation) is of paramount importance to control spread and eradicate disease. Approximately, 90% of positive epithelial samples received during the 2001 UK FMD outbreak were so defined by the antigen ELISA on prepared suspensions, the remainder being serotyped after amplification and isolation of virus following cell culture passage. Negatives could only be classified following double passage in cell culture, which took 4 to 5 days. Consequently, the introduction by the UK Government towards the end of March of a 24/48 hour culling policy (all animals to be slaughtered on infected premises within 24 hours of diagnosis and those on neighbouring premises within 48 hours) meant that confirmation of disease was subsequently made by clinical judgement alone. This policy, although playing a critical role in controlling disease, caused huge controversy and provoked much debate on the likelihood of many animals being slaughtered unnecessarily, fuelled by the finding that neither virus nor antibody could be detected in samples received from many of the confirmed cases. 261 Recently, the development of a real-time reverse transcription polymerase chain reaction (RT-PCR) procedure has provided an additional tool which can be used for FMD diagnosis (Reid et al., 2002). Furthermore, this real-time RT-PCR method can be automated allowing increased throughput of samples with fewer user-dependent steps (Reid et al., 2003). The authors have compared the performance of a fully automated real-time RT-PCR (Reid et al., 2003) with VI and ELISA for the detection of FMDV on the majority of epithelium samples received at the WRL for FMD from overseas during a recent eighteen-month period (August 2002 until January 2004) and on two subsets from confirmed cases from the recent UK outbreak : firstly, samples which were negative by both ELISA and VI and secondly, others which were negative by ELISA on epithelial suspension but positive by VI. Materials and Methods: Three hundred and thirty four samples of vesicular epithelium were received from eighteen countries during the period of the study and were examined by ELISA, VI and RT-PCR (Table II). Upon receipt, the pH of the transport buffer containing epithelial tissue was estimated using phenol red indicator. A suspension of the epithelium (ES) was made in 0.04 M phosphate buffer (ideally this should be done using a 10% concentration (w/v), but during this study a lower concentration was frequently used due to paucity of material). A 1.4 ml aliquot was taken for the antigen detection ELISA (Ferris and Dawson, 1988; OIE, 2004) whilst 0.2 ml aliquots were used to inoculate cell cultures of primary bovine thyroid (Snowdon, 1966) and IB-RS-2 cells (De Castro, 1964) (five tubes of each cell type per specimen). Additionally, 0.2 ml of ES were added to a 1 ml aliquot of Trizol solution and stored at 80°C until assay. Real-time RT-PCR was usually performed on the ES within one to two days of preparation, with a diagnostic result typically being obtained within a single working day. Three hundred and eighty samples from 331 separate premises (‘confirmed’cases) in the UK, which had been found to be negative by both ELISA and VI in 2001, and 199 samples from a further 188 separate premises, which had been found to be positive by VI but negative by ELISA on epithelial suspension, were re-evaluated by RT-PCR. In the majority of cases, the original epithelial suspension (which had been stored at -80oC) was used but new suspensions were prepared for others, while aliquots of the transport buffer were examined in a few cases (in the absence of both stored suspension and submitted epithelium). The real-time RT-PCR assay used in this study has been described elsewhere (Reid et al., 2003). Briefly, total nucleic acid was extracted from the solution of Trizol/ES using a fully automated robot system. This robot was then also used to pipette viral nucleic acid into a reverse transcription mix for reverse transcription and complementary deoxyribonucleic acid into PCR reaction mix (including an oligonucleotide primers/probe set targeting the internal ribosomal entry site of the FMDV genome). Thermal cycling and concurrent fluorescence detection were performed and a cycle detection threshold (i.e. the cycle at which a target sequence is detected [CT]) was recorded for each test sample (Oleksiewicz, Donaldson and Sorensen, 2001). Sequencing of viruses was performed as described by Knowles and Samuel (2003). Results: Samples submitted to the WRL for FMD from overseas The results achieved by ELISA, VI in cell culture and RT-PCR are summarised in Table II for comparison of the performance of the three assays. FMDV was detected in 195 samples by VI (58.4%), in 125 by antigen ELISA (37.4%) and in 204 samples by VI/ELISA combined (61.1%). These viruses represented four out of the seven FMDV serotypes (O, A, SAT 2 and Asia 1), although the majority (68.6%) were of serotype O. There was broad agreement between RT-PCR and VI for 265/334 (79.3%) and VI/ELISA combined for 274 (82%) of the epithelial samples tested when a cut-off CT value of ‘<39’ was used to assign the samples as either FMDV positive or negative (results not shown). In general, VI positive samples corresponded to a real-time RT-PCR CT value of <35, with only eight VI positive samples producing a CT value in the range ≥35-<39. Furthermore, there were no false negative results obtained with RTPCR since all samples assigned negative by RT-PCR were also negative by VI/ELISA. Of particular interest was the finding that RT-PCR was able to detect FMDV in an additional sixty (18%) of the samples tested. Fresh ribonucleic acid (RNA) was prepared from these epithelial suspensions and the RT-PCR was repeated with similar results confirming the presence of FMDV genome in these samples. In addition, there were seventeen ‘suspect positive’ samples that generated CT values in the range ≥39 to <50. Repeat procedures were also performed for these samples with similar results, suggesting that FMDV was also present in these samples although the FMDV copy number was lower than the diagnostic threshold. The majority (96.9%) of VI positive samples were detected on the first passage in cell cultures (within two days). However, in order to ensure detection of infectious FMDV 262 and to define NVD samples, a second cell culture passage was required (in total, up to four days). Indeed six samples required this second passage in cell culture before the presence of FMDV was demonstrated. Calculations of the relative sensitivity and specificity of the RT-PCR at unit incremental increases in CT value in comparison to either VI alone or to VI/ELISA combined are recorded in Table III. It shows that 100% relative sensitivity was achieved by the RT-PCR at a CT value of <39, which justifies using this as the ideal diagnostic cut-off for the RT-PCR, since it is the lowest CT value that achieves this. 2001 UK FMD outbreak samples – ELISA/VI negative It can be seen from Table IV that 367 samples produced a CT value of 50 (FMDV genome not detected by RT-PCR). However, 7 sample suspensions yielded CT values of <35, and 2 in each of the ranges >35-<40, >40-<45 and >45-50. Subsequent to these findings, attempts were made to isolate virus from available epithelial suspensions, with success (i.e. positive type O FMDV) in 3 out 5 samples from the first category (paucity of material precluded examination of 2 samples) and 0, 1 and 0 samples from the other categories, respectively. 2001 UK FMD outbreak samples – ELISA negative/VI positive FMDV genome was detected in 179/199 samples by RT-PCR (for simplicity, using <40 as the CT value cut-off point; Table V). A further 10 and 2 samples produced CT values in the range >40-<45 and >45-<50, respectively, while the remaining 8 samples generated a CT value of 50. FMDV failed to be isolated in 3/6, 0/2 and 4/5 of the samples which were available from each category for subsequent cell culture passage. The resulting cell culture grown antigens were tested by RT-PCR with positive results, except one virus (UKG 13795/2001, originating from Crickhowell, Powys, Wales), which consistently yielded a CT value of 50 from replicate tests. This virus isolate was sequenced to see if the analysis might indicate why this virus was undetected by RT-PCR. It was previously known that the Pan Asia type O FMDVs, whilst detectable by the real-time RT-PCR assay, contain a nucleotide substitution within the TaqMan® probe region (Fig.1). The causative FMDV strain of the UKG outbreak contains an additional substitution resulting in two mis-matches with the universal probe. The Welsh virus was found to have a further, additional nucleotide substitution resulting in three mis-matches within the probe region. This was evidently sufficient to make it unrecognised by the conventional diagnostic probe. This prompted a mini-study to examine the reaction of other virus isolates from the Crickhowell region (all of which had originally typed as positive by ELISA on epithelial suspension) in RT-PCR procedures using either the conventional diagnostic probe or one (P282T) designed to match the Welsh mutant virus. The results of these tests are shown in Table V1 and indicate that all these isolates were related, failed to be detected by the RT-PCR using the conventional diagnostic probe but were recognised by the new probe. Discussion: The ELISA and VI have been recommended laboratory procedures for FMD diagnosis for nearly twenty years based on their suitability to detect the presence of FMDV antigen in tissue samples. It is evident from the present study that these procedures are deficient for certain specimens. The results for the overseas samples show that while all VI/ELISA positive FMDV samples were also positive by real-time RT-PCR (100% sensitivity at a CT cut-off value of <39) (Table III), FMD viral genome was detected in a significant proportion of the samples examined in which FMDV antigen was not. The low relative specificity values (a range in value from 63.1%. down to 38.1% depending upon cutoff) of the RT-PCR result from the doubtful premise that the combination of VI and ELISA is 100% efficient for detection of FMDV. The authors are confident that this is not the case, that the RT-PCR procedure has specific reaction for FMDV genome and that the low value is actually related both to the performance of the comparative assays and to the quality of the samples submitted. If one considers what the VI and ELISA procedures actually measure then it is evident that their effectiveness for diagnostic use is inherently compromised. Virus isolation is dependent upon the presence of infectious virus in sample submissions and while the ELISA can detect both infectious and non-infectious FMD viral antigen, it is dependent upon the antigen being present in sufficient concentration (1-2 ng/ml of antigen or 5-6 log10/ml of live virus) to work (N.P. Ferris, unpublished results). If neither of these two conditions is met then FMDV will not be recognised. Ideally, vesicular epithelium should be collected from an animal during the acute stage of FMD when the concentration of virus associated with the sample is high. Unfortunately, samples submitted to the WRL for FMD from overseas are very often collected late in the course of disease when the amount of virus may either be waning or indeed be absent after clearance. Delayed reporting of 263 disease and late sample collection can arise for a variety of reasons, e.g. a lack of resources, communication difficulties in areas of rugged terrain or a low perceived importance of FMD. Delays can be worse depending on who has responsibility for sample collection; in some countries it is local staff and in others countries staff from a specialised laboratory (often many miles away) carry out the collection (Ferris et al., 1992). Secondary bacterial infection is a common sequel to virus infection and can lead to a reduction in FMDV infectivity. Additionally, samples may be in transit for lengthy periods and subjected to physico-chemical stresses (e.g. elevated temperatures between collection and laboratory receipt) with the result that on arrival only small amounts of infectious virus, at best, may be present. FMDV survival can be adversely affected by harsh environmental conditions, including excessive temperature, extremes of pH, disinfectants and desiccation. It is therefore advantageous to protect samples during the interval between collection and testing (especially for VI). Their dispatch to reference laboratories should follow specific guidelines to ensure their security (Kitching and Donaldson, 1987). These features, which influence sample quality, are likely to be less important for the RT-PCR as it can detect a small fragment of FMDV genomic RNA, not just live virus. There was broad correlation between the results achieved by RT-PCR examination of the UK outbreak sample subsets with the original (2001) diagnostic results derived from ELISA/VI suggesting that definitive sample classification of both positivity and negativity by RT-PCR is achievable within a relatively short timescale. However, the failure to detect the group of related virus isolates from Wales (using the routine diagnostic probe), which had evidently evolved during the epidemic and had a further nucleotide substitution in the RT-PCR probe site, illustrates the importance of constant monitoring of representative field FMDV strains by nucleotide sequencing to ensure that the primers/probe set selected for the diagnostic RT-PCR is fit for purpose. Conclusions: • The real-time RT-PCR currently used at the WRL for FMD provides an extremely sensitive and rapid additional procedure for improved laboratory diagnosis of FMD Recommendations: • No specific recommendations Acknowledgements: This work was funded by the Department for the Environment, Food and Rural Affairs, United Kingdom (project number SE1119). References: De Castro. M.P. 1964. Behaviour of the foot-and-mouth disease virus in cell cultures: susceptibility of the IB-RS-2 line. Arch. Inst. Biol., São Paulo, 31: 63-78. Ferris, N.P. & Dawson, M. 1988. Routine application of enzyme-linked immunosorbent assay in comparison with complement fixation for the diagnosis of foot-and-mouth and swine vesicular diseases. Vet. Microbiol., 16 (3): 201-209. Ferris, N.P. & Doanaldson, A.I. 1992. The world reference laboratory for foot and mouth disease: a review of thirty-three years of activity (1958-1991). Rev. Sci. Tech. Off. Int. Epiz., 11 (3): 657-684. Ferris, N.P., Donaldson, A.I., Shrestha, R.M. & Kitching, R.P. 1992. A review of foot and mouth disease in Nepal. Rev. Sci. Tech. Off. Int. Epiz., 11 (3): 685-698. Kitching, R.P. & Doanldson, A.I. 1987. Collection and transportation of specimens for vesicular virus investigation. Rev. Sci. Tech. Off. Int. Epiz., 6 (1): 263-272. Knowles, N.J. & Samuel, A.R. 2003. Molecular epidemiology of foot-and-mouth disease virus. Virus Res., 91: 65-80. OIE (World Organisation for Animal Health) 2004. Manual of diagnostic tests and vaccines for terrestrial animals. 5th Ed. Parts I and II. OIE, Paris, 1178 pp. Oleksiewicz, M.B., Doanldson, A.I. & Alexandersen, S. 2001. Development of a novel real-time RT-PCR assay for quantitation of foot-and-mouth disease virus in diverse porcine tissues. J. Virol. Meth. 92 (1): 23-35. 264 Reid, S.M., Ferris, N.P., Hutchings, G.H., Zhang, Z., Belsham, G.J. & Alexandersen S. 2002. Detection of all seven serotypes of foot-and-mouth disease virus by real-time, fluorogenic reverse transcription polymerase chain reaction assay. J. Virol. Meth., 105 (1): 67-80. Reid, S.M., Grierson, S.S., Ferris, N.P., Hutchings, G.H. & Alexandersen, S. 2003. Evaluation of automated RT-PCR to accelerate the laboratory diagnosis of foot-and-mouth disease virus. J. Virol. Meth. 107 (2): 129-139. Roeder, P.L. & Le Blanc Smith, P.M. 1987. Detection and typing of foot-and-mouth disease virus by enzyme-linked immunosorbent assay: a sensitive, rapid and reliable technique for primary diagnosis. Res. Vet. Sci. 43: 225-232. Snowdon, W.A. 1966. Growth of foot-and-mouth disease virus in monolayer cultures of calf thyroid cells. Nature, 210 (40): 1079-1080. Table 1. Number of samples received by the WRL for FMD from overseas between 1994 and 2003 and the numbers (and percentages) found to be positive or negative for FMD virus by passage in cell culture and antigen detection ELISA Year a 1994 1995 1996 1997 1998 1999 2000 2001 2002 2003 Total number of Countriesa Number of samples Total Positive 29 655 348 (53%) 29 697 421 (60%) 31 535 238 (44%) 29 526 334 (63%) 27 441 248 (56%) 43 595 357 (60%) 29 434 209 (48%) 32 619 197 (32%) 27 390 162 (42%) 19 475 262 (55%) 5,367 2,776 (52%) countries which submitted samples to the WRL for FMD Negative 307 (47%) 276 (40%) 297 (56%) 192 (37% 193 (44%) 238 (40%) 225 (52%) 422 (68%) 228 (58%) 213 (45%) 2,591 (48%) Fig. 1. Nucleotide sequence changes within the TaqMan® probe site of the Pan Asia type O FMD virus, the causative UKG 2001 FMD virus and the related Welsh viruses compared to the FMD virus consensus sequence. FMD virus consensus sequence: Pan Asia Strain: UKG 2001 FMD virus: Related Welsh UKG 2001 FMD virus: TaqMan® Probe Site 5’-GGATGCCCTTCAGGTACCCCGAGG-3’ 5’-GGATGCCCTTCAGGTACCCTGAGG-3’ 5’-GGATGCCCTTTAGGTACCCTGAGG-3’ 5’-GGATGCCCTTTAAGTACCCTGAGG-3’ Nucleotide substitutions are indicated X 265 Table II. Detection of FMD virus in suspensions of submitted epithelia achieved by ELISA, passage in cell culture (VI), ELISA and VI combined and by RT-PCR (August 2002-January 2004) Country No VIb ELISA FMDV NVDa FMDV ELISA/VI NVD FMDV Bhutan 58 2 56 17 41 Botswana 5 5 5 Burundi 7 4 3 5 2 Hong 7 1 6 3 4 Kong Iran 43 17 26 31 12 Iraq 18 1 17 6 12 Laos 35 18 17 33 2 Lebanon 5 3 2 3 2 Libya 1 1 1 Malaysia 12 8 4 12 Nepal 4 4 4 Pakistan 90 36 54 41 49 1 1 1 PATd Philippines 9 8 1 7 2 Thailand 7 6 1 7 Turkey 24 10 14 17 7 3 1 2 3 UAEe Vietnam 5 5 4 1 Total 334 125 209 195 139 a NVD, no virus detected b VI, serotype of FMD virus isolated by passage (VI) in c CT, threshold cycle value d PAT, Palestinian Autonomous Territories f UAE, United Arab Emirates NVD 17 5 3 41 5 2 4 31 7 33 5 1 12 4 44 1 8 7 18 3 5 204 12 11 2 46 1 6 130 RT-PCR for FMD virus (CTc value) <35 >35 >39 >45 50 <39 <45 <50 36 4 6 12 1 4 1 4 1 1 6 1 32 11 33 5 12 4 59 1 8 7 21 3 5 244 1 1 2 1 5 1 20 2 1 6 16 1 1 1 8 5 20 3 53 cell culture characterised by ELISA Table III. Relative sensitivity of the real-time RT-PCR for FMD virus at successive threshold cycle value cut-off points in comparison with VI in cell culture or VI plus ELISA combined Threshold cycle Value cut-off <35 <36 <37 <38 <39 <40 <41 <42 <43 <44 <45 <46 <47 <48 <49 <50 a VIa Sensitivity 95.9 97.0 99.0 99.5 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 100.0 Specificity 59.0 56.1 54.0 51.1 50.4 46.0 43.2 42.4 40.3 40.3 38.8 38.8 38.8 38.1 38.1 38.1 VI/ELISA combined Sensitivity Specificity 96.1 63.1 98.0 60.0 99.0 57.7 99.5 54.6 100.0 53.8 100.0 49.2 100.0 46.2 100.0 45.4 100.0 43.1 100.0 43.1 100.0 41.5 100.0 41.5 100.0 41.5 100.0 40.8 100.0 40.8 100.0 40.8 VI, virus isolation in cell culture with specificity of cytopathic effect confirmed by ELISA 266 Table IV. Detection of FMD virus by RT-PCR in suspensions of submitted epithelia from clinically confirmed cases found to be ELISA/VI negative after sample receipt in 2001 Animal No of samples Sheep 253 Cattle 98 Pig 14 Cattle + Sheep 5 Deer 1 Not known 9 Total 380 a CT, threshold cycle value <35 2 5 7 RT-PCR for FMD virus (CTa value) >35-<40 >40-<45 >45-<50 2 2 2 2 2 2 50 245 93 14 5 1 9 367 Table V. Detection of FMD virus by RT-PCR in suspensions of submitted epithelia from clinically confirmed cases found to be ELISA negative/VI positive after sample receipt in 2001 Animal No of samples Sheep 131 Cattle 60 Cattle+Sheep 2 Goat 3 Not known 3 Total 199 a CT, threshold cycle value <35 87 41 2 1 2 133 RT-PCR for FMD virus (CTa value) >35-<40 >40-<45 >45-<50 33 6 1 12 3 1 1 1 46 10 2 50 4 4 8 Table VI. Detection of ‘mutant’ FMD virus by RT-PCR in suspensions of submitted epithelia from clinically confirmed cases in Powys, Wales found to be positive after sample receipt in 2001 Sample ref Animal Origin FMD No. UKG 13795/2001e Sheep Crickhowell, Powys 1861 RT-PCR for FMD virus (CTb value) P269Rc P282Td 50 35.81 UKG 13705/2001 Cattle Crickhowell, Powys 1846 50 32.35 UKG 13708/2001 Cattle Crickhowell, Powys 1848 50 31.52 UKG 13724/2001 Cattle Powys 1852 50 29.75 UKG 13734/2001 Cattle Powys 1857 50 27.6 UKG 13777/2001 Cattle Powys 1860 50 40.91 Crickhowell, Powys 1868 50 31.06 UKG 13798/2001 NKf UKG 13949/2001 Cattle Abergavenny, Powys 1889 50 32.85 UKG 14004/2001 Cattle Crickhowell, Powys 1896 50 31.46 UKG 14221/2001 Cattle Abergavenny, Powys 1938 50 27.59 UKG 14339/2001 Cattle Crickhowell, Powys 1945 50 31.08 a OD, optical density value originally achieved on the epithelial suspension b CT, threshold cycle value c P269R, the normal diagnostic probe d P282T, ‘UK’ probe e UKG 13795/2001 serotyped on first passage cell culture antigen, all other virus isolates listed serotyped on epithelial suspension f NK, not known 267