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Appendix 61 Positive RT-PCR test results in tonsils of foot and mouth disease virus infected piglets after more than 28 days K. Orsel1*, H.I.J. Roest 2, E. van der Linde2, F. van Hemert-Kluitenberg2 and A. Dekker2 1. Faculty of Veterinary Medicine, Department of Farm Animal Health, Utrecht University, Utrecht, The Netherlands 2. Central Institute for Animal Disease Control Lelystad (CIDC-Lelystad), Wageningen UR * P.O. box 80.151,3508 TD Utrecht, The Netherlands. Tel.: +31 30 2537551, Fax: +31 30 2521887 Email address: [email protected] Abstract: The aim of this study was to determine whether FMDV can be detected in tonsils of pigs, at least 28 days after initial infection. Material and Methods: Piglets originated from two different transmission experiments with FMDV O/NET/2001. Infection with FMDV was confirmed in these piglets either with positive test results in the virus titration test on oro-pharyngeal fluid collected with swabs or heparinised blood, or with a positive serological response with the NS ELISA or by showing clinical signs of FMD. Fourteen days after infection all results from the virus titration tests of OPF and heparinised blood, were negative, indicating the piglets to be recovered from FMDV infection. At the end of the studies, i.e. 28 days after inoculation of the seeder pigs in experiment 1, and 28 days after infection of contact exposed pigs in experiment 2, all pigs were killed, and the tonsils were collected. From all tonsil plates a part of 1x1 cm was prepared in the MagNALyser® or ground with a pestle and morter for testing in RT-PCR. This resulted in positive test result in 13 tonsil samples of 2 vaccinated and 11 nonvaccinated piglets; all piglets showed clinical signs. Discussion: Our findings confirm the persistence of viral genome in tonsils of piglets at least 28 days after inoculation of seeder pigs. Hereby we provide more evidence of pigs being carriers of FMDV, and the tonsil tissue as a location of persistence. Introduction: Foot-and-mouth disease (FMD) is a highly contagious viral disease and can cause severe outbreaks in susceptible populations. Not only effect on animal health and welfare, but also the economic impact of the disease is of great importance, especially for exporting countries. Limitations on export are partly based on the possible role of carriers in transmission of FMD virus. It has been suggested that carrier animals imported from infected countries may have been responsible for outbreaks in importing countries (Moonen & Schrijver, 2000, Salt, 1993). Persistent infection in carrier animals has been defined as presence of virus more than 28 days after infection (Sutmoller & Gaggero, 1965). In 1959 van Bekkum et al reported the recovery of tissue culture infectious FMDV in the oropharyngeal scrapings collected with a probang sampling cup from FMD convalescent and sub-clinically infected vaccinated cattle. This method has been the ‘standard’ detection method for carrier animals ever since (Van Bekkum, 1959). Occurrence of carriers in cattle can been observed in approximately 50% of the cattle after clinical and sub clinical infection and in animals with and without vaccination (Salt, 1993), and also other ruminants like sheep, goats and African buffalo have been recognized as carriers of FMD virus. Many studies tried to define carrier state in pigs as well; it was shown that pigs cleared the virus after 3-4 weeks (Alexandersen et al., 2003), although Mezencio et al describes the finding of pigs being carriers by identifying FMDV-RNA in sera of pigs after 266 days (Mezencio et al., 1999). Oleksiewicz showed PCR positive test results in tonsil samples of pigs at 2 and 3 days post infection (Oleksiewicz et al., 2001), and also with in situ hybridization techniques positive results on tonsil tissues were reported (Brown et al., 1995). The aim of this study was to determine whether FMDV can be detected in tonsils in pigs with RTPCR, at least 28 days after initial infection. 392 Materials and Methods: Tonsils were available from two transmission experiments performed with piglets of 10 weeks old (publication in preparation). The first study included 11 non-vaccinated seeder piglets inoculated intradermally with approximately 10.000 plague forming units of FMDV O/NET/2001 in the heelbulb, and 12 contact-exposed pigs, which were either vaccinated (n=6) or non-vaccinated (n=6). Vaccination was performed with a standard dose of double oil emulsion O1Manisa vaccine 15 days prior to contact with inoculated piglets. The second experiment included pigs infected through contact with FMDV O/NET/2001. Six vaccinated but not-infected animals were included as vaccine virus controls. 30 Days after inoculation (dpi) or 29 days after in-contact exposure all piglets were slaughtered. Tonsils were also collected from eight piglets (slaughtered for other purposes) as additional negative controls. From the central part of the left tonsil a biopsy of 1 by 1 cm was collected. The tonsil biopsies were stored at -700 C until analysis. The biopsies were ground with a pestle and mortar using sterile sand and EMEM containing 2% fetal calf serum (FCS) and 10% mixed antibiotics. All samples were handled in a class II laminar flow cabinet to prevent contamination. The tissue suspensions were tested for presence of FMDV on a monolayer of secondary lamb kidney cells. In total 200 µl of tonsil tissue suspension was added to one well of a 6-well plate (Greiner®). After one hour incubation, the wells were washed with fresh medium and 2,5 ml of fresh medium was added. The cells were macroscopically observed for cytopathogenic effects for 2 days. If no cyto-pathogenic effect was observed the cell and supernatant were frozen and thawed and 200 µl was handled once like the original suspension. All incubations were performed at 370 C in a humidified atmosphere containing 5% CO2. The tonsil suspensions were also tested by automated real-time RT-PCR; RNA isolation was performed using the Magna Pure LC total Nucleic Acid Isolation kit (3 038 505) in the MagNA Pure ® system (Roche). The isolated RNA was tested in a Light Cycler based RT-PCR with use of Light Cycler RNA Master Hybridization Probes (3 0180954), all in accordance with the manufacturers instructions (Roche). Using the Light Cycler system with hybridisation probes performed in a closed glass capillary, real time detection of the amplification product is allowed with minimised risk of crosscontamination. In each run a low positive and negative control was included. The primers and probes used are fully described by Moonen (Moonen et al., 2003). The RT-PCR for FMD virus is validated within the laboratory facilities of CIDC-Lelystad. Results: No virus could be isolated from the tonsil samples of experimentally infected animals or vaccineand negative controls. The positive test controls gave a positive result as expected, indicating a technically correct test result. In contrast, 2 out of 25 vaccinated animals and 11 out of 31 nonvaccinated animals tested positive in the RT-PCR (table 1-2-3) (Fisher’s exact p=0.024). All positive RT-PCR samples originated from animals that had shown clinical signs of FMD. Discussion: We detected viral RNA using RT-PCR in tonsils of two vaccinated an 23 non vaccinated pigs, which indicates that pigs could also become carriers according to the definition. However, we did not detect virus using the less sensitive VI tests, so it cannot be determined whether these pigs would pose a risk for transmission of infectious virus. In carrier animals FMD virus is know to be located in the oro-pharyngeal region and in more detail, in the dorsal soft palate (Zhang & Alexandersen, 2004, Zhang & Kitching, 2001). In pigs, tonsillar tissue can be found (Dyce et al., 2002) throughout the oropharynx, so we therefore directed our sampling towards the tonsils of pigs. Expected amounts of virus in op-fluid samples are generally low during persistence (Zhang et al., 2004) and highly variable (Alexandersen et al., 2002). Due to the high sensitivity of the real time RT-PCR for FMD (Callahan et al., 2002, Moonen et al., 2003, Reid et al., 2002, Shaw et al., 2004) samples with a low concentration of viral genome still produced a positive result, though negative by virus isolation. This technique can be used as a quantitative assay at higher concentrations, but at low concentrations the crossing point observed in our real time RT-PCR is almost identical (table 1-2-3).. But virus isolation techniques on probang samples are laborious, time-consuming and have low sensitivity (Zhang & Alexandersen, 2003), so RT-PCR is preferred. Zhang & Alexandersen showed a correlation between the presence of viral RNA in pharyngeal tissues and the recovery of infectious virus in OP fluid samples in cattle (Zhang & Alexandersen, 2004). We did not test probang samples or OP fluid samples, but with this relation presented it might also lead to positive probang samples in pigs. All our positive test results on tonsillar tissue originated from piglets that tested positive in OPF during the acute infection with FMD. 393 Recommendations: * All samples originate from experimental studies designed for other purposes, based on the current findings, it would be interesting to follow up the pigs for a longer period in time, both for OPF samples and tissue samples. * Discussion in literature on carrier status of pigs: To overcome discussions on false positive test results, other independent diagnostic techniques could help (in situ pcr incorporating digoxigeninlabeled dUTP (Prato Murphy et al., 1995), histopathologicall and in-situ hybridization (Brown et al., 1995), in situ hybridization also by using biotin-labelled oligodeoxynucleotides and tyramide signal amplification (Zhang & Kitching, 2000, Zhang & Kitching, 2001)), but these tests are not as sensitive as the RT-PCR. * Combine knowledge on carrier status, location of persistent virus and diagnostic possibilities to sustain the carrier status in pigs. Acknowledgements: •Financial support from the ministry of agriculture in the Netherlands (BOP 7-428) •Financial support from the European Union (EU SSPE-CT-2003-503603) •Analists from the FMD-laboratory in Lelystad •Animal technicians in Lelystad References: Alexandersen, S., Zhang, Z. & Donaldson, A. (2002). Aspects of the persistence of foot-andmouth disease virus in animals-the carrier problem. Microbes Infect 4, 1099. Alexandersen, S., Zhang, Z., Donaldson, A. I. & Garland, A. J. (2003). The pathogenesis and diagnosis of foot-and-mouth disease. J Comp Pathol 129, 1-36. Brown, C. C., Olander, H. J. & Meyer, R. F. (1995). Pathogenesis of foot-and-mouth disease in swine, studied by in-situ hybridization. J Comp Pathol 113, 51-8. Callahan, J. D., Brown, F., Osorio, F. A., Sur, J. H., Kramer, E., Long, G. W., Lubroth, J., Ellis, S. J., Shoulars, K. S., Gaffney, K. L., Rock, D. L. & Nelson, W. M. (2002). Use of a portable real-time reverse transcriptase-polymerase chain reaction assay for rapid detection of foot-and-mouth disease virus. J Am Vet Med Assoc 220, 1636-42. Dyce, K. M., Sack, W. O. & Wensing, C. J. G. (2002). Textbook of veterinary anatomy, 3rd edn. Philadelphia: Saunders. Mezencio, J. M., Babcock, G. D., Kramer, E. & Brown, F. (1999). Evidence for the persistence of foot-and-mouth disease virus in pigs. Vet J 157, 213-7. Moonen, P., Boonstra, J., van der Honing, R. H., Leendertse, C. B., Jacobs, L. & Dekker, A. (2003). Validation of a LightCycler-based reverse transcription polymerase chain reaction for the detection of foot-and-mouth disease virus. J Virol Methods 113, 35-41. Moonen, P. & Schrijver, R. (2000). Carriers of foot-and-mouth disease virus: a review. Vet Q 22, 193-7. Oleksiewicz, M. B., Donaldson, A. I. & Alexandersen, S. (2001). Development of a novel realtime RT-PCR assay for quantitation of foot-and-mouth disease virus in diverse porcine tissues. J Virol Methods 92, 23-35. Prato Murphy, M. L., Rodriguez, M., Schudel, A. A. & Meyer, R. F. (1995). Localization of foot and mouth disease virus RNA in tissue culture infected cells via in situ polymerase chain reaction. J Virol Methods 54, 173-8. 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 Methods 105, 6780. 394 Salt, J. S. (1993). The carrier state in foot and mouth disease--an immunological review. Br Vet J 149, 207-23. Shaw, A. E., Reid, S. M., King, D. P., Hutchings, G. H. & Ferris, N. P. (2004). Enhanced laboratory diagnosis of foot and mouth disease by real-time polymerase chain reaction. Rev Sci Tech 23, 1003-9. Sutmoller, P. & Gaggero, A. (1965). Foot-and mouth diseases carriers. Vet Rec 77, 968-9. Van Bekkum, J. G., Frenkel HS, Frederiks H and Frenkel S. (1959). Observations on the carrier state of cattle exposed to foot-and-mouth disease virus. tijdschr Diergeneeskd 20, 1159-64. Zhang, Z. & Alexandersen, S. (2003). Detection of carrier cattle and sheep persistently infected with foot-and-mouth disease virus by a rapid real-time RT-PCR assay. J Virol Methods 111, 95-100. Zhang, Z. & Alexandersen, S. (2004). Quantitative analysis of foot-and-mouth disease virus RNA loads in bovine tissues: implications for the site of viral persistence. J Gen Virol 85, 256775. Zhang, Z. & Kitching, P. (2000). A sensitive method for the detection of foot and mouth disease virus by in situ hybridisation using biotin-labelled oligodeoxynucleotides and tyramide signal amplification. J Virol Methods 88, 187-92. Zhang, Z., Murphy, C., Quan, M., Knight, J. & Alexandersen, S. (2004). Extent of reduction of foot-and-mouth disease virus RNA load in oesophageal-pharyngeal fluid after peak levels may be a critical determinant of virus persistence in infected cattle. J Gen Virol 85, 415-21. Zhang, Z. D. & Kitching, R. P. (2001). The localization of persistent foot and mouth disease virus in the epithelial cells of the soft palate and pharynx. J Comp Pathol 124, 89-94. 395 Table 1: Results from experiment 1 Animal Vaccination NS ELISA Clinical no status signs 8671 8677 8669 8667 8663 8659 8665 8675 8679 8681 8661 8670 8672 8674 8676 8678 8680 8668 8666 8658 8662 8664 8660 Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated + + + + + + + + + + + + + + + + + - VI OPF acute phase Yes Yes Yes Yes Yes Yes No Yes Yes No No Yes Yes Yes Yes Yes No Yes Yes Yes Yes No No Table 2: Results from control samples Animal Vaccination NS Clinical no status ELISA signs VI blood VI or CPE RT-PCR CP acute > 28 > 28 lightcycler days days phase + + + + + + + + + + + + + + + + + - VI OPF acute phase + + + + + + + + + + + + + + + + + - Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated - No No No No No No - - positive control n=2 - - + - - - - n=8 + + + + + + + + + + + + - 28,6 28,6 28,8 28,8 28,9 28,9 28,6 28,8 28,8 29,7 28,7 28,9 - VI blood VI or RT-PCR CP lightcycler acute CPE > > 28 days phase 28 days 8682 8683 9546 9547 9548 9549 negative control - - - - + + 28,2 + + + 28.5 - - - - - 396 Table 3: Results from experiment 2 Animal Vaccination NS ELISA Clinical status signs no 9043 9044 9045 9046 9047 9048 9506 9509 9510 9511 9512 9513 9514 9515 9521 9522 9523 9524 9525 9526 9527 9529 9530 9536 9537 9538 9539 9540 9541 9542 9543 9544 9545 Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated Vaccinated + + + + + + + + + + + + + + + + + + + + + + + + + + Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes No No Yes No No Yes No Yes Yes Yes No Yes No No Yes No Yes No VI OPF acute phase + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + + VI blood VI or CPE RT-PCR CP acute > 28 > 28 lightcycler days days phase + + + + + + + + + + + + + + + + + + + + + + - - + - 27.05 - Captions to table 1-2-3 + : at least tested positive once - : negative in all samples VI : virus isolation CP = crossing point (2nd derivative) OPF = oro-pharyngeal fluid collected with swabs CPE = cytopathogenic effect 397