Download Positive RT-PCR test results in tonsils of foot and mouth disease virus infected piglets after more than 28 days

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