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1 Institute for Animal Health (IAH), Division of Immunology, Compton, Newbury RG20 7NN, UK
2 Department of Virology, Division of Investigative Science, Faculty of Medicine, Imperial College London, St Mary's Campus, Norfolk Place, London W2 1PG, UK
Correspondence
Shirley A. Ellis
shirley.ellis{at}bbsrc.ac.uk
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). FMDV shows high genetic and antigenic variability, which is reflected in the seven serotypes and multiple subtypes reported to date (Domingo et al., 2003). The virus causes a highly contagious infection (foot-and-mouth disease; FMD) in cloven-hooved animals that is characterized by the formation of vesicles on the mouth, tongue, nose and feet; most infected animals develop viraemia. The 2001 UK outbreak resulted in the slaughter of four million animals (Scudamore & Harris, 2002) and an estimated total loss to the UK economy of between £8.2 billion and £9.2 billion (between US$ 12.3 billion and 13.8 billion) (Thompson et al., 2002).
The virus elicits a rapid humoral response in both infected and vaccinated animals (Grubman & Baxt, 2004). Virus-specific antibodies protect animals in a serotype-specific manner against reinfection, or against infection in the case of vaccination, and protection is generally correlated with high levels of neutralizing antibodies (McCullough et al., 1992). Control of the disease is achieved by vaccination with a chemically inactivated whole-virus vaccine emulsified with adjuvant; however, this provides only short-term, serotype-specific protection (Barteling & Vreeswijk, 1991). Such chemically inactivated vaccines have a number of disadvantages, including the requirement for a cold chain to preserve capsid stability, the need for periodic revaccination with virus strains antigenically similar to circulating viruses, and the risk of virus release during vaccine production (Mason et al., 2003). The introduction of the killed FMDV vaccine has been very successful in areas of the world where the disease is enzootic. However, one of the major difficulties in implementing vaccination is the inability to distinguish vaccinated from infected/recovered animals, which may still be shedding virus. Currently, a number of assays developed specifically for this purpose are being validated (Mezencio et al., 1998; Oem et al., 2007; Shen et al., 1999).
Identification and characterization of T-cell epitopes are important for understanding protective immunity mediated by CD8+ and CD4+ T lymphocytes. Such T-cell responses are pathogen-specific and are restricted by major histocompatibility complex (MHC) genes, which encode the class I and II molecules responsible for the presentation of foreign peptides to the immune system (Townsend et al., 1985, 1986). The role of cellular immunity in the protection of animals from FMD is still a matter of some controversy. Specific T-cell-mediated antiviral responses have been observed in cattle and swine following either infection or vaccination (Bautista et al., 2003; Blanco et al., 2001; Childerstone et al., 1999; Glass et al., 1991) and it has been suggested that cell-mediated immunity is involved in clearance of the virus from persistently infected animals (Ilott et al., 1997). CD4+ T-cell responses seem to play an important role in protection against FMD, and several publications demonstrate the presence of FMDV-specific MHC class II-restricted responses in cattle (Glass et al., 1991; Van Lierop et al., 1995) and pigs (Blanco et al., 2001; Garcia-Briones et al., 2000; Gerner et al., 2006). CD8+ T-cell-mediated immune responses to FMDV have been reported in pigs (Blanco et al., 2001; Garcia-Briones et al., 2004) and cattle (Childerstone et al., 1999); however, MHC class I restriction has not been demonstrated, and the significance of cell-mediated immune responses in terms of protective immunity to FMDV remains unclear.
Relative to humans, MHC class I expression in cattle is unusually complex. Recent molecular studies have shown that haplotypes may express one, two or three classical class I genes from a putative total of six (Birch et al., 2006; Ellis et al., 1999). At present, about 60 full-length validated cattle MHC class I cDNA sequences are available (http://www.ebi.ac.uk/ipd/mhc/bola), and the haplotypes commonly found in the Holstein breed are well-characterized.
In the present study, we evaluated the presence of FMDV-specific CD8+ T-cell responses in cattle of known MHC class I phenotype. By using a gamma interferon (IFN-
) ELIspot, we detected antigen-specific MHC-restricted CD8+ T-cell responses following infection with FMDV or vaccination with a chemically inactivated vaccine.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The O1 Manisa vaccine was prepared from antigen concentrate stored over liquid nitrogen, held at a commercial facility as part of a new UK strategic reserve. In accordance with the European Pharmacopoeia Monograph, this commercially produced oil-adjuvant vaccine had been shown to have a PD50 value of 18.
Fowlpox virus (FPV) recombinants expressing the P1 and 2A regions of FMDV serotype O were constructed by using an approach described previously for an antigen from an avian pathogen (Qingzhong et al., 1994). Briefly, wild-type FPV FP9 was grown in primary chicken embryo fibroblasts (CEFs; IAH, Compton, UK) for 5 days in E199 medium supplemented with 10 % fetal calf serum (FCS; Autogen Bioclear) and purified by centrifugation. The polyprotein P1 and 2A of FMDV O Manisa/67 was amplified by RT-PCR using the following primers: 5'-ATTCCCGGGATGGGAGCTGGGCAATCCAGCCCAG-3' (sense) and 5'-CGGCCCGGGTCACTCCAGGGTTGGACTCCACATC-3' (antisense) (Sigma Genosys). An initiation and a termination codon were introduced (underlined) and SmaI sites were introduced on each flank (bold) for blunt-end cloning into the FPV vector pEFL29 (Qingzhong et al., 1994). Primary CEFs grown in E199 medium supplemented with 10 % FCS (Autogen Bioclear) in T75 flasks were infected with parenteral attenuated FPV FP9 (Laidlaw & Skinner, 2004) at an m.o.i. of 2 for 1 h, washed with serum-free E199 medium and then transfected with 10 µg pEFL29-FMD-P12A by using Lipofectin (Invitrogen), following the manufacturer's instructions. The cells were then incubated at 37 °C for 5 days. Recombinant viruses were identified as blue plaques on monolayers of CEFs overlaid with Sea Kem agarose (Invitrogen) containing 5-bromo-4-chloro-3-indolyl-β-D-galactosidase (X-Gal). Recombinant FPV was plaque-purified three times. All constructs were confirmed by DNA sequencing (IAH Sequencing Service).
P815-BoLA class I transfectants were prepared as described previously (Gaddum et al., 1996; Kydd et al., 2006).
Animal experiments.
For the vaccination study, seven Holstein cattle (Bos taurus) were vaccinated intramuscularly with one bovine dose of commercially available serotype O inactivated vaccine (Merial) and boosted 11 weeks later. For the infection study, six male Holstein calves aged between 6 and 9 months and weighing approximately 150 kg were exposed for 24 h to calves that had been infected 24 h previously by subepidermolingual injection with approximately 1x105 TCID50 FMDV O UKG/34/2001. Clinical signs and rectal temperatures were recorded over an 8 day period and the animals were maintained in isolation for up to 60 days. The needle-infected calves were not used in this study.
The majority of animals used in this study (Table 1) were from a partially inbred herd, generated by back-crossing, in which MHC class I haplotypes have been characterized at the level of expressed genes (Ellis et al., 1996, 1998). Details of cattle MHC haplotypes, alleles and nomenclature can be found at http://www.ebi.ac.uk/ipd/mhc/bola/. Animals were typed by PCR using sequence-specific primers (PCR-SSP), reference strand conformational analysis (RSCA; Arguello et al., 1998; Birch et al., 2006) or indirect immunofluorescence (Table 2).
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), following the manufacturer's instructions. CD8+ 
/β T cells were positively selected by incubating peripheral blood mononuclear cells with the mAb CC58 (MacHugh et al., 1991) (IAH, Compton, UK) and then with anti-mouse IgG1 microbeads (Miltenyi Biotec). To prepare antigen-presenting cells (APCs), CD14+ cells were incubated for 7 days at 37 °C in RPMI 1640 medium (Invitrogen) containing 10 % heat-inactivated FCS, 2 mM L-glutamine, 55 µM 2-mercaptoethanol and 1 % penicillin/streptomycin (IAH, Compton, UK) and supplemented with recombinant bovine granulocyte–macrophage colony-stimulating factor (a gift from Jayne Hope, IAH, Compton, UK).
Flow cytometry.
Lymphocyte-subset purities were monitored by using the following mAbs: CC8 (IgG2a, anti-bovine CD4) (Howard et al., 1989; Morrison et al., 1994); CC58 (IgG1, anti-bovine CD8 /β) (MacHugh et al., 1991); CC63 (IgG2a, anti-bovine CD8 /β and / chain) (MacHugh et al., 1991); CC15 (IgG2a, anti-bovine WC1) (Fikri et al., 2000); and GB21A (IgG2b, anti-bovine 
T-cell receptor) (Davis et al., 1996). The purity of the CD8+ T cells was tested by flow cytometry prior to each assay and >95 % of the cells separated by MACS were found to be CD8+ β T cells. Flow cytometry was performed as described previously (Bruce et al., 1999) in a BD Biosciences FACScalibur flow cytometer and data were analysed by using CellQuest Pro v5.2 software (BD Biosciences).
Ex vivo IFN- ELIspot assay.
MultiScreen-HA plates (mixed cellulose ester membranes; Millipore) were coated overnight at 4 °C with 100 µl anti-IFN- CC330 (2 µg ml–1; IAH, Compton, UK) in distilled water. The plates were washed with PBS+0.05 % (v/v) Tween 20 (PBS-T), blocked with 5 % BSA in PBS for 2 h and then washed prior to use. APCs were prepared by infecting CD14+ cells with FMDV or rFPV-FMD-P12A (m.o.i.=5). Alternatively, mouse P815 cells expressing individual BoLA class I genes were infected with rFPV-FMD-P12A (m.o.i.=5). APCs infected with wild-type FPV (m.o.i.=5) or incubated with CEF or BHK-21 cell lysate were used as negative controls. Effector cells (CD8+ β subset) were added to wells in triplicate and stimulated with autologous or heterologous APCs or P815 transfectants at an APC : CD8+ T-cell ratio of 5 : 1. After 24 h incubation at 37 °C, supernatants were discarded and 200 µl cold distilled water per well was added for 10 min. Plates were washed with PBS-T, and 100 µl biotinylated mouse anti-bovine INF- mAb CC302 (5.5 µg ml–1; IAH, Compton, UK) per well was added. After an overnight incubation at 4 °C, plates were washed with PBS-T, and 100 µl Vectastain ABC peroxidase complex (Vector Laboratories) per well was added, following the manufacturer's instructions. After 90 min incubation at room temperature, the plates were washed with PBS-T and spots were developed by using AEC substrate (KemEnTec) following the manufacturer's instructions. Spots were counted and analysed with an AID ELISpot automatic reader using the AID ELISpot v2.5 software (Autoimmun Diagnostika GmbH).
Cultured IFN- ELIspot assay.
For cultured IFN- ELIspot assays, 1x104 or 5x104 CD8+ T cells per well were stimulated with autologous virus-infected APCs for 7 days in 24-well plates (TPP) and in the presence of 10 U recombinant human interleukin-2 (IL-2; Roche) ml–1 at an APC : CD8+ T-cell ratio of 3 : 1. CD8+ T cells were harvested and restimulated in vitro following the protocol described above. One set of three wells per plate was stimulated by adding 20 µl of the mitogen concanavalin A (1 µg ml–1; Sigma) to generate a positive control.
Data analysis.
INF- ELIspot results are shown as the mean±SD of triplicate spots per 106 CD8+ T cells. CD8+ T-cell responses are considered to be positive when the means are at least twice the negative control or more than 2SD higher than the negative control, whichever is highest. Calculation of descriptive statistics (geometric statistics and SD) and all statistical analyses (parametric analysis of variants for normally distributed data, followed by a Student–Neuman–Keuls test) were performed by using InStat (GraphPad).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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ELIspot assay was used to measure virus-specific IFN- release from CD8+ T cells. Six weeks after infection, we were unable to detect circulating antigen-specific CD8+ T cells by using the ex vivo assay (data not shown), but by using the cultured ELIspot assay, we were able to detect virus-specific memory CD8+ T-cell responses against the challenge virus (wild type), a tissue culture-adapted strain of FMDV (heparin binding) and a recombinant FPV expressing the P1 and 2A proteins of FMDV serotype O (Fig. 1). Only one of the six animals (VN90) did not show a statistically significant CD8+ T-cell response (P<0.005). The other five animals had statistically significantly higher responses than the control samples, ranging from 500 to 2500 spots per 106 CD8+ T cells. The responses of the individual animals to the three viruses used were not statistically significantly different, with the exception of VR57, where the response to the tissue culture-adapted FMDV was higher than the response to wild-type FMDV or rFPV-FMDV-P12A (P<0.005).
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) vaccinated with the commercially available vaccine against FMDV were tested for the presence of CD8+ T-cell activity directed against the structural proteins of the virus. An ex vivo IFN- ELIspot was performed to analyse the induction of FMDV P12A-specific CD8+ T cells in FMDV-vaccinated cattle by using the recombinant FPV expression system. Prior to vaccination, none of the seven animals were found to have a CD8+ T-cell response against FMDV and, 4 weeks after vaccination, there was still no significant response. Ex vivo CD8+ T-cell responses 7 days after boosting were statistically significant (P<0.05), albeit very low (>50 spots per 106 CD8+ T cells; data not shown).
Freshly isolated CD8+ T cells from cattle 12 weeks after boost were stimulated in vitro by using autologous APCs infected with rFPV-FMD-P12A and in the presence of IL-2. After a 7 day incubation, the CD8+ T cells were harvested and their activation was measured by IFN--release ELIspot. The response to FMDV P12A was increased up to 500-fold (Fig. 2), suggesting that there is a strong memory CD8+ T-cell response after vaccination. Of the seven animals, four had responses of between 500 and 1000 spots per 106 CD8+ cells and three had responses of between 2500 and 4500 spots per 106 CD8+ cells. All responses were statistically significantly different (P<0.005) from the responses to wild-type FPV or the negative controls.
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and 2.
Fig. 3 demonstrates clearly that the antigen-specific CD8+ T-cell responses in these FMDV-vaccinated animals are MHC-restricted. All four animals show a response to rFPV-FMD-P12A-infected P815 cells expressing at least one of their own expressed MHC class I alleles, and none of the four animals responded to P815 cells expressing irrelevant or no MHC class I alleles. FMD9, which expresses three alleles, responded to two of the three, FMD6 responded to one of two (it was not tested on the second), FMD17 responded to two of five and FMD18 responded to two of four (it was not tested on one).
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). The MHC class I haplotypes and associated alleles expressed by these animals are shown in Tables 1 and 2. In this case, the animals were tested for their ability to respond to MHC class I-transfected P815 cells infected with either wild-type FMDV or rFPV-FMD-P12A. Animal VR57 responded to two alleles when wild-type FMDV-infected P815 transfectants were used, but only to one of these when rFPV-FMD-P12A-infected P815 transfectants were used, which may reflect recognition of an epitope(s) derived from a different protein in the former case. Neither animal responded to P815 cells expressing irrelevant or no MHC class I alleles.
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). Both vaccinated animals responded to autologous infected cells and to cells expressing one of their haplotypes (A14 in the case of FMD17 and A10 in the case of FMD18). However, both animals failed to respond to cells expressing their other haplotype (A31 in both cases). This is in contrast to the result shown in Fig. 3, where both animals showed a low response to P815 cells expressing N*02201, one of two alleles expressed on the A31 haplotype; in this case, the APCs were infected with rFPV-FMD-P12A rather than wild-type FMDV. Both infected animals responded to autologous infected cells and to cells expressing shared haplotypes. None of the animals responded to infected, MHC-mismatched cells.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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MHC class I-restricted virus-specific CD8+ T cells can mediate antiviral protection via two pathways: direct cytotoxicity or release of cytokines such as IFN-. Due to the highly cytopathic nature of the virus, it has never been possible to establish a classical cytotoxic T lymphocyte (CTL) assay for FMDV, and we have therefore employed an IFN--release ELIspot assay to measure CD8+ T-cell activity. We show that a CD8+ T-cell immune response is present following vaccination and that the CD8+ T-cell response is almost 100 times higher following restimulation. Conversely, we could not detect a CD8+ T-cell response following infection, although a memory response was detectable and measurable. The level of response varied greatly between animals in both the vaccinated and infected groups, with one animal in the latter group failing to respond.
An MHC class I-restricted CD8+ T-cell response is generally induced by endogenously synthesized antigens, such as those generated during viral infections, whilst induction of a class II response is mediated by exogenous soluble proteins. Consequently, an infectious virus that can replicate in the host is the optimum vaccine candidate for priming a CD8+ T-cell response. By using a recombinant FPV expressing the structural polyprotein of FMDV (P12A), we were able to stimulate CD8+ β memory T cells in animals vaccinated with chemically inactivated virus. This result is not unprecedented, and similar observations have been reported using a CD8+ T-cell clone derived from FMDV-vaccinated pigs (Rodriguez et al., 1996). The induction of pathogen-specific CD8+ CTLs after vaccination with killed virus has been reported in other systems, such as hepatitis B virus and influenza A virus (Bohm et al., 1995; Mbawuike & Wyde, 1993; Schirmbeck et al., 1995).
The commercial vaccine used in this study relies on chemical inactivation of highly purified virions grown in tissue culture, essentially creating replication-incompetent virions similar to virus-like particles (VLPs). The vaccine formulation is then prepared by using a proprietary adjuvant into water-in-oil emulsion. Whilst it is conceivable that the adjuvant used could be involved in priming the observed CD8+ T-cell immune response, it is more likely to be a function of FMDV VLPs. Thus, non-replicating papillomavirus particles are capable of efficient induction of CTL responses in the absence of adjuvants in the mouse model (Sedlik et al., 1997, 1999). The mechanism(s) responsible for this VLP-induced CD8+ T-cell priming is unclear, although it has been proposed that FcR targets VLPs to dendritic cells, inducing a maturation state that facilitates antigen presentation to naïve T cells (Da Silva et al., 2007).
MHC restriction of IFN- production was demonstrated in both infected and vaccinated animals by using target cells from animals matched, half-matched or mismatched for MHC, and P815 cells transfected with single MHC class I alleles. Results show that animals responded consistently only to cells expressing either a complete shared haplotype or individual shared alleles, and generally responded preferentially through the same haplotypes/alleles. There were some anomalies between the results obtained by using rFPV-FMD-P12A and wild-type FMDV, and between results obtained by using half-matched targets and targets expressing single alleles. Specifically, CD8+ T cells from animal VR57 recognized peptides presented by the allele N*00201 when expressed on P815 cells infected with wild-type FMDV, but not when infected with rFPV-FMD-P12A. This suggests the presence of N*00201-restricted epitopes within proteins other than FMDV P12A. Vaccinated animals FMD17 and FMD18 both failed to recognize the A31 haplotype on half-matched targets infected with wild-type FMDV, but showed a small response to one of the A31 alleles (N*02201) when expressed on P815 cells together with FMDV P12A (Fig. 3). One possible explanation for this is that, on the half-matched targets, other expressed alleles compete more efficiently for peptides or, alternatively, the N*02201 allele binds preferentially to peptides derived from FMDV proteins other than P12A that are not CD8+ T-cell epitopes.
The allele N*01301 (the only allele expressed on the A18 haplotype) generated the most consistently high response. This is interesting because, in previous studies of T-cell responses to the protozoan parasite Theileria parva, most animals studied showed a CTL response restricted entirely by the products expressed by one of their MHC class I haplotypes, and the broad ‘w6’ serological specificity was often dominant (Morrison, 1996). This specificity encompasses a number of related haplotypes, one of which is A18, as used in the current study.
Our results confirm the presence of circulating effector and memory MHC class I-restricted CD8+ T cells specific for FMDV in the natural host; however, the correlation between FMDV-specific CD8+ T-cell recognition and protection against disease remains to be defined. It is possible that the CD8+ T-cell function is not confined to MHC-restricted cytotoxicity. For example, during FMDV infection, secretion of IFN- may be important in controlling the virus, at least in some animals. Therefore, an analysis of lymphokine profiles would be valuable in determining the roles of the different T-cell subsets during the immune response to FMDV. Our future studies are directed towards identifying specific CD8+ T-cell epitopes in the structural and non-structural regions of the virus genome and to a better understanding of the role of genetic variability in protection against FMDV.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Received 31 August 2007;
accepted 19 November 2007.
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