Institute for Animal Health, Pirbright Laboratory, Woking, Surrey GU24 0NF, UK
Correspondence
Peter Mertens
peter.mertens{at}bbsrc.ac.uk
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ABSTRACT |
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Present address: Novartis Horsham Research Centre, Wimblehurst Road, Horsham, West Sussex RH12 4AB, UK.
Present address: Instituto Gulbenkian de Ciencia, Aportado 14, 2781 Oeiras Codex, Portugal.
Published ahead of print on 15 October 2002 as DOI 10.1099/vir.0.18705-0.
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Introduction |
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AHSV has also survived in Spain from 1987 to 1990, where the silent periods ranged from 8 to 12 months (Mellor, 1994a). Although very small numbers of adult vector insects were present in some locations in southern Spain during these inter-epizootic periods, no disease or evidence of seroconversion was seen in any equids, despite intensive surveillance by the national veterinary authorities (Mellor, 1993
; Mellor & Boorman, 1995
). These observations strongly suggest that transmission of AHSV did not occur during these inter-epizootic periods.
This pattern of annual episodes of disease in the late summer or autumn, separated by periods of quiescence, is a feature common to many orbivirus diseases that are transmitted by Culicoides. Reappearance of the disease coincides with major increases in the abundance of adults of the vector species (Mellor, 1996). The survival of virus from one vector season to the next is popularly termed overwintering (Yonguc et al., 1982
; Taylor & Mellor, 1994
; Mellor & Boorman, 1980
; Mellor, 1998
). It is evident that a reservoir of the virus must exist through the winter but its location and the mechanism by which it re-establishes overt infection of either the vector insect or the mammalian host has not been satisfactorily explained or demonstrated.
Here, we demonstrate that ovine T-cells that are persistently infected with BTV can be converted to lytic infection through interaction with skin fibroblasts or an antibody to the
T-cell-restricted surface protein WC-1. On the basis of these observations we suggest a novel mechanism by which overwintering of BTV could occur in the absence of adult vector insects.
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Methods |
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Animals.
Eight 6-month-old female Dorset Horn sheep were obtained from the Institute for Animal Health (Compton Laboratory, Reading, UK) and were held in insect-proof, high-security accommodation at the Pirbright Laboratory (Surrey, UK) throughout the experiments. At 18 days prior to BTV infection, six of the sheep were exposed to the bites of Culicoides midges (see below). Four of these and the two untreated control sheep were inoculated intradermally with 106 TCID50 of BTV-1 (Takamatsu & Jeggo, 1989). The two remaining midge-bitten sheep were used as uninfected controls. Viraemia was monitored by virus isolation from peripheral blood (see below). At 9 weeks post-infection (p.i.), the sheep were exposed to feeding midges (400 midges per animal per exposure) three times within a 24 h period (7:00 am, 3:00 pm and 7:00 am). Peripheral blood, skin biopsy tissues, fed midges, tissue fluid and blood from skin biopsy sites were all collected at 0, 8, 24 and 32 h after the first midge feeding point. All animal experiments were carried out humanely and in accordance with the requirements of the Animal (Science Procedure) Act 1986, under project licence number PPL90/00866. Isolations of BTV from these samples were performed as described below.
Insects.
Adult C. sonorensis (previously named C. variipennis) (a known vector of BTV) from a colony maintained at IAH Pirbright (Boorman, 1974) were used in these experiments.
Virus isolation.
Heparinized whole blood, homogenized skin biopsy samples, tissue fluids and blood collected from skin biopsy sites were inoculated into BHK cell cultures and chick embryos (Mellor et al., 1984) in attempts to isolate BTV. Similarly, the supernatants from skin biopsy tissue cells (collected after culturing for 1 week at 37 °C) and lymphocyte cultures (established from tissue fluids and bloods recovered from skin biopsy sites and cultured with IL-2 for 1 week) were also assayed for BTV. Midges that had been fed on infected sheep were incubated for 10 days at 25 °C and were then homogenized in ground glass grinders before assay for infectious virus (Mellor et al., 1984
).
Cell lines and cultures.
Ovine (S-59 and S-96) T-cell lines, a bovine (Th-67)
T-cell line (Takamatsu et al., 1997
), a short-term cultured ovine (S-40)
T-cell line [
T-cell receptor (TCR)+, WC1+, CD2- and CD8-1], a bovine (TC-6) CD4+
T-cell line (Takamatsu et al., 1990
) and an ovine (TS-47)
T-cell line (
TCR-, WC-1-, CD2+ and CD4+) were used in these experiments. All of the T-cell lines used were IL-2 dependent and were cultured with Iscove's modified MEM (IMEM) supplemented with recombinant human IL-2 (10 U ml-1), 10 % heat-inactivated FCS and antibiotics, as described previously (Takamatsu et al., 1997
). Primary sheep skin fibroblast cell lines (skin 59, skin 59-3, skin 73, skin 82, skin 83, skin 85 and skin 96) were prepared from skin biopsy samples and were maintained as described previously (Takamatsu & Jeggo, 1989
). A sheep endothelial cell line was prepared from mechanically harvested large vessel endothelium from sheep aorta and cultured as described by Ryan et al. (1978)
. The
T-cell line S-59, as well as skin fibroblast cell lines skin 59 and skin 59-3, were all derived from the same animal.
The T-cells from BTV-infected sheep were cultured as follows: peripheral blood lymphocytes (PBLs) were isolated from heparinized blood by gradient centrifugation on Nycoprep Animal (Nycomed) and plastic-adherent cells were removed by repeated incubation at 37 °C for 30 min in plastic tissue culture flasks. Non-adherent PBLs were then incubated with anti-sheep CD2 (mAb 36F, IgG2a), anti-sheep CD8 (mAb 7C2, IgG2a) and polyclonal rabbit anti-sheep immunoglobulin (Dacopatts) at 4 °C for 30 min. After washing twice with IMEM, the antibody-coated
T-cells and B-cells were lysed by rabbit complement (Cedarlene Laboratories) by incubating at 37 °C for 30 min. Live cells were isolated by gradient centrifugation on Nycoprep Animal and cultured with IL-2-containing IMEM. The purity of the
T-cells in the cultures was assessed by FACS analysis (see below).
FACS analysis.
FACS analysis was performed with FACScan (Becton Dickinson) using the LYSIS II program and the following mouse mAbs: anti-bovine WC-1, which cross-reacts with sheep; mAb CC 15 (IgG2a), anti-sheep WC-1; mAb SC-29 (IgG1), anti-sheep TCR; mAb 86D (IgG1), anti-sheep CD4; mAb 17D (IgG1), anti-sheep CD8; mAb 7C2 (IgG2a), anti-sheep CD45; mAb 1D (IgG1) and goat anti-sheep Ig conjugated with FITC. Secondary antibodies, conjugated with either FITC or R-phycoerythrin (PE), were obtained from Southern Biotechnology. For BTV antigen staining, cells were fixed with 1 % paraformaldehyde, permeabilized with 0·3 % saponin/PBS (containing 0·25 % BSA) and stained with mAb 3-17-3A (mouse anti-BTV VP7) in 0·3 % saponin/PBS, followed by anti-mouse PE-conjugated immunoglobulin in saponin/PBS. mAbs 124 (anti-human Bcl-2, which cross-reacts with sheep cells) and ILA-30 (anti-bovine IgM, which does not cross-react with sheep cells) were used as positive and negative controls, respectively.
Immunohistology.
Skin biopsy samples were frozen in Tissue-Tek OCT (Miles) by floating in isopropanol cooled with dry ice. Cryostat sections (5 µm) were prepared using a Jung Frigocut cryostat (ReichertJung) and standard immunochemical staining techniques were used following the protocol of the Vectastain ABC kit (Vector Laboratories).
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Results |
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A total of seven sheep skin fibroblast cell lines and a sheep endothelial cell line were also examined for their ability to induce growth arrest of the T-cell line S-59. Three of the skin fibroblast lines [skin 82 (Fig. 3
A), skin 59-3 and skin 96 (data not shown)] did induce growth arrest, a process that was therefore not MHC-restricted. However, two of these cell lines (skin 59-3 and skin 96) later lost this ability. The absence of growth arrest by two further cell lines (skin 59 and skin 73) provides MHC-matched and MHC-mismatched negative controls for the induction of growth arrest by skin 82 (Fig. 3A
).
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Six sheep were infected with BTV by intradermal inoculation. All of them developed mild but typical clinical signs of bluetongue, including fever, lymphopenia and swollen muzzles. Viraemia was detected in each case from 4 or 5 days until 21 to 28 days p.i. The T-cells recovered from peripheral blood samples taken on days 313 p.i. from each of the infected sheep were shown to be infected with BTV. BTV was rarely detected in the freshly isolated and purified
T-cells but the virus was readily recovered when the cells were cultured with IL-2. Few, if any,
T-cells were recovered from blood samples taken between 14 and 34 days p.i. due to lymphopenia, which reduced very significantly the number of these cells in circulation. Virus was not recovered from the
T-cells that were isolated from peripheral blood samples taken between 35 and 63 days p.i., even when they were cultured with IL-2.
The T-cells isolated from the blood of BTV-infected sheep (313 days p.i.) were cultured for up to 6 weeks and it was confirmed by recovery of infectious virus that they were persistently infected. At 4 weeks into the culturing period, five of the
T-cell cultures were co-cultured with
-irradiated skin 82 cells. As observed previously with established ovine
T-cell lines, this treatment resulted in conversion to a lytic form of BTV infection. The treatment also increased the amount of virus recovered from these cultures (Table 1
), although the degree of enhancement (between log 0·2 and 2·6) was less dramatic than that observed with the established S-59 cell line (Fig. 3B
).
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Severe skin inflammation was caused by feeding Culicoides midges on the sheep, which can be seen in Fig. 4(A), extending across the whole diameter (6 cm) of the cup used previously to keep the insects in place. In addition, immunohistological studies of skin biopsy tissues taken at 8 h post-feeding from three of the midge-sensitized and two untreated sheep demonstrated a massive accumulation of
T-cells in the midge feeding areas (more than 350
T-cells per field at xtimes;400 magnification) (Fig. 4B
). In samples from untreated sheep, or from non-inflamed areas of the sensitized sheep, an average of 2030
T-cells per field were detected. These cells were spread evenly and not clustered as in the inflamed tissue samples. Both CD4+ (120210 cells per field at xtimes;400 magnification) and CD8+ T-cells (6577 cells per field at xtimes;400 magnification) were observed in samples from midge feeding areas but in smaller numbers than
T-cells (Fig. 4C, D
).
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Discussion |
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One possible overwintering mechanism could involve survival of infected adult vector insects. The lifespan of an adult midge is usually less than 10 days. However, in exceptional conditions, some individuals can survive for several weeks, but there is no evidence to suggest that they could survive for an entire intra-epizootic period of 912 months (Mellor, 1990). Survival of the virus in persistently infected, long-lived adult Culicoides is therefore considered to be very unlikely indeed. Culicoides midges usually survive winter periods as larvae but there is no evidence of transovarial transmission of either BTV or AHSV in these insects. Persistence of virus in larvae is therefore also considered to be highly improbable (Mellor, 1990
).
The re-introduction of either infected adult midges or viraemic vertebrate hosts from other enzootic areas could also give the appearance of overwintering. Although this possibility is often difficult to exclude with certainty, in at least some cases it cannot be reconciled with epidemiological data. For example, during the outbreak of African horse sickness (AHS) in Spain in 19871990, annual disease episodes were each caused by AHSV-4, a serotype that had never been recorded previously outside southern Africa. At the time of the outbreaks there was no other evidence of AHSV, of any serotype, within 2000 miles of Spain (Mellor & Boorman, 1995). Under these circumstances, the possibility of annual re-introduction of AHSV-4 over four consecutive years is remote in the extreme. Precisely the same situation prevailed during the BTV-10 outbreaks in Spain and Portugal from 1955 to 1960, the BTV-4 outbreaks in western Turkey from 1977 to 1981 and the current outbreaks of BTV-2, -4, -9 and -16 in the Mediterranean region. In each case there was an initial introduction of virus to a location that was geographically remote from the nearest enzootic zones, followed by recrudescences of the same virus type(s) over a period of years.
A third possibility is that an unknown vector or vertebrate host species could be involved, providing a natural reservoir in which the virus could persist. This is also most unlikely, as the winter conditions that result in the absence of known BTV and AHSV vector species of Culicoides will have precisely the same effect on other less abundant and less efficient vector species of Culicoides and other biting insects (Ortega et al., 1998). Attempts to infect a wide range of animals, including mice, rats, hamsters, guinea pigs, rabbits, ferrets, dogs, other carnivores, camel and elephant, have been carried out with either AHSV and/or BTV. In all cases these species are not considered to play a significant role in the epidemiology of either disease (Coetzer & Erasmus, 1994
; Alexander et al., 1994
; Akita et al., 1994
). In the absence of an identifiable overwintering reservoir, an alternative vector or mammalian host species, we have considered the possibility that orbiviruses might persist via some unidentified mechanism in their usual vertebrate hosts.
The experiments described here demonstrate that infectious BTV can be recovered from ovine skin biopsies for more than 9 weeks p.i. (i.e. for the entire duration of the experiment), although the viraemic phase had ended after only 34 weeks. It appears likely that the source of the virus is T-cells that became persistently infected during the initial stages of virus replication and viraemia in the host. BTV association with leukocytes and virus replication in lymphocytes has been demonstrated previously by many authors (Morrill & McConnell, 1985
; Whetter et al., 1989
; Stott et al., 1990
). Stott et al. (1990
, 1992
) also observed a persistent BTV infection in bovine CD8+ T-cells and in null cells in vitro. The null cells were so described due to an absence of expression of CD2, CD4, CD6, CD8 or WC-1 molecules, as recognized by mAb IL-A29. Although the authors started with
T-cell-enriched cultures, IL-A29-positive cells were gradually lost from the culture (Stott et al., 1992
). Activated
T-cells isolated from ruminants are not only negative for CD2, CD4 and CD8 expression, they also show low or zero expression of CD6 and loss of the WC-1 epitope recognized by mAb T-19 or IL-A29 (Lund et al., 1993
; H. Takamatsu unpublished data). However, they do show increased expression of the WC-1 epitope recognized by mAb SC-29 (H. Takamatsu, unpublished data). The null cells identified by Stott et al. (1992)
are therefore likely to be activated
T-cells. In this study we have confirmed persistent BTV infection of IL-2-dependent ovine and bovine
T-cells cultures in vitro. These BTV-infected
T-cell cultures grow continuously, with no obvious CPE. A similar persistent infection of BTV has been demonstrated in some insect cell cultures without shut-off of host cell DNA or protein synthesis (Jennings & Boorman, 1979
; Wechsler et al., 1989
; Fu et al., 1999
) where there was also little or no sign of CPE.
BTV and AHSV possess many characteristics in common with other members of the family Reoviridae and the model for BTV replication is partly based on orthoreovirus replication (Eaton et al., 1990). Orthoreovirus non-structural protein
1 is essential for cell lysis and CPE (reviewed by Oberhause et al., 1998
). The protein
1 inhibits host cell DNA synthesis and blocks the host cell from progressing to the S phase so that they remain in the G1 phase. BTV infection also causes host cell shut-off in lytically infected mammalian cell lines, including BHK cells, but does not induce shut-off in cultured
T-cells. It was therefore considered possible that anti-WC-1 antibodies, which also cause growth arrest at the G1 phase (Takamatsu et al., 1997
; Kirkham et al., 1997
, 1998
), might mimic the processes of host cell shut-down that occur in lytically BTV-infected mammalian cells, thereby causing the conversion to a lytic mechanism, and indeed, this proved to be so. The same conversion from persistent to lytic BTV infection could also be induced in BTV persistent
T-cells by co-culture with some, but not all, skin fibroblast cell lines, incidentally suggesting that the latter bear a WC-1 counter receptor.
If persistent infection of T-cells in vivo plays a central role in a BTV overwintering mechanism, then it appears likely that the conversion of persistence to the more productive lytic form of infection would occur at a location and at a time when midges interact with the mammalian host. Recruitment of activated sheep
T-cells to inflamed areas of the skin has been observed previously (Bowles et al., 1992
; Egan et al., 1996
). Biting by Culicoides midges has been shown to induce a severe inflammatory response in equid and bovine skin (Mellor & McCaig, 1974
; P. S. Mellor & M. H. Jeggo, unpublished data). A similar response was also observed in the sheep used in the study described here. Interactions between skin fibroblasts and the WC-1 molecules on the surface of
T-cells may also result in a similar shut-down of those activated
T-cells that become involved in lymphoid infiltration to the site of vector biting. If these cells were already persistently infected with BTV, the data presented here indicate that this would be accompanied by conversion to a lytic infection and enhanced virus replication and release. In practice, the release of BTV particles from persistently infected
T-cells might be restricted. Low levels of virus production might be masked by the presence of neutralizing antibodies, until sufficient biting insects were present in the environment to induce a sufficiently intense dermal inflammatory response to trigger the release of infectious virus at the site of inflammation and consequently, of insect biting.
In addition, proteases, which are associated with such an inflammatory response (Harvima et al., 1994), may also play an important role in transmission. Treatment of BTV or AHSV particles with proteases results in cleavage of the outer capsid protein VP2, forming infectious subvirus particles (ISVPs) (Marchi et al., 1995
, Mertens et al., 1987
, 1996
; Burroughs et al., 1994
). These ISVPs have a specific infectivity for insect cells or adult Culicoides of
100 times greater than that of intact virus particles and it is considered likely that infection of vector insects is by these protease-modified particles (Mertens et al., 1996
).
A vital component of the proposed BTV overwintering mechanism is the requirement for persistent or latent infection of T-cells. At this moment we do not know how persisting BTV escapes detection during immune surveillance. One of the possibilities is that the low level of virus replication in resting lymphocytes (Stott et al., 1992
) could result in little or no production of potential cytotoxic T-cell epitopes. Thus BTV-infected, resting
T-cells may remain undetected during immune surveillance or attempts at virus isolation. These observations may explain why, in the present study, virus was only recovered when lymphocytes were cultured in vitro with IL-2, thus activating
T-cells. Confirmation of this proposed BTV overwintering mechanism requires the demonstration of infected ovine
T-cells in vivo for periods of up to 912 months. In relation to this, it is well known that BTV infection in cattle is clinically milder than in sheep and the viraemic period is longer, so that the virus may be more closely adapted to bovines than ovines. Thus, cattle may be a more appropriate mammalian host species in which to study BTV persistence further. Indeed, although previous reports of latent-infection and showering phenomenon after insect bites in cattle (Luedke et al., 1977
) are now largely disregarded, they might be at least partially explained by persistent BTV infection of bovine
T-cells.
In the present bluetongue epizootic in Europe and North Africa (19982001), penetration of the virus further north than ever before has occurred, particularly in eastern Europe, so that regions of Bulgaria, Serbia and Kosovo have become affected (Mellor & Wittmann, 2002). Astonishingly, the virus also appears to have succeeded in overwintering in this northerly region where adult vectors are absent for several months each year. In these locations, the traditional vector, C. imicola, does not occur and a novel vector species with a distribution that extends across central and northern Europe is operating (Mellor & Wittmann, 2002
). Should the mechanism outlined in the present study prove to be the method by which BTV is overwintering in the region, then the (seasonal) presence of a more northerly vector could provide the virus with the means to extend its range much further northwards and much more widely across Europe.
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ACKNOWLEDGEMENTS |
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Received 10 July 2002;
accepted 7 October 2002.