Evaluation of lumpy skin disease virus, a capripoxvirus, as a replication-deficient vaccine vector

Kate Aspden1, Jo-Ann Passmore1, Friedrich Tiedt1 and Anna-Lise Williamson1,2

1 Division of Medical Virology, Department of Clinical Laboratory Science & Institute of Infectious Disease and Molecular Medicine, University of Cape Town, Observatory 7925, Cape Town, South Africa
2 National Health Laboratory Service, University of Cape Town, Observatory 7925, Cape Town, South Africa

Correspondence
Anna-Lise Williamson (at Division of Medical Virology)
annalise{at}curie.uct.ac.za


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Lumpy skin disease virus (LSDV), a capripoxvirus with a host range limited to ruminants, was evaluated as a replication-deficient vaccine vector for use in non-ruminant hosts. By using the rabies virus glycoprotein (RG) as a model antigen, it was demonstrated that recombinant LSDV encoding the rabies glycoprotein (rLSDV-RG) was able to express RG in both permissive (ruminant) and non-permissive (non-ruminant) cells. The recombinant LSDV, however, replicated to maturity only in permissive but not in non-permissive cells. Recombinant LSDV-RG was assessed for its ability to generate immunity against RG in non-ruminant hosts (rabbits and mice). Rabbits inoculated with rLSDV-RG produced rabies virus (RV) neutralizing antibodies at levels twofold higher than those reported by the WHO to be protective. BALB/c mice immunized with rLSDV-RG elicited levels of RV-specific cellular immunity (T-cell proliferation) comparable with those of mice immunized with a commercial inactivated rabies vaccine (Verorab; Pasteur Merieux). Most importantly, mice immunized with rLSDV-RG were protected from an aggressive intracranial rabies virus challenge.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Poxviruses have been widely investigated as vaccine vectors because they activate both humoral and cellular immunity (Zavala et al., 2001; Willey et al., 2003), have the capacity to accommodate over 25 kb of extra DNA (Smith & Moss, 1983; Merchlinsky & Moss, 1992) and simultaneous expression of several foreign genes has been achieved (Perkus et al., 1985; Carroll et al., 1998; Welter et al., 2000). Vaccinia virus (VV), in particular, has been used extensively because of its well-defined molecular characteristics (Moss, 2001) and its success in the WHO vaccine programme to eradicate smallpox (WHO, 1980). It is not considered absolutely safe, however, since virus replication is not host-restricted and disseminated infections are a risk to immune-compromised individuals (Redfield et al., 1987). As a result, a number of replication-deficient and/or host-restricted poxviruses have been investigated. These include highly attenuated strains of VV (Rodriguez et al., 1989; Moss et al., 1996) and the host-restricted avipoxviruses (Taylor et al., 1988, 1991; Cadoz et al., 1992; Somogyi et al., 1993; Stannard et al., 1998).

Modified vaccinia virus Ankara (MVA) was attenuated by more than 570 passages of vaccinia virus (strain Ankara) in chicken embryo fibroblasts (Mayr & Danner, 1978), during which multiple deletions and mutations occurred that severely restricted the virus's host range (Wyatt et al., 1998; Meyer et al., 1991). Complete genomic sequence analysis of MVA in comparison with other orthopoxviruses confirmed that the genome of MVA is ~14 kb smaller and that 25/177 MVA-encoded genes are either split and/or mutated resulting in truncated proteins (Antoine et al., 1998). MVA was found to be non-pathogenic in animals and replication-deficient in mammalian cells. Experiments have indicated that recombinant MVA viruses can provide protection against a wide variety of viral pathogens. Where comparisons were made, the immunogenicity of MVA recombinants was equal to or better than that of VV recombinants (Sutter et al., 1994; Bender et al., 1996; Hirsch et al., 1996; Wyatt et al., 1996). The efficacy of this vector, however, may be limited by pre-existing immunity to other VV strains (Baxby & Paoletti, 1992).

The avipoxviruses were initially investigated and developed as vaccine vectors for immunizing birds against avian diseases. Fowlpoxvirus (Boyle & Coupar, 1988) and canarypoxvirus (commercially known as ALVAC; Taylor et al., 1988) have also been investigated as safe and efficient vaccine vectors for use in humans and other mammals. Although the efficacy of canarypox-based vaccines has been demonstrated in pre-clinical studies for a number of recombinant antigens (Taylor et al., 1991; Cadoz et al., 1992; Paoletti, 1996), this vector has only demonstrated limited immunogenicity to recombinant antigen in phase I/II clinical trials (Belshe et al., 2001; Gupta et al., 2002).

In this study, lumpy skin disease virus (LSDV), a capripoxvirus with a host range limited to ruminants (Alexander et al., 1957), was evaluated as a replication-deficient vaccine vector for use in non-ruminant hosts, using the rabies virus glycoprotein (RG) as a model recombinant antigen. Two possible insertion sites have been identified in the LSDV genome for the insertion of foreign genes: the ribonucleotide reductase gene and an intergenic region (Cohen et al., 1997). Recently, the complete genome sequences of several capripoxviruses, including LSDV (Tulman et al., 2001), sheeppoxvirus and goatpoxvirus (Tulman et al., 2002), have been published, revealing other potential sites for insertion of foreign genes. LSDV has already been used in ruminants to express recombinant antigens and has been shown to be a successful dual vaccine in cattle against both the recombinant antigen and LSDV. We have previously demonstrated that recombinant LSDV (Neethling strain) expressing RG (rLSDV-RG) elicits both neutralizing antibodies and cell-mediated immunity against rabies virus (RV) in cattle (Aspden et al., 2002). Romero et al. (1993, 1994a, b) reported on an LSDV–rinderpest recombinant vaccine that protected cattle against both LSDV and rinderpest.

To investigate host-restricted LSDV as a vaccine vector in non-ruminants, we have evaluated the morphogenesis of rLSDV-RG (Neethling strain) in cells of ruminant as well as non-ruminant origin by electron microscopy. We found that rLSDV-RG only replicates to maturity in permissive Madin–Darby bovine kidney (MDBK) cells but immature virions were clearly evident in non-permissive cells. Despite limited replication in non-permissive cells, we were able to demonstrate expression of RG by rLSDV-RG in both permissive and non-permissive cells. Furthermore, rLSDV-RG was shown to elicit an RV neutralizing antibody response, an RG-specific cell-mediated immune response and protect against intracranial RV challenge in non-ruminant hosts.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Generation of rLSDV-RG.
LSDV (Neethling strain) was kindly supplied by H. G. Jaeger of Onderstepoort Biological Products (Onderstepoort, South Africa). The virus was passaged through primary lamb testes (LT) cells to obtain a titre of 3x106 focus-forming units (f.f.u.) ml-1 (Aspden, 2002). The ribonucleotide reductase gene was ascertained to be non-essential for replication of LSDV by the creation of a LSDV gene library and experimental insertional inactivation (Cohen et al., 1997). Recombinant LSDV (Neethling strain) expressing RG under the control of a fowlpoxvirus early/late promoter was constructed using a shuttle vector that targeted the RG gene into the non-essential ribonucleotide reductase gene in the LSDV genome according to the method recently described by Aspden et al. (2002). The recombinant LSDV-RG was passaged through LT cells eight times with selection (Aspden et al., 2002). The cells and growth medium were harvested, frozen and thawed three times and centrifuged for 20 min at 2000 r.p.m. to remove cell debris. The supernatant was centrifuged for 2 h at 19 000 g through a 36 % sucrose solution to purify the recombinant virus. The rLSDV-RG pellet was resuspended in FCS-free Dulbecco's modified Eagle's medium (DMEM) (1x106 f.f.u. ml-1) and frozen at -70 °C in 500 µl aliquots.

Assessment of transient gene expression.
The ability of rLSDV-RG to express foreign genes under the control of different promoters was assessed in a transient expression system using plasmids in which the reporter gene lacZ was under transcriptional control of the VV promoters P11 (late) on plasmid PAL1 (supplied by M. Mackett, Paterson Laboratories) and P7.5 (early/late) on plasmid PSC65 (a VV shuttle vector). Non-permissive CV-1 (African green monkey kidney cell line) or permissive MDBK and LT cell monolayers were infected with rLSDV-RG at an m.o.i of 5 for 1 h. Virus inoculum was removed and cells transfected with 5 µg plasmid using liposomes (DOTAP; Roche Diagnostics) according to the manufacturer's instructions. At 24, 30 and 48 h post-infection, culture medium was removed and cells covered with PBS containing X-Gal (1 mg ml-1). After 18 h, {beta}-galactosidase activity was determined by counting the number of blue cells (MacGregor et al., 1991).

Electron microscopy of rLSDV-RG-infected cells.
Duplicate cultures of CV-1, MDBK and LT cells were grown overnight to 70 % confluence in 5 ml culture flasks before being infected with rLSDV-RG. Each culture flask was infected with 1 f.f.u. per cell of recombinant virus and grown for another 72 h. Cells were detached using a rubber policeman, centrifuged at 250 g for 10 min and washed with PBS. The pellets were stabilized in agarose and fixed in glutaraldehyde. Ultra-thin sections were prepared from rLSDV-RG-infected CV-1 cells, MDBK cells and LT cells harvested at 48 h post-infection (p.i.). Cell cultures were rinsed three times with PBS, fixed in 2 % glutaraldehyde for 2 h at 4 °C, post-fixed in 1 % OsO4, then dehydrated and embedded in Spurr's resin by conventional methods. Ultra-thin sections were post-stained with uranyl acetate and lead citrate before examination with an electron microscope.

Assessment of infection efficiency and kinetics using X-Gal staining.
Triplicate cultures of CV-1, 3T3 (murine fibroblast cell line), MDBK and LT cells were grown overnight to 70 % confluence in 24-well tissue culture plates (Nunc) before being infected with rLSDV-RG. Each well was infected with 2·5x103 f.f.u. ml-1 of recombinant virus for 2 h in 10 % FCS DMEM, after which the medium was removed and 10 % FCS DMEM containing mycophenolic acid, xanthine and hypoxanthine (Aspden et al., 2002) was added. At 24, 48 and 72 h p.i., medium was removed, the cells washed once with PBS, fixed for 10 min with 4 % paraformaldehyde, washed with PBS and then stained with X-Gal stain according to the method described by MacGregor et al. (1991). The number of LT, MDBK, CV-1 and 3T3 cells infected with rLSDV-RG (i.e. expressing the {beta}-galactosidase reporter gene product) was evaluated by counting at least 200 cells in each of the triplicate culture wells. To evaluate the number of infected cells that were either present as single cells or present as foci (defined as >2 adjacent cells staining positive by X-Gal staining), at least 100 infected cells were counted.

Quantification of {beta}-galactosidase activity.
{beta}-Galactosidase reporter gene activity was determined using a commercial kit provided by Sigma. Briefly, triplicate cultures of CV-1, 3T3, MDBK and LT cells were grown overnight to 70 % confluence in six-well tissue culture plates (Nunc) before being infected with rLSDV-RG. Each well was infected with 2·5x103 f.f.u. recombinant virus ml-1 for 2 h in 10 % FCS DMEM, after which the medium was removed and 10 % FCS DMEM containing mycophenolic acid, xanthine and hypoxanthine (Aspden et al., 2002) was added. At 72 h p.i., medium was removed and cells washed three times with PBS. Cells were lysed using lysis buffer (25 mM CHAPS, 250 mM HEPES, pH 7·5) for 15 min at room temperature. Cell lysate was collected and centrifuged at 5000 r.p.m. for 5 min to remove cell debris. Infected cell lysate (150 µl) was added to an equal volume of 2x assay buffer (1·33 mg ONPG substrate ml-1, 2 mM MgCl2, 100 mM mercaptoethanol, 200 mM sodium phosphate, pH 7·3). This was incubated at 37 °C for 30 min and then stopped by the addition of 500 µl 1 M sodium carbonate stopping solution. The absorbance (A) was read at 420 nm and {beta}-galactosidase activity calculated using the following equation: units per sample=(Axfinal reaction volume)/(4·6xtmin). To normalize {beta}-galactosidase activity to infection efficiency, the results were transformed using the following equation: units per % cells infected=(Axfinal reaction volume)/(4·6xtmin) per % cells infected with rLSDV-RG (i.e. expressing {beta}-galactosidase).

Immunofluorescence of rLSDV-RG-infected cells.
CV-1, 3T3, MDBK and LT cells were grown overnight to 70 % confluence in eight-well chamber slides (Nunc) before being infected with rLSDV-RG. Each well was infected with 2·0x103 f.f.u. recombinant virus ml-1 for 2 h in 10 % FCS DMEM, after which the medium was removed and 10 % FCS DMEM containing mycophenolic acid, xanthine and hypoxanthine (Aspden et al., 2002) was added. Controls on each slide included one uninfected well and one well infected with LSDV-wt (2x103 f.f.u. ml-1). At 72 h p.i., medium was removed and cells washed twice with PBS. The cells were fixed with acetone at 4 °C for 10 min, air dried and then washed with PBS for 10 min at room temperature with shaking. Ovalbumin (2 %) was added as blocking reagent for 20 min at room temperature with shaking, then the wells were washed twice with PBS for 10 min each at room temperature with shaking. A 1 : 70 dilution of unlabelled anti-rabies glycoprotein monoclonal antibody (Biodesign, ME) containing 1·5 % BSA in PBS was added and the cells incubated for 1 h at 37 °C. The cells were washed twice with PBS for 10 min with shaking. A 1 : 10 dilution of FITC-conjugated rabbit anti-mouse IgG (Dako) containing 1·5 % BSA in PBS was added and the cells incubated for 20 min at 37 °C. The cells were washed twice with PBS for 10 min with shaking, rinsed in distilled water and mounted using Dako fluorescent mounting medium. Immunofluorescence was evaluated using a Zeiss Axioscop 2 fluorescent microscope at 450–490 nm wavelength and 40x magnification.

Evaluation of antibody responses.
Rabbits were either unvaccinated (n=3) or vaccinated with LSDV-wt intramuscularly (Neethling strain, n=3), rLSDV-RG intramuscularly (n=2, 1x105 f.f.u. ml-1), rLSDV-RG intradermally (n=4, 1x105 f.f.u. ml-1) or Verorab intramuscularly (n=3; 1 equivalent human/animal dose; Pasteur Merieux, France). All groups were vaccinated on day 0 and boosted on days 28 and 70. Verorab-vaccinated rabbits were vaccinated on day 0 and boosted on day 28 only. Blood was taken every 2 weeks from all rabbits and stored at -70 °C until tested. ELISAs (Trousse Platelia Rage ELISA kit; Diagnostics Pasteur) to measure RG-specific antibodies were performed according to the manufacturer's specifications on serum derived from each of the rabbits. RV neutralization assays were performed at Onderstepoort Veterinary Institute (Onderstepoort, South Africa) on selected rabbit serum samples (according to the methods of Cliquet et al., 1998). BALB/c mice were either unvaccinated or vaccinated with rLSDV-RG intramuscularly, intradermally and orally (107 f.f.u. ml-1) on days 0, 14 and 28 (n=3 per group). Blood was drawn from the tail vein of mice every 14 days and the serum tested by ELISA for the presence of RG-specific antibodies.

Lymphoproliferation assay.
BALB/c mice (8–12 weeks, n=3 per group; Animal Unit, University of Cape Town) were inoculated intradermally on days 0 and 28 with PBS (50 µl), rLSDV-RG (5x104 f.f.u. ml-1 in 50 µl), LSDV-wt (5x104 f.f.u. ml-1 in 50 µl) or Verorab (50 µl; 1/10 human equivalent dose). On day 38, the mice were sacrificed and splenocytes were isolated by passage through a steel mesh (Sigma) to obtain a single cell suspension. Contaminating red blood cells were removed by centrifugation over Ficoll–Hypaque density gradients (Sigma). Freshly isolated splenocytes were seeded in quadruplicate into round-bottomed 96-well culture plates (2x105 cells per well; Nunc) and incubated in the presence of inactivated rabies virus (two virus particles per cell; State Vaccine Institute, Cape Town, South Africa) or unstimulated for 6 days at 37 °C, 5 % CO2. [3H]Thymidine (1 µCi per well; Sigma) was added to each well for the last 18 h of the assay. Cells were harvested using an automated cell harvester (PHD, Cambridge Technology) and the radioactivity was measured by using a liquid scintillation counter (Tricard-4640). Statistical analysis was carried out using Student's t-test for paired samples.

Rabies virus potency test.
NMRI mice (3–4 weeks of age, n=4 per group; South African Vaccine Producers) were kept in an isolated animal unit at the State Vaccine Institute (Pinelands, Cape Town). All personnel handling the mice were rabies immune and prior approval from the ethics committee was granted (UCT Research Ethics Committee approval reference: 01/16). Mice were inoculated intracranially with various dilutions (8x10-3, 8x10-2, 8x10-1, 1·8x101 or 2·8x101 LD50) of CVS-11 (Rabies Unit, Onderstepoort Veterinary Institute, South Africa, supplied at 4·8x101 LD50). The mice were monitored for 16 days post-challenge to ascertain which dose of the intracranial challenge would give the most reproducible results. From this experiment, it was ascertained that CVS-11 doses of >=8x10-1 resulted in 100 % fatality following intracranial challenge, while CVS-11 doses of <8x10-1 resulted in 25–50 % fatality (data not shown). CVS-11 doses of 8x10-1 LD50 (100 % fatality in unvaccinated animals), 8x10-2 LD50 (25–50 % fatality in unvaccinated animals) and 8x10-3 LD50 (25 % fatality in unvaccinated animals) were used in the subsequent challenge experiments. The mice were monitored for 16 days post-challenge. Any survivors were euthanized and brains were stained with FITC-labelled RV nucleoprotein antibodies to ascertain whether or not mice were infected with RV. All mice that had died by day 16 had positive brain impression stains, while those that were euthanized on day 17 were negative. It was thus concluded that death up to day 16 was an accurate end-point for rabies infections in mice.

Rabies virus challenge experiments.
NMRI mice (3–4 weeks, n=30 per group; South African Vaccine Producers) were inoculated intramuscularly on days 0 and 14 with either PBS (50 µl), rLSDV-RG (5x104 f.f.u. ml-1 in 50 µl), LSDV-wt (5x104 f.f.u. ml-1 in 50 µl) or Verorab (1/10 human equivalent dose in 50 µl). On day 21, ten mice from each group were challenged intracranially with 8x10-1 LD50, 8x10-2 LD50 or 8x10-3 LD50 dilution of live rabies virus (CVS-11; supplied by Onderstepoort Veterinary Institute's Rabies Unit, South Africa). All mice that died within the first 5 days post-challenge were considered non-rabies-related deaths (Seligmann, 1973). Rabies symptomatic and non-symptomatic deaths were screened using a fluorescent antibody test (FAT) of a brain impression (Dean et al., 1996). Survival was monitored between days 5 and 14. During the monitoring period, mice that displayed advanced rabies symptoms were euthanized and considered to be rabies positive after confirmation using the FAT (Dean et al., 1996). Statistical analysis was carried out using the Mann–Whitney U-test for paired samples.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Transient gene expression
A transient expression system was used to determine whether LSDV transcriptases are capable of recognizing both early/late and late poxvirus promoter sequences in cells of ruminant and non-ruminant origin (MacGregor & Caskey, 1989). LSDV-infected LT (ovine, permissive), MDBK (bovine, permissive) and CV-1 (monkey kidney, non-permissive) cells were transfected with plasmids carrying the lacZ reporter gene under the control of either the late VV promoter P11 or the early/late VV promoter P7.5. No endogenous {beta}-galactosidase activity was evident in uninfected MDBK, LT or CV-1 cells. Both P7.5 (early/late) and P11 (late) promoters in permissive (MDBK and LT) and non-permissive (CV-1) cells stained positive for {beta}-galactosidase activity (data not shown), indicating that both early/late and late poxvirus promoters are recognized in rLSDV-RG-infected cells and that these promoters can direct the expression of foreign genes.

Electron microscopy of LSDV-infected permissive and non-permissive cells
We next examined the extent of rLSDV-RG maturation within permissive bovine MDBK and ovine LT and non-permissive primate CV-1 cell types by thin-section electron microscopy (Fig. 1). Fig. 1(A) and (B) are representative of rLSDV-RG infections in permissive MDBK cells. These displayed evidence of advanced infection as demonstrated by intra- and extracellular virions. In permissive cells (MDBK, Fig. 1A, B) but not in CV-1 cells (Fig. 1C), LSDV virions condensed and became oblong in shape, typical of poxvirus morphology. rLSDV-RG virus morphology and maturation in infected ovine LT cells was comparable with that observed for MDBK cells (data not shown).




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Fig. 1. Morphogenesis of recombinant LSDV-RG within permissive and non-permissive host cells. (A, B) Permissive bovine MDBK cells were infected with rLSDV-RG (1 f.f.u. per cell, 48 h, 6600x magnification). The electron micrograph demonstrates mature virions inside (A) and outside (B) the cell. The inserts (26 000x magnification) show high-power virion structure. (C) Non-permissive primate CV-1 cells were infected with rLSDV-RG (1 f.f.u. per cell, 48 h, 9000x magnification). The insert is a 40 000x magnification of the ‘viral factory’.

 
rLSDV-RG replication in non-permissive CV-1 (Fig. 1C) cells proved to be incomplete as virions appeared circular, less condensed and were not observed budding or as extracellular virions.

Expression of foreign genes by rLSDV-RG in permissive and non-permissive cells
Having established that LSDV transcriptases were capable of transcribing foreign genes in non-permissive cells but that rLSDV-RG undergoes only incomplete replication in these cells, we wanted to investigate the expression of foreign gene products. Using the expression of the reporter gene {beta}-galactosidase in rLSDV-RG as a marker, the number of permissive (LT and MDBK) versus non-permissive (primate CV-1 and murine 3T3) cells infected with rLSDV-RG over a 72 h period was evaluated (Fig. 2A). Both non-permissive cell lines were poorly infected with rLSDV-RG with only 3·8±1·2 (mean±SD) % of primate CV-1 and 4·7±1·0 % murine 3T3 cells staining positive for {beta}-galactosidase at 72 h. In comparison, rLSDV-RG infected 53·0±7·4 % of LT and 28·5±6·2 % MDBK cells. Whereas both LT and MDBK cells displayed evidence of foci formation and productive infection (Fig. 2B; as evidenced by staining of >2 adjacent cells for {beta}-galactosidase activity) by 48 and 72 h of infection, no foci formation was observed for CV-1 and 3T3 cells. When the level of {beta}-galactosidase activity was investigated, however, both permissive MDBK and non-permissive CV-1 cells showed similar levels of {beta}-galactosidase activity (Fig. 2C). In comparison, the level of {beta}-galactosidase activity in permissive LT and non-permissive 3T3 cells, although similar to each other, showed levels of activity that were threefold lower than MDBK and CV-1. Since it is clear from Fig. 2(A) and (B) that rLSDV-RG replicates faster in LT than in MDBK cells and that it replicates comparably in CV-1 and 3T3 cells, it is unlikely that efficiency of replication directly impacts on the level of recombinant antigen expression. It may be that the different cellular and/or species origin of the cell lines used in this study play a role in the efficiency of recombinant antigen expression within individual cells. Although non-permissive cell lines are infected with rLSDV-RG with six- to 13-fold lower efficiency than permissive cell lines, this data indicates that the level of foreign gene expression in each individual cell is generally comparable.



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Fig. 2. Recombinant gene ({beta}-galactosidase) expression by rLSDV-RG in permissive (lamb LT and bovine MDBK) and non-permissive (murine 3T3 and monkey CV-1) cell lines. Permissive and non-permissive cells lines were infected with rLSDV-RG containing the {beta}-galactosidase reporter gene (2·5x103 f.f.u. ml-1) for 72 h. (A) The number of cells expressing the {beta}-galactosidase gene product was determined using X-Gal staining and counting at least 200 cells per well in three independent experiments. Results were expressed as a percentage (mean±SD). (B) Formation of foci (indicative of productive spread of virus) was evaluated by determining the number of infected ({beta}-galactosidase-expressing) cells that were bordered by similarly infected cells (>2 adjacent cells infected) or present as a single infected cell (1 cell infected). For this, 100 infected cells from three independent experiments were assessed. (C) The level of {beta}-galactosidase activity in permissive and non-permissive cells was determined. {beta}-Galactosidase activity was normalized by expression as a function of the number of cells infected and expressed as µunits per % infected cell. Each bar represents the mean level of activity (±SD) of three independent experiments.

 
The expression of RG in permissive and non-permissive cells was evaluated using immunofluorescence (Fig. 3). A high proportion of permissive MDBK cells stained positive for RG and clear foci were evident at 72 h p.i. (Fig. 3B). Both non-permissive cell lines (CV-1 and 3T3; Figs 3D and F, respectively) stained positive for RG indicating efficient expression of recombinant antigen by rLSDV-RG. The proportion of cells expressing RG was low and clear foci formation was absent at 72 h infection.



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Fig. 3. Rabies glycoprotein expression in permissive and non-permissive cell lines. Permissive (MDBK, A and B) and non-permissive (CV-1, C and D; 3T3, E and F) cells lines were infected with LSDV-wt (4x103 f.f.u. ml-1; A, C and E) or rLSDV-RG (4x103 f.f.u. ml-1; B, D and F) for 72 h. Expression of rabies glycoprotein was determined by immunofluorescence using purified anti-rabies glycoprotein monoclonal antibody (Biodesign, ME) and rabbit anti-mouse FITC-conjugated secondary antibody (Dako). Both antibodies were used at concentrations suggested by the manufacturers. Fluorescence was viewed using a Zeiss Axioscop 2 microscope at 40x magnification.

 
Evaluation of RG-specific antibody responses in non-ruminant hosts
To assess the immunogenicity of rLSDV-RG in non-ruminant hosts, rabbits and mice were immunized with rLSDV-RG and RG-specific humoral immune responses were measured. Antibodies to RG were detected in all of the rabbits immunized with rLSDV-RG after the first booster inoculation, but not in LSDV-wt- or non-immunized rabbits (Table 1). The World Health Organization (WHO) has defined the critical antibody level necessary for protection against rabies to be >=0·5 IU ml-1 (in humans and other species except cattle and horses; Dutta et al., 1992; Cliquet et al., 1998). This level of RG-specific antibodies was attained in all rLSDV-RG-immunized rabbits tested after the first booster inoculation and increased substantially after the second booster inoculation. Similar levels of RV-specific antibodies were elicited in rabbits by rLSDV-RG irrespective of the route of immunization (intramuscular versus intradermal). None of the PBS-, LSDV-wt- or DMEM-inoculated control animals produced detectable antibodies to rabies antigen.


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Table 1. Rabies virus glycoprotein-specific antibodies (IU ml-1) in rabbits immunized with rLSDV-RG or Verorab

All rabbits were immunized intramuscularly (IM) except for one group that was immunized intradermally (ID). A titre of 0·5 IU ml-1 has been defined by the World Health Organization as the minimum antibody titre required to elicit protection against rabies. Values are expressed as the mean IU ml-1 (±SD). The detection limit of the ELISA was >=0·3 IU ml-1.

 
The antibody responses of two of the rLSDV-RG rabbits were tested for their ability to neutralize live RV (Table 2). By 12 weeks post-immunization, both were found to have levels of neutralizing antibodies that were approximately twice that required to provide protection (as defined by the WHO), whereas the control animals remained negative. Neutralizing antibody titres were not determined in Verorab-immunized rabbits.


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Table 2. Rabies virus neutralizing antibodies (IU ml-1) in rabbits immunized with rLSDV-RG

 
BALB/c mice vaccinated with rLSDV-RG failed to produce measurable antibodies to rabies virus despite a booster inoculation (data not shown). Verorab, by comparison, induced moderate RG-specific humoral immunity in mice with the levels of 0·5 IU ml-1 being attained following a booster inoculation.

rLSDV-RV immunization induces RV-specific T-cell proliferation in mice
To investigate the ability of rLSDV-RG to induce cell-mediated immune responses to rabies antigens, T-cell proliferative activity was investigated following immunization. Splenocytes from mice immunized with rLSDV-RG showed significantly stronger proliferation to inactivated rabies virus than LSDV-wt-inoculated mice (P=0·02) and was comparable with levels induced by Verorab (Fig. 4).



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Fig. 4. rLSDV-RV immunization induces strong T-cell proliferative responses to inactivated rabies virus following in vitro restimulation. BALB/c mice (8–12 weeks) were immunized twice intradermally (days 0 and 28) with PBS, LSDV-wt, rLSDV-RV or Verorab, and in vitro splenocyte proliferation to either no antigen (control) or inactivated rabies virus (two particles per cell) was determined by [3H]thymidine incorporation. Results were expressed as stimulation index and were calculated as: (c.p.m. of test wells)/(c.p.m. of control wells). Each bar represents the mean stimulation index (±SD) of quadruplicate wells. * P<0·05; ** P<0·01 (comparing rLSDV-RG- or Verorab-immunized animals with LSDV-wt-immune controls using Student's t-test for paired samples).

 
Protection against rabies virus challenge
To assess the efficacy of rLSDV-RG against virus challenge, mice immunized with rLSDV-RG, LSDV-wt, Verorab or PBS were infected intracranially with live rabies virus and their survival was monitored for 14 days post-challenge (Fig. 5). Fig. 5(A) shows the survival curve of the mice challenged with a high dose of rabies virus [8x10-1 LD50 of CVS-11 (challenge virus standard)]. Immunization with rLSDV-RG and Verorab provided strong protection against high-dose rabies virus challenge compared with LSDV-wt and PBS immunization (P=0·0021 comparing rLSDV-RG with LSDV-wt immunization). The onset of rabies disease was later in the rLSDV-RG- and Verorab-vaccinated groups than the LSDV-wt and PBS groups. Fig. 5(B) shows the survival curve of the mice challenged with a lower dose of rabies virus (8x10-2 LD50 of CVS-11). None of the rLSDV-RG- and Verorab-vaccinated mice died, while 50 and 60 % of the inoculated mice in the PBS and LSDV-wt groups, respectively, died (P=0·0002 comparing rLSDV-RG with LSDV-wt immunization). This experiment conclusively shows that protection against a live RV infection can be brought about by inoculation with rLSDV-RG and that the protective ability of this candidate vaccine is comparable with that of Verorab.



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Fig. 5. Immunization with rLSDV-RG protects mice from intracranial rabies virus challenge. Survival plot of NMRI mice inoculated on days 0 and 14 with PBS ({blacklozenge}, n=20), rLSDV-RG ({blacksquare}, 5x104 f.f.u. ml-1, n=20), LSDV-wt ({bullet}, 5x104 f.f.u. ml-1, n=20), or Verorab ({Delta}, 10 % human dose, n=20) and challenged with 8x10-1 LD50 RV (CVS-11) (A) or 8x10-2 LD50 RV (CVS-11) (B).

 

   DISCUSSION
Top
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Many studies are currently focusing on the use of attenuated MVA as a recombinant vaccine vector because of its proven safety record in immunocompromised primates (Barouche et al., 2001; Stittelaar et al., 2001). The efficacy of this vector, however, may be limited by pre-existing immunity to other VV strains (Baxby & Paoletti, 1992). Although all countries have discontinued their VV vaccination programmes because of the eradication of smallpox, the renewed threat of bioterrorism has led some countries such as the USA to prepare for the reintroduction of VV vaccination. Studies on fowlpox virus (Somogyi et al., 1993) and canarypox virus (Taylor et al., 1991; Moss, 1996) have shown that such poxviruses undergo incomplete or abortive replication cycles in mammalian cells. This feature has become known as ‘host restriction’ and has subsequently been applied to other attenuated viral vectors such as MVA. Host-restricted vaccine vectors are assumed to be a lower biological containment risk than vaccine vectors with broad host ranges because such replication deficiency limits the threat of environmental spread.

We have investigated a host-restricted capripoxvirus, LSDV, as a novel vector for effective but safe replication-deficient recombinant vaccines. LSDV has been described as highly species-specific and its associated disease is limited to ruminants (Weiss, 1968; Young et al., 1970). In order to explore the stringency of this host restriction, we compared the replication competence of recombinant LSDV in cells of bovine (MDBK), ovine (LT) and primate (CV-1) origin. Although our study was limited to only a few cell types, this represents the first ultrastructural comparison of the morphogenesis of LSDV in cultured cells of ruminant and non-ruminant origin. The ultrastructure of LSDV in bovine MDBK cells observed in this study is consistent with the results of Prozesky & Barnard (1982) who investigated the pathogenesis of wild-type LSDV in naturally infected cattle using electron microscopy.

LSDV viral factories – electron dense areas consisting of nucleic acid material and proteinaceous membrane structures and viral particles – were seen in non-permissive primate CV-1 cells, providing evidence of incomplete replication. No mature particles were observed.

To confirm that both early/late and late expression of the foreign gene occurs in LSDV-infected cells, we used a transient expression assay to demonstrate that both late (p11) and early/late (p7.5) poxvirus promoters were able to drive the expression of foreign genes in both LSDV-infected permissive (ovine LT and bovine MDBK) and non-permissive (primate CV-1) cells. We further demonstrated that both the recombinant antigen (RG) and {beta}-galactosidase (the reporter gene product) were detectable in rLSDV-RG-infected cells of both ruminant and non-ruminant origin. Consistent with the morphogenesis results, we found that rLSDV-RG did not productively infect non-permissive cells as evidenced by the lack of clear foci formation but that expression levels of recombinant antigen were similar in permissive and non-permissive cells.

This finding, together with the result from the ultrastructural study, was extremely encouraging as it provides preliminary evidence that non-permissive cells can support the early stages of rLSDV-RG replication. While complete maturation of LSDV particles is restricted to cells of ruminant origin, expression of rLSDV-encoded foreign genes can occur in both ruminant and non-ruminant cells.

We have previously presented evidence that rLSDV-RG expresses RG adequately in ruminant hosts so that immunity to that protein is elicited (Aspden et al., 2002). In cattle, rLSDV-RG induces cell-mediated immunity to RV and neutralizing antibody titres of up to 3000 times the WHO prescribed threshold level of protection (0·5 IU ml-1 in humans and other species except cattle and horses; Cliquet et al., 1998). These findings confirm that RG is expressed by the bovine cells infected with rLSDV-RG in such a manner that RV-specific cell-mediated and humoral immunity is elicited in cattle. Furthermore, Romero et al. (1993, 1994a, b) have demonstrated immunity against rinderpest in cattle that had been vaccinated with a recombinant capripoxvirus expressing an antigenic protein of rinderpest virus.

To evaluate further the efficacy of LSDV as a recombinant vaccine vector in non-ruminant hosts, we investigated the immunogenicity of rLSDV-RG in rabbits and mice. The rLSDV-RG vaccine induced RV neutralizing antibody titres in rabbits following a boost that exceeded those that have been found to provide protection from rabies infection (Dutta et al., 1992; Cliquet et al., 1998). In mice, immunization with rLSDV-RG generated significant T-cell proliferative activity towards inactivated RV indicating that a cell-mediated immune response was induced. No RV-specific antibody responses were detectable in mice.

Most importantly, we have clearly shown that vaccination with a relatively low dose of rLSDV-RG protects mice against an aggressive intracranial challenge with live RV. As no antibodies to RG were detected in mice, a direct interpretation would be that this protection was cell mediated. This is contentious, however, since previous studies have demonstrated that neutralizing antibodies are central to protection against rabies challenge but that other effector mechanisms may suffice for protection in the absence of an antibody response (Mifune et al., 1981; Xiang et al., 1995). It is possible that induction of a cellular response (as demonstrated in this study) allows for an accelerated antibody response following rabies virus challenge that is sufficient to protect the mice. Whatever the mechanism, since intracranial challenge with live RV is the gold standard against which all rabies vaccines are compared, we can conclude that rLSDV-RG is comparable with other commercially used rabies vaccines in its ability to protect against RV infection.

Recombinant LSDV-RG virus titres used in this study were 500–50 000-fold lower (104–105 f.f.u. per dose) than titres reported in studies using non-replicating poxvirus vectors (ranging from 5x106 p.f.u. per dose for canarypox to 5x108 p.f.u. per dose for MVA; Allen et al., 2000; Horig et al., 2000; Men et al., 2000; Amara et al., 2002). While we have clearly shown that these low rLSDV-RG titres provide protection from RV challenge following only two immunizations thereby strongly supporting the efficacy of our vaccine, this result is tempered by the difficulty of raising high titres of LSDV in tissue culture. This is an important consideration that could affect future developments of the capripoxvirus vectors in non-ruminant hosts.

In conclusion, this study demonstrates that rLSDV-RG shows potential as a novel vaccine against rabies in non-ruminant hosts. Furthermore, the advantages and efficacy we report here of LSDV as a host-restricted vaccine vector strongly suggest that this virus may prove useful as a potentially safe, replication-restricted vaccine vector.


   ACKNOWLEDGEMENTS
 
Research funding from the NRF and bursaries from the PRF and DR McIntosh Trust Fund is gratefully acknowledged. We acknowledge Keith Dumbell for helpful discussions and initiating LSDV research at UCT. We thank Maureen Dennehy and Darren Martin for critically reading the manuscript, Vincent Sharp and Rodney Lucas for their expert technical assistance with the rabies virus challenge experiments, the State Vaccine Institute for making their facilities available, John Bingham from the Rabies Unit at Onderstepoort Veterinary Institute for providing the rabies virus and doing the neutralization assays, and Nicky Johnston for providing useful advice about immunofluorescent staining. Students and staff in the Division of Medical Virology (UCT) are thanked for advice and support.


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Received 22 January 2003; accepted 19 March 2003.



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