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In order to assess the intracellular localisation of VP2 in virus infection, cells were first infected with BTV-10 and then VP2 detected by immunofluorescence microscopy. At 4 hours post infection VP2 was found to be restricted to punctuate areas within the cytoplasm (Fig. 2A). A similar distribution of VP2 was observed when cells were transfected with plasmids encoding VP2 or a VP2-GFP fusion protein where the GFP was located at the C-terminus of VP2 (Fig. 2B and 2C, respectively). However, fusion of GFP to the N-terminus of VP2 altered the subcellular localisation of the protein and resulted in a more diffuse pattern of fluorescence (Fig. 2D). Western blot analysis of the VP2 GFP fusion proteins confirmed that they were expressed in cells as full-length proteins (Fig. 2F).
Since the fusion of GFP to the N-terminus but not the C-teriminus of VP2 altered its sub-cellular localisation we hypothesised that the N-terminus of the protein contained a signal necessary for intracellular localisation that was disrupted in the fusion protein. To test this possibility, we expressed an N-terminal fragment of VP2 consisting of the first 118 residues of the protein fused at its C-terminus to GFP (Fig. 3A, VP21–118GFP). The distribution of this deletion mutant was indistinguishable from that of the full-length VP2-GFP protein (Fig. 3A, VP2 GFP), suggesting that it contained the signals necessary for the subcellular localisation of VP2. The punctuate pattern of VP21–118GFP accumulation was distinct from what would be expected if the chimeric protein had been incorporated into aggresomes as there was no evidence of co-localisation with ubiquitin (Fig. 3A, ubiquitin).
Earlier electron microscopic studies of BTV infected cells have suggested that the virus associates with the cell membrane and with vimentin intermediate filaments [22,26]. In order to test where VP21–118GFP was localised within cells, subcellular fractionation studies were carried out in the presence of detergents at both 4°C and 37°C (Fig. 3B). These studies were designed to separate soluble cytoplasmic proteins and lipid raft associated proteins from cytoskeletal and nuclear fractions of the cell. As expected, actin was present as both soluble and cytoskeletal associated insoluble forms under the conditions of the study. Tubulin and untagged GFP were present predominantly in the detergent soluble cytosolic fraction. The nuclear membrane protein lamin was present predominantly in the detergent insoluble fractions at both 4°C and 37°C, although at the higher temperature the proportion of solubulised lamin was increased. Vimentin and VP2 co-segregated in these fractionation studies, and were present only in the detergent insoluble fractions (Fig. 3B). Thus, fractionation studies supported association with VP2 with detergent insoluble cell fractions containing vimentin. Immunofluorescence was performed with each of the proteins in the cell fractionation studies in Vero cells expressing VP21–118GFP. While there was no evidence that either actin, tubulin or lamin co-localised with the VP2 deletion mutant, all of the VP21–118GFP was detected in areas of the cell rich in vimentin (Fig. 3C). Thus, in both cell fractionation and colocalisation studies VP21–118GFP and vimentin were detected together.
In order to confirm the association between vimentin and full-length VP2, the distribution of vimentin and VP2 in virus infected cells was monitored by immunoflorescence. BTV infection of mammalian cells in tissue culture leads to rapid changes in cell morphology with apoptosis, which is triggered during virus entry . Thus the distribution of VP2 and vimentin was assessed at early (8 hours) and late (24 hours) timepoints following virus infection (Fig. 3D). At 8 hours post infection vimentin was distributed throughout the cell but was also present in concentrated foci within the cytoplasm. At the same timepoint, VP2 was detected in concentrated foci within the cells. Intriguingly, the largest foci of VP2 co-localised with the largest foci of vimentin. All the VP2 in detected in virus infected cells was found associated with vimentin, although not all the vimentin at this timepoint was associated with VP2 (Fig. 3D). In contrast, at 24 hours post infection, by which time there are substantial virus-induced changes in cell morphology almost all the vimentin and VP2 present in the cell colocalised (Fig. 3D). Thus, data from virus infected cells confirm the VP2-vimentin association detected with the VP21–118GFP deletion mutant.
Having established that VP21–118GFP and full-length VP2 both colocalised with vimentin we went on to perform deletion mutagenesis of the VP21–118GFP mutant in order to determine the regions of the protein critical to this interaction. Five deletions were introduced to the 1–118 region of VP2, to produce in-frame truncations of this region of the protein (Fig. 4A). Of these, deletion of the 49 amino acids between position 65 and 114 resulted in a diffuse distribution of the truncated protein (Fig. 4B). Further truncation of the protein, by introducing deletions from amino acids 65–92 or 93–114 gave no further resolution to the analysis, as both of these deletions abolished punctuate localisation of the VP2 protein (Fig. 4B). Deletions between amino acids 13 and 64 had no effect on the localisation of the VP21–118GFP protein, suggesting that this region does not contribute to the subcellular localisation of the protein.
The most likely reasons that deletions from 65–92 and 93–114 both resulted in diffuse localisation of the VP2 truncation mutant were that there were either multiple localisation signals, or the signal extended across the amino acid 92–93 junction, or these deletions had long-distance effects on folding of the remaining VP2 sequences. In order to resolve these possibilities we carried out site directed mutagenesis of conserved amino acids in this region of the protein. Sequence alignment of the sequence of the VP2 protein from 13 different BTV serotypes revealed that there were four amino acid regions within this sequence which were either completely conserved or very similar. These were the regions G70 to L81, D86 to K88, F98 to T100 and W104 to I109 (Fig. 5A). Five sets of mutations were generated; the first four were designed to mutate charged or conserved amino acids in each of the four conserved regions to alanine. The rationale for selecting the charged amino acids for mutation was that it was expected that these proteins would be more likely to be solvent exposed and thus available to interact with vimentin. The fifth mutation was to target the conserved GXV motif present at position 70–72. Since mutation of these amino acids to alanine would have been a very conservative change, and these residues are part of the only strongly hydrophobic region of the truncated protein, GVV was mutated to DVD. This mutation has the effect of altering the mean hydrophobicity of this region of the truncated VP2, such that the hydrophobic patch normally present in this region of protein has mean hydrophilic properties (Fig. 5B).
On expression, all of the four alanine mutations had similar punctate accumulation patterns to the unmodified VP2 sequence (Fig. 5C). In contrast, the mutation of the GVV to DVD at positions 70–72 resulted in a diffuse pattern of expression similar to that seen with the ΔVP265–92GFP and ΔVP293–114GFP deletion mutants. Vimentin targeted immunofluorescence with cells expressing the DVD (M5) mutant revealed that the association between vimentin and VP2 had been broken (Fig. 5D). Because the M5 mutation was designed to alter the hydrophobicity profile of the vimentin binding region of VP2 it was possible that the changes in accumulation seen were a result of global changes in the folding of the VP2 protein. In order to address this possibility, the M5 mutation was introduced into the full-length VP2 protein, and this protein expressed in insect cells using the baculovirus expression system. Unmodified VP2 multimerises to form a trimeric triskelion structure present on the outer surface of the mature virus particle . By separating purified, unmodified VP2 under native conditions it is possible to detect monomeric, dimeric and trimeric forms of the protein (Fig. 5E). Separation of the full-length M5 VP2 mutant under the same conditions resulted in the formation of a similar proportion of each of the multimeric states of the VP2 protein as the unmodified protein. Thus the mutation that prevented the association of VP2 with vimentin did not prevent the formation of higher order multimers of the VP2 protein. This suggests that while it is possible that the local protein architecture was affected by the M5 mutation the overall folding of the protein was similar enough to the unmodified protein that multimerisation was unaffected.
The vimentin intermediate filament network has a radial organisation, extending outwards from the cell centre. This localization partially overlaps with that of microtubules there is evidence that the two filament systems interact [29,30]. Indeed, vimentin structures move along microtubules [29,31-33]. Therefore, if the vimentin-VP2 interaction was important for BTV assembly and/or egress we predicted that pharmacological disruption of either microtubules or vimentin would result in changes of the amount of virus produced or the amount of virus released from infected cells. To test this possibility we carried out pharmacological experiments with colchicine and acrylamide which specifically disrupt the microtubule and vimentin intermediate filament networks respectively. Consistent with our previous experiments, in untreated cells vimentin was detected throughout the cytoplasm and this distribution was unaffected by the expression of full-length, untagged VP2. However, in the presence of colchicine, as expected, there was a redistribution of vimentin and VP2 towards the nucleus (Fig. 6A). This observation supports other observations that vimentin intermediate filaments are organised by the microtubule network [29,30]. The redistribution of vimentin was even more dramatic on the addition of acrylamide which resulted in a collapse of the intermediate filament network and a change in cell morphology to a more rounded shape (Fig. 6A). To test the effect of these changes in vimentin distribution on virus replication and release, virus was adsorbed to cells and then cells were treated with either colchicine (Fig. 6B) or acrylamide (Fig. 6B). Indirect disruption of vimentin with colchicine resulted in 2.5 times more cell associated virus and a five-fold reduction in the amount of the released virus detected compared to untreated control cells. Addition of acrylamide resulted in a 50 fold reduction in the amount of released virus and a 250 fold increase in the amount of cell associated virus at 24 hours post infection. These data suggest that disruption of vimentin either directly or indirectly inhibits the trafficking of mature virus particles out of virus infected cells.
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