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INSERM E 0345, EFS-Alsace, 10 rue Spielmann, BP 36, 67065 Strasbourg Cédex, France
Correspondence
Robert Drillien
robert.drillien{at}efs-alsace.fr
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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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; Meyer et al., 1991; Carroll & Moss, 1997; Drexler et al., 1998). Primary or established human cell lines are non-permissive for MVA, although some tumour cell lines enable very low levels of virus multiplication when high multiplicities of infection are employed (Carroll & Moss, 1997; Blanchard et al., 1998). MVA infection in HeLa cells is arrested at a late stage of the virus cycle, after the accumulation of immature virus particles (Sutter & Moss, 1992) but before the association of these particles with nascent DNA (Sancho et al., 2002), a step required for the formation of intracellular mature virus. The defectiveness of MVA makes it an attractive candidate for use as a viral vector, as well as a smallpox vaccine if such a vaccine is again needed. Studies in animal models have demonstrated that MVA is indeed an effective vaccine vector (Moss et al., 1996; Sutter & Staib, 2003) and that it is safe even in immunocompromised animals (Stittelaar et al., 2001). Although higher MVA titres are often used to induce an immune response similar to that in a standard VACV infection (Belyakov et al., 2003), the titres employed may be lower than the final body titres of replicating VACV strains. This suggests that MVA displays some unique features that enable it to induce an immune response without propagating. One explanation may lie in the fact that MVA lacks many of the genes found in standard VACV strains that counteract the immune response (Antoine et al., 1998; Blanchard et al., 1998). Another key factor may be the ability of MVA to induce human fibroblasts and human leukocytes to secrete type I interferons known to boost T cell responses (Buttner et al., 1995; Blanchard et al., 1998). In fact, MVA has been shown to efficiently re-stimulate human cytotoxic T lymphocytes in vitro (Dorrell et al., 2000). In this study we focus on the interactions of MVA with human dendritic cells (DCs), as a considerable amount of work has demonstrated the importance of these antigen-presenting cells in the induction of the immune response (Banchereau & Steinman, 1998). Prior investigations into the interaction of VACV with DCs have mainly been conducted with replicating VACV strains. Such studies have shown that DCs allow the expression of early virus genes, but are non-permissive for viral DNA replication or late protein synthesis (Bronte et al., 1997; Engelmayer et al., 1999; Drillien et al., 2000; Jenne et al., 2000). Furthermore, VACV-infected DCs fail to respond to agents that induce their activation (Engelmayer et al., 1999; Jenne et al., 2000), a step also termed maturation, which is required for de novo stimulation of T lymphocytes but not necessarily for memory responses. In contrast, it has been reported that MVA-infected DCs can be activated by tumour necrosis factor alpha (TNF-
) (Trevor et al., 2001), but no study has examined whether MVA on its own may activate DCs.
Thus we investigated the level of cell surface co-stimulatory molecules and pro-inflammatory cytokine secretion in MVA-infected DCs. We also studied the influence of MVA-infected HeLa cells on DCs by following the level of co-stimulatory molecules and the ability of such mixed populations to induce gamma interferon (IFN-
) secretion from T lymphocytes. These studies highlight the unique interactions of MVA with human dendritic cells.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Fractions containing either lymphocytes or monocytes were frozen separately in liquid nitrogen. When required, the monocytes were thawed and differentiated into DCs for 5–6 days at a density of 106 cells ml–1 in RPMI 1640 medium supplemented with 40 ng interleukin 4 (IL4) ml–1, 50 ng granulocyte macrophage–colony stimulating factor (GM–CSF) ml–1, 0·11 mg sodium pyruvate ml–1, non-essential amino acids for MEM, and 10 % FBS (Sallusto & Lanzavecchia, 1994). After the first 3 days' culture, half of the initial volume of fresh medium was added. When DCs were used for T cell stimulation, monocytes were differentiated in the presence of 2 % pooled human serum (PHS) and heparin (50 IU ml–1) as described previously (Xia & Kao, 2002). All media contained 50 µg gentamicin ml–1. When required, bulk lymphocyte preparations maintained in liquid nitrogen were thawed and cultured in RPMI 1640 supplemented with 0·11 mg sodium pyruvate ml–1, non-essential amino acids for MEM and 5 % PHS. Hamster BHK21 cells were cultured in BHK21 medium supplemented with tryptose phosphate broth and 10 % FBS. Human HeLa cells were grown in MEM with 10 % FBS.
Virus.
VACV strains Copenhagen (VACV-Cop), Lister (VACV-Lis) and MVA (a clonal isolate designated MVA-N33 and kindly provided by Jean-Marc Balloul, Transgène, Strasbourg, France) were grown in BHK21 cells and purified by the standard sucrose gradient procedure (Joklik, 1962). The ratio of virus particles to p.f.u. was very similar for different virus stocks (30 % variation). To construct VACV-Cop and MVA recombinant viruses encoding the enhanced green fluorescent protein (GFP), the corresponding gene was obtained from BD Biosciences Clontech. The GFP gene was first positioned downstream of a synthetic early promoter, which we used previously to drive foreign gene expression from the virus genome (Howley et al., 1996), and this expression cassette was ligated as a 5' NaeI–3' XbaI fragment into the pG08 vector (Sutter et al., 1994) cut with SmaI and NaeI so as to remove the resident promoters. This procedure generated a plasmid designated pETS 32 that also harboured the xanthine–guanine phosphoribosyl transferase gene behind a 7·5 kDa early/late promoter. BHK21 cells were then infected with VACV-Cop or MVA and transfected with pETS 32 using lipofectamine (Invitrogen). Recombinant viruses designated MVA 32 and VACV-Cop 32 were selected on BHK21 cells in the presence of mycophenolic acid, xanthine and hypoxanthine as described previously (Falkner & Moss, 1988). Virus plaques were picked, amplified and cloned several times, and after each series of plaque isolations the clones were tested for expression of GFP by examination under UV light. Single plaques were selected for further study after several rounds of plaque isolation during which all plaques were positive for GFP expression. The MVA 32 recombinant isolated in this manner was identical to the MVA parental strain except for the inserted genes whereas, due to the strategy employed, the VACV-Cop 32 recombinant was mutated and deleted in ORFs A52R to A55R corresponding to deletion III in MVA. This deletion was not associated with any difference in plaque formation or host range, as reported previously (Howley et al., 1999). In some experiments we used a VACV-Cop recombinant (VACV-Cop 21) encoding GFP inserted in the I4L locus (large subunit of the viral ribonucleotide reductase). In this case the GFP gene was positioned behind the same synthetic early promoter as in VACV-Cop 32, and both viruses induced the same level of GFP expression. When required, virus was subjected to UV inactivation for 20 min in the presence of 25 µg ml–1 psoralen using a 365 nm UV bulb positioned 2·8 cm over the sample, according to a procedure described previously (Tsung et al., 1996).
Flow cytometry.
Five to 6 days after differentiation of monocytes into DCs, the cells were pelleted, counted and resuspended at 5x105–106 cells ml–1 in fresh culture medium containing 50 ng GM–CSF ml–1 and 40 ng IL4 ml–1. The DCs were then infected with purified virus that had been subjected to 5 s ultrasonic treatment to disaggregate clumps. Lipopolysaccharide (LPS) endotoxin from Escherichia coli (serotype O128 : B12) was routinely employed to check for the ability of DCs to undergo maturation. Eighteen to 24 h after infection the cells were pelleted, washed once in PBS and resuspended for 30 min in PBS containing 5 % normal goat serum (NGS) and a 20-fold dilution of a specific primary mouse monoclonal antibody or a control antibody of the same isotype. The cells were then pelleted, washed once in cold PBS and incubated for 30 min in PBS containing 5 % NGS and F(ab')2 sheep antibodies directed against mouse IgG that had previously been coupled with FITC. The cells were pelleted again, washed once in PBS, resuspended in PBS and fixed with 1 % paraformalehyde. Cells were then examined by flow cytometry using a Becton Dickinson Facscan and CellQuest software. In some experiments, direct immunolabelling was performed with FITC- or phycoerythrin (PE)-coupled mouse antibodies. Alternatively, HeLa cells in monolayers were infected with MVA or VACV-Cop for 15 h, then detached from the culture flasks with 0·02 % EDTA in PBS, pelleted, washed twice with PBS to remove any input virus, and added to fresh DCs for an additional 24 h (one HeLa cell per four DCs). The mixed population was then double-labelled with a mouse anti-CD1a antibody (HI149) coupled to FITC, which binds only to DCs, and anti-CD86 antibodies coupled with PE. The cells were then examined by flow cytometry in the FL1 and FL2 channels, and only CD1a+ cells in the FL1 channel (FITC) were gated to plot histograms of fluorescence in the FL2 channel (PE).
Cytokine assays.
Dendritic cells cultured at 106 cells ml–1 as described above were infected at different multiplicities for the chosen time interval, after which they were pelleted and the culture medium was recovered and stored in aliquots at –80 °C for cytokine assays. TNF- and IL6 were determined by standard sandwich ELISA tests using reagents from Beckman Coulter. For IFN- assays, 105 HeLa cells in 2 cm2 tissue culture plates were infected with MVA or VACV-Cop for 1 h in PBS, then fresh RPMI containing 5 % PHS was added. After 4 h infection, the culture medium was removed and the cells were washed twice with PBS. Human DCs (105) in RPMI containing 5 % PHS were then added to each culture plate; alternatively, DCs were directly infected with MVA or VACV-Cop. Autologous T lymphocytes were then prepared by depletion of B cells, NK cells, monocytes, activated T cells and granulocytes from total lymphocytes using a combination of antibodies coupled to magnetic beads (anti-CD14, anti-CD16, anti-CD56, anti-HLA class II DR/DP from Dynal) and added 2 h later to the mixture of infected HeLa cells and DCs (106 T lymphocytes per 105 DCs and 105 HeLa cells) or to 105 infected DCs. After overnight incubation, the cells in suspension were removed from the culture dishes, pelleted once and resuspended in fresh RPMI and 5 % PSH. The cells were assayed for IFN- production using an enzyme-linked immunospot (ELISPOT) method and reagents obtained from Diaclone.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). When analysed by flow cytometry, uninfected DCs displayed almost no CD83 on their surface and only moderate levels of CD80 and CD86, indicating that they were in an immature state (Fig. 1a). A striking increase in all three cell surface antigens was observed after 1 day's culture in the presence of 10 ng ml–1 LPS, which is a potent maturation factor for DCs (Fig. 1b). Infection of the DCs for 1 day with MVA (5 p.f.u. per cell) led to a moderate increase in cell surface CD80 and CD83, and a more marked increase in CD86 (Fig. 1c). The extent of increase in these surface molecules was proportional to the m.o.i. employed, being barely detectable at 0·1 p.f.u. per cell and reaching a maximum at 20 p.f.u. per cell (not shown). Infection of the DCs with VACV-Cop resulted in no change in the level of CD83 and CD86, and a decrease in the baseline level of CD80 (Fig. 1d), consistent with our previous studies (Drillien et al., 2000). Thus MVA, in contrast to VACV-Cop, is able to induce a moderate level of DC activation as judged by an increase in cell surface maturation markers. However, in no instance did MVA infection, whatever the multiplicity (0·1–20 p.f.u. per cell), lead to the striking increase in all three maturation markers that was observed in the presence of LPS.
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). Infection by MVA led to a significant, albeit low, level of TNF- and IL6 secretion that increased with m.o.i. The level of cytokine secretion after MVA infection was much lower than that induced by LPS. IL1
secretion was also detected in some experiments, but not consistently (results not shown). VACV-Cop failed to induce any TNF- and induced IL6 secretion barely above the background level of uninfected DCs.
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). Whereas polymyxin B significantly reduced the high level of DC activation observed in the presence of as much as 1 mg ml–1 LPS, it was unable to alter the moderate activation of DCs induced by MVA (Fig. 2).
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, compare c and a). Interestingly, both GFP– and GFP+ cells showed an increase in the level of CD86, indicating that virus gene expression may not be an absolute requirement for DC activation. However, the absence of GFP expression did not necessarily suggest that the DCs had not been infected, as some DCs may undergo abortive infection characterized by virus binding/uptake but no gene expression. To reduce virus gene expression, DCs were infected with MVA 32 that had undergone prior inactivation by UV light in the presence of psoralen. Considering both CD86– and CD86+ DCs, only 8·8 % (7·2+1·6) of cells infected with the UV-inactivated MVA 32 expressed GFP (Fig. 3d), whereas 21·3 % (10·9+10·4) of the DCs infected with the control virus expressed GFP (Fig. 3c), indicating that UV inactivation of the virus led to a significant reduction in its gene expression. However, no decrease but rather an increase in the number of CD86+ cells was observed in the DCs infected with UV-inactivated MVA 32, suggesting that gene expression was not a prerequisite for DC activation. VACV-Cop 21, as reported above for VACV-Cop, failed to increase the level of CD86 on the DC surface (Fig. 3e). However, DCs infected with UV-inactivated VACV-Cop 21 displayed a reduced level of GFP expression and, most surprisingly, an increase in CD86 expression (Fig. 3f) which was nearly comparable to the level achieved after MVA 32 infection.
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). Heating MVA at a higher temperature (1 h at 65 °C) completely destroyed its ability to activate DCs (Fig. 4c). As endotoxins are notoriously resistant to mild heat treatments (such as 1 h at 65 °C), these results also indicate that they are not involved in DC activation by MVA. We also examined the ability of heat-treated VACV to activate DCs and, in this case, used either VACV-Cop or VACV-Lis (results shown for Lister strain). Untreated VACV-Lis failed to activate DCs (Fig. 4a) but prior heat treatment of the virus (55 °C for 1 h) unveiled its ability to activate DCs (Fig. 4b). This feature was further confirmed by the ability of VACV-Cop (treated at 55 °C) to induce TNF- and IL6 secretion (not shown). Heat treatment of VACV-Lis (65 °C for 1 h) destroyed its capacity to activate DCs (Fig. 4c).
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release by T lymphocytes, compare thin lines in a and b). Addition of MVA-infected HeLa cells to DCs induced a large increase in the level of CD86 (Fig. 5b, thick line) demonstrating that MVA-infected cells activated DCs, as reported above for direct virus infection. However, HeLa cells infected with MVA did not induce as high a level of DC activation as was reached after LPS treatment (Fig. 5a). When HeLa cells were infected with VACV-Cop instead of MVA, and then mixed with DCs, there was no increase in the level of CD86 (Fig. 5b, dashed line). To determine whether DC activation required active virus infection, HeLa cells infected with MVA for 12 h were fixed for 30 min with 1 % paraformaldehyde in PBS, then pelleted, washed twice with PBS and added to DCs for another 24 h, and examined for CD86 expression. Under these conditions, HeLa cells infected with MVA, or even with VACV-Cop, activated DCs (Fig. 5c).
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-producing T cells. For these experiments DCs were differentiated from monocytes in the presence of 2 % PHS, instead of FBS, to avoid presentation of bovine antigens to T cells, and 50 units ml–1 heparin to ensure they were bona fide DCs expressing the CD1a antigen (Xia & Kao, 2002). We found that DCs infected directly with MVA or VACV-Cop did not induce a significant number of IFN--producing T cells above the background level (Fig. 6a). Therefore, further experiments were conducted in which HeLa cells were infected, washed to remove any input virus, then mixed with DCs, and finally autologous T cells were added. The combination of MVA-infected HeLa cells and DCs induced a much higher number of IFN--producing lymphocytes than the combination of VACV-Cop-infected cells with DCs (Table 2). No IFN--producing lymphocytes were noticed in the absence of DCs or in the presence of uninfected HeLa cells, and only a small number of spots were visualized with uninfected DCs alone. To extend this analysis we investigated the influence of m.o.i. on the number of IFN--producing lymphocytes (Fig. 6b). The DCs combined with VACV-Cop-infected HeLa cells induced no IFN--producing lymphocytes, or only a small increase above background level in some experiments (not shown). The DCs combined with MVA-infected HeLa cells induced IFN--producing lymphocytes significantly above background level, and the number of spots increased proportionally with the m.o.i.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Jenne et al., 2000). The latter two studies demonstrated that infection with standard VACV strains interferes with DC activation by a number of agents. In contrast, MVA-infected DCs responded to activation by TNF- treatment (Trevor et al., 2001), but no direct activation of DCs by MVA was reported previously, possibly because only a low m.o.i. (0·1 p.f.u. per cell) was employed.
The use of UV- or heat-treated virus (55 °C) enabled us to demonstrate that DC activation did not require virus gene expression, suggesting that virus binding or uptake is sufficient. Similar conclusions have been reached for activation of human DCs by measles virus (Bieback et al., 2002) or activation of murine DCs by mouse mammary tumor virus (Burzyn et al., 2004) or human adenovirus 5 (Hirschowitz et al., 2000). Intriguingly, we found that VACV-Cop or VACV-Lis subjected to UV light or heat treatment at 55 °C activated DCs as efficiently as MVA. Furthermore, VACV-Cop-infected cells, which failed to activate DCs, acquired this capacity if they had previously been fixed with paraformaldehyde. These data indicate that standard VACV strains are capable of inducing direct or indirect DC activation if virus gene expression is prevented, and suggest that standard VACV strains, but not MVA, express one or several genes that prevent activation. It should be noted that poxviruses belonging to the genus Avipoxvirus have been shown to activate human DCs (Ignatius et al., 2000) and murine DCs (Brown et al., 2000), a finding that may be correlated with the lack of virus functions that interfere with the activation pathway in mammalian cells. However, it is not yet known whether the avipoxviruses activate DCs in the absence of virus gene expression.
What might be the mechanism that enables MVA to activate DCs? MVA infection was found to induce NF-
B activation in human embryonic kidney cells, while a replication-competent VACV strain or cowpox virus inhibited TNF--induced NF-B activation (Oie & Pickup, 2001). Relevant to these findings, it has been reported that NF-B activation was essential for DC activation and antigen presentation, as both are prevented by overexpression of the natural NF-B inhibitor, IB (Yoshimura et al., 2001). Taken together, these data suggest that MVA could activate DCs via an NF-B-dependent mechanism. Interestingly, the VACV A52R ORF, which is deleted from the MVA genome (Antoine et al., 1998), has been shown to belong to the IL1/Toll-like receptor family and to interfere with the activation of NF-B by a number of effectors (Bowie et al., 2000; Harte et al., 2003). However, our data show that deletion of this gene from the MVA genome cannot be the only critical factor that allows DC activation, as the recombinant VACV-Cop encoding GFP used in this work was also deleted in A52R yet was unable to activate DCs.
Although DCs infected with MVA or VACV-Cop were able to stimulate the proliferation of autologous T lymphocytes equally well, when the cells were derived from individuals vaccinated against smallpox (not shown) we could not detect a significant boost in the number of IFN--secreting lymphocytes using infected DCs as stimulators. On the other hand, we report here a very effective induction of IFN--secreting lymphocytes by a mixture of MVA-infected HeLa cells and DCs, which was much higher than if VACV-Cop-infected HeLa cells were employed. These results highlight the efficiency of the cross-presentation pathway for the stimulation of IFN--producing T cells, and presumably cytotoxic T cells, compared with the direct presentation pathway. Other studies have documented the ability of DCs to cross-present antigens from VACV-infected cells (Larsson et al., 2001; Ramirez & Sigal, 2002). These studies employed cells that were non-permissive for VACV replication (monocytes, macrophages, DCs), a situation that may closely mimic MVA infection of HeLa cells. Clearly the stimulation of T lymphocytes with a mixture of VACV-Cop-infected HeLa cells and DCs is very poor, and this may be explained by the continual production of virus in HeLa cells which will then inhibit DC activity. We can speculate from these findings that, under circumstances of in vivo infection with replication-competent VACV, non-permissive cells are the most important for transferring antigens to DCs, in much the same way as MVA-infected HeLa cells are able to do so in vitro. Finally, our data also help explain why a non-replicating virus such as MVA is an efficient vaccine strain provided high input titres are used.
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ACKNOWLEDGEMENTS |
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Received 29 January 2004;
accepted 29 April 2004.
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  Z. Waibler, M. Anzaghe, T. Frenz, A. Schwantes, C. Pohlmann, H. Ludwig, M. Palomo-Otero, A. Alcami, G. Sutter, and U. Kalinke Vaccinia Virus-Mediated Inhibition of Type I Interferon Responses Is a Multifactorial Process Involving the Soluble Type I Interferon Receptor B18 and Intracellular Components J. Virol., February 15, 2009; 83(4): 1563 - 1571. [Abstract] [Full Text] [PDF]
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 D. Teoh, L. A. Johnson, T. Hanke, A. J. McMichael, and D. G. Jackson Blocking Development of a CD8+ T Cell Response by Targeting Lymphatic Recruitment of APC J. Immunol., February 15, 2009; 182(4): 2425 - 2431. [Abstract] [Full Text] [PDF]
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Z. Waibler, M. Anzaghe, H. Ludwig, S. Akira, S. Weiss, G. Sutter, and U. Kalinke Modified Vaccinia Virus Ankara Induces Toll-Like Receptor-Independent Type I Interferon Responses J. Virol., November 15, 2007; 81(22): 12102 - 12110. [Abstract] [Full Text] [PDF]
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H. Yang, T. Dong, E. Turnbull, S. Ranasinghe, B. Ondondo, N. Goonetilleke, N. Winstone, K. di Gleria, P. Bowness, C. Conlon, et al. Broad TCR Usage in Functional HIV-1-Specific CD8+ T Cell Expansions Driven by Vaccination during Highly Active Antiretroviral Therapy J. Immunol., July 1, 2007; 179(1): 597 - 606. [Abstract] [Full Text] [PDF]
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L. Deng, P. Dai, W. Ding, R. D. Granstein, and S. Shuman Vaccinia virus infection attenuates innate immune responses and antigen presentation by epidermal dendritic cells. J. Virol., October 1, 2006; 80(20): 9977 - 9987. [Abstract] [Full Text] [PDF]
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 M. G. Cottingham, A. van Maurik, M. Zago, A. T. Newton, R. J. Anderson, M. K. Howard, J. Schneider, and M. A. Skinner Different Levels of Immunogenicity of Two Strains of Fowlpox Virus as Recombinant Vaccine Vectors Eliciting T-Cell Responses in Heterologous Prime-Boost Vaccination Strategies. Clin. Vaccine Immunol., July 1, 2006; 13(7): 747 - 757. [Abstract] [Full Text] [PDF]
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) or VACV-Cop (
) and cultured in the presence of T lymphocytes overnight. (b) HeLa cells were infected with MVA (