1 Department of Virology, Biomedical Primate Research Centre, Lange Kleiweg 139, 2288 GJ Rijswijk, the Netherlands
2 Robert Koch Institut, Nordufer 20, D-13353 Berlin, Germany
3 GSF-Institut für Molekulare Virologie, Trogerstrasse 4b, 81675 München, Germany
4 Karolinska Institute, PO Box 280, SE-171 77 Solna, Sweden
Correspondence
Gerrit Koopman
koopman{at}bprc.nl
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ABSTRACT |
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Present address: Paul-Ehrlich-Institute, Langen, Germany.
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INTRODUCTION |
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METHODS |
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Vaccines.
For DNA-expression-based vaccines, the following vectors were used: pTH.UbgagPk, pTH.UbpolPk, pTH.UbnefPk, pTH.rev and pTH.tat; all express the gag, pol, nef, rev and tat genes of SIVmacJ5 under the control of the human cytomegalovirus immediateearly (HCMV IE) enhancer/promoter (Hanke et al., 1998). Vector pND14-G4 contained the SIVmac251 envelope gp120 coding sequence under the control of the HCMV IE enhancer/promoter. All constructs also contained the HCMV intron A sequence 5' of the expressed genes, in order to increase expression from the HCMV enhancer/promoter sequence, and carried the bovine growth hormone (BGH) poly(A) signal/terminator sequence (Rhodes et al., 1994
; Hanke et al., 1998
). Vector control animals were immunized with empty pTH.
MVA recombinants used in this study expressed the Gag, Pol, Nef, Rev, Tat and envelope gp160 proteins of SIVmacJ5 under control of the vaccinia virus earlylate promoter P7.5 (Nilsson et al., 2001; Horton et al., 2002
; Vogel et al., 2002b
). Vector control animals were immunized with wild-type MVA.
SFV recombinants used in this study expressed the Gag, Pol, Nef, Rev, Tat and envelope gp160 proteins of SIVmacJ5. Genes were expressed in general-expression vectors based on the SFV replicon (Nilsson et al., 2001). Recombinant RNA molecules were transcribed from the inserted genes and subsequently packaged into suicide SFV particles for use as a vaccine. Vector control animals were immunized with SFV-LacZ.
SIVmac251 virus stock.
Challenge virus stock was prepared from the supernatant of peripheral blood mononuclear cells (PBMCs) from rhesus macaques that were infected with SIVmac251 (kindly provided by Dr Aubertin, Strasbourg, France). Rhesus macaques were inoculated intrarectally by using tenfold dilutions from 101 to 104, with four rhesus macaques per dilution apart from the 104 dilution, for which three monkeys were used. The resulting number of infected animals in each group was 4, 4, 2 and 0, respectively, giving an intrarectal titre of 103 MID50 ml1.
Immunization and challenge schedule.
The study comprised three groups of animals. Six animals received the DNA, MVA and SFV vectors expressing the SIV proteins Gag, Pol, Nef, Rev, Tat and Env; four animals received the empty DNA, MVA and SFV vectors (vector controls); two animals were not immunized (naïve controls). Animals received four immunizations at 8-weekly intervals, starting with the DNA vector, followed by MVA, then SFV and finally a second MVA immunization. DNA immunization (100 µg per construct) was performed as a total of six intradermal injections, 100 µl per injection site (100 µg DNA), proxolateral from both inguinal regions. MVA immunization (1x108 p.f.u. per construct) was given as five intramuscular injections, 500 µl per injection site (1x108 p.f.u. per site). SFV (1x108 p.f.u. per construct) was given as two subcutaneous injections, 1000 µl per injection site (3x108 p.f.u.), proxolateral from both inguinal regions. Eight weeks after the last immunization, all animals were challenged by intrarectal administration of 50 MID50 of the pathogenic SIVmac251 stock. Fasted monkeys were sedated by ketamine injection and laid on their stomachs with the pelvic region slightly elevated. A feeding tube was inserted 4 cm into the rectum and 3 ml virus diluted 1 : 60 in sterile, pyrogen-free RPMI 1640 containing 20 % inactivated fetal calf serum (FCS) was delivered slowly.
Cellular immunology assays.
Th cell responses were determined by a standard [3H]thymidine incorporation assay, performed 4 weeks after each immunization and after challenge, as described by Verschoor et al. (1999). Briefly, PBMCs were cultured in RPMI 1640 medium supplemented with 5 % heat-inactivated FCS in U-shaped, 96-well microtitre plates at a concentration of 2x105 cells per well using SIVmac251 rgp130 (NIBSC, EVA655), SIVmac251 rGag (NIH catalogue no. 1845), SIV J5Nef (NIBSC, ARP668) and SIV-Tat (NIBSC, ARP681) (5 µg ml1) to stimulate proliferation. Concanavalin A stimulation (5 µg ml1) was used as a positive control. Cells were incubated for 90 h. During the last 18 h, cells were pulsed with 2·5 µCi [3H]thymidine per well. Subsequently, cultures were harvested on glass-fibre filters and label uptake was determined by counting simultaneously in an open-well Packard Matrix counter (direct beta-counter) with 96 counting tubes. Stimulation indices (SIs) were calculated by dividing the mean c.p.m. of antigen-stimulated wells by the mean of the unstimulated wells. An SI of >3·0 was considered to be positive.
Induction of gamma interferon (IFN-), interleukin 2 (IL2) and IL4 cytokine responses was measured by using an ELISpot assay, performed 4 weeks after each immunization and after challenge (Verschoor et al., 1999
). In brief, 4x106 PBMCs ml1 were cultured in a 24-well tissue culture plate for 24 h in RPMI 1640 medium supplemented with 5 % pooled rhesus serum, using SIVmac251 rgp130, SIVmac251 rGag, SIV J5Nef and SIV-Tat (5 µg ml1) to stimulate cytokine production. Phorbol myristate acetate (PMA; 20 ng ml1) plus ionomycin (1 µg ml1) stimulation was used as a positive control. Two weeks after each immunization, IFN-
ELISpot responses against a panel of peptides that was selected to cover the immunodominant epitopes of SIV Tat, Rev, Nef and Gag were measured. Peptide pools were composed as follows: Tat pool 1 (EVA7069.15), Tat pool 2 (EVA7069.610), Rev pool 1 (EVA7068.14), Rev pool 2 (EVA7068.58), Nef pool 1 (EVA7067.15), Nef pool 2 (EVA7067.610), Nef pool 3 (EVA7067.1115), Nef pool 4 (EVA7067.1620), Nef pool 5 (EVA7067.2125), Nef pool 6 (EVA7067.2629), Gag pool 1 (EVA7066.14), Gag pool 2 (EVA7066.58), Gag pool 3 (EVA7066.912) and Gag pool 4 (EVA7066.1316). For enumeration of antigen-specific cytokine production, non-adherent cells were collected and plated at 2x105 cells per well in a 96-well ELISpot plate with the same antigens added. Microtitre plates were pre-coated with mAbs that were specific for the lymphokine of interest, i.e. anti-IFN-
mAb MD-1 (Ucytech), anti-IL4 mAb QS-4 (Ucytech) and anti-IL2 mAb B-G5 (Diaclone Laboratories).
Detection of cytokine-secreting cells took place after either 15 h for IL4 or 4 h for IFN- and IL2. Cells were lysed and debris was washed away before adding detector antibodies. IFN-
, IL2 and IL4 were detected by using biotinylated rabbit anti-rhesus IL2, biotinylated rabbit anti-rhesus IFN-
or biotinylated mouse anti-rhesus IL4. Spots were visualized by using a gold staining/silver enhancement technique (Ucytech). IFN-
, IL2 or IL4 ELISpot results are expressed as spot-forming cells per 106 PBMCs minus background (mean of medium control+2SD). The assay was discarded if PMA/ionomycin stimulation gave no response.
Determination of virus load.
A quantitative competitive RNA-PCR was used to estimate the virus load in plasma, as described by ten Haaft et al. (1998). For determining the cell-associated virus load, PBMCs or mononuclear cells isolated from peripheral lymph nodes, mesenteric lymph nodes or spleen were cultured in a three- or fourfold dilution range with C8166 indicator cells for 36 weeks. PBMCs taken at week 2 after challenge were plated in a range from 250 000 to 250 cells per well by using fourfold dilutions. For the two non-infected macaques (Ri404 and Ri437), 107 cells were plated over 24 wells. PBMCs taken at week 31 after challenge were plated in a range from 500 000 to 2000 cells per well by using threefold dilutions. From four macaques with low or undetectable plasma virus loads (Ri404, Ri406, Ri421 and Ri437), 5x106 PBMCs were plated over 10 wells. Medium was renewed twice a week. Positive cultures were scored via cytopathic effect formation, which was confirmed by RT-PCR.
Intracellular IFN- and IL2 staining.
PBMCs (5x106 ml1) were incubated at 37 °C for 2 h with anti-CD28 and anti-CD49d antibodies (2 µg each antibody; BD Pharmingen) and either staphylococcal enterotoxin B (1·25 µg ml1; Sigma), pooled peptides (1·25 µg each peptide per sample) or SIV-1 Env protein (1·25 µg ml1). Peptide pools for induction of intracellular cytokines were composed as follows: Tat pool (EVA7069.110), Rev pool (EVA7068.18), Nef pool (EVA7067.129) and Gag pool (EVA7066.116).
Cells were treated with brefeldin A (GolgiPlug 1 : 1000; BD Pharmingen) to inhibit protein trafficking and incubated for 16 h at 37 °C. Cells were then washed with PBS/1 % BSA and stained for surface markers by using fluorescein isothiocyanate (FITC)-labelled anti-CD8 (DAKO) and peridinin chlorophyll protein (PerCP)-labelled anti-CD4+ (clone L200; BD Pharmingen) for 30 min at 4 °C in the dark. Subsequently, cells were washed with PBS/BSA and fixed with cytofix/cytoperm solution (BD Pharmingen) for 20 min at 4 °C. The cells were then washed twice with permeabilization buffer (diluted tenfold in water) and resuspended in permeabilization buffer containing phycoerythrin (PE)-labelled anti-IL2 and allophycocyanin (APC)-labelled anti-IFN- mAb. After 30 min incubation at 4 °C, cells were washed twice with permeabilization buffer and fixed in 2 % paraformaldehyde in PBS for 16 h. Acquisition was performed on a FACSort flow cytometer collecting 100 000200 000 lymphocyte-gated events per sample.
FACS subset analysis.
Macaques were monitored for changes in their T-lymphocyte subsets by flow cytometry analysis, as described by Koopman et al. (2001). Briefly, 100 µl EDTA-treated blood was incubated with 10 µl mAb mix in 5 ml polystyrene, round-bottom tubes (Falcon 2058; Becton Dickinson) at room temperature for 15 min. After this incubation, 1·5 ml lysing solution (Becton Dickinson) was added, followed by incubation at room temperature for 10 min and then centrifugation for 10 min at 500 g. The supernatant was aspirated and the cells were resuspended in 5 ml PBS with 12 % formaldehyde and stored overnight at 4 °C. Flow cytometry was performed on a FACSort using CellQuest software (Becton Dickinson). The following mAbs were used: (a) CD3FITC, CD16PE, CD8PerCP, CD4APC; (b) HLA-DRFITC, CD20PE, CD8PerCP, CD4APC and (c) CD45RAFITC, CD62LPE, CD8PerCP, CD4APC. These mAbs were obtained from BD Pharmingen (CD8PerCP clone SK1, CD4APC clone SK3, CD62LPE clone SK11, HLA-DRFITC clone L243, CD20PE clone L27, CD3eFITC clone SP34, CD16PE clone 3G8) or Diaclone (CD45RAFITC clone B-C15).
Humoral responses.
Anti-SIV antibody titres were determined by using a standard whole-SIVmac ELISA. Briefly, serially diluted plasma was assayed in duplicate in microtitre plates that were coated with whole SIVmac lysate. After incubation with anti-human IgG peroxidase conjugate (Sigma)/substrate solution and absorbance measurement, curves were fitted to the data points and used to calculate individual titres, i.e. the dilution at which the curves crossed the cut-off absorbance. Neutralizing antibodies were measured as described by Norley et al. (1996). Briefly, serial dilutions of SIVmac251 (six replicates) were incubated with a 1 : 100 dilution of plasma before addition of C8166 cells. After incubation for 7 days, wells were tested for SIVmac Gag p27 by using an antigen-capture ELISA and the virus titres (TCID50) determined. Yield reduction for each sample was calculated as the titre of virus in the absence of plasma divided by the titre in the presence of plasma.
Statistical analysis.
An unpaired t-test was used to compare virus load between the group of six vaccinated animals and the four control animals that became infected. For reasons explained in Results, the two non-infected animals were excluded from the analysis. Statistical analysis was performed for peak virus load, measured at week 2 after infection, and steady-state plasma virus load, measured at week 10 after infection. In order to study the effect of vaccination on the persistence of CD4+ T cells, the change in CD4+ T-cell count observed between the time before infection and week 31 after infection was calculated in the group of vaccinated animals, as well as in the group of control animals that became infected. Both within the vaccine group and within the control group, statistical significance of the change in CD4+ T-cell count was calculated by using Student's paired t-test. An unpaired t-test was used to compare anti-SIV antibody levels and SIV-neutralizing antibody levels, measured at week 20 after infection, between the group of six vaccinated animals and the four control animals that became infected.
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RESULTS |
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In order to establish whether CD4+ Th1 cells or CD8+ cytotoxic cells were responsible for the IFN- production seen in the ELISpot assay, an intracellular cytokine assay was performed (example shown in Fig. 1
). As shown in Table 2
, in agreement with the ELISpot results, Gag-specific IFN-
production was seen in three of the six vaccinated macaques (Ri420, Ri445 and Ri447). Importantly, only CD4+ cells produced IFN-
. In contrast to the ELISpot data, no response was seen against SIV Env. However, as no peptides were available for Env, we used the protein in this assay and this may have been less optimal than the peptides. Strikingly, macaque Ri408 showed Nef-specific IFN-
production, both by CD4+ and by CD8+ T-cells (Table 2
). Although such a response was not seen in the ELISpot assay on this macaque (Table 1
), it must be stated that it gave a high background, which may have precluded detection. None of the control animals showed a response. The proportion of positive cells in the medium control was below 0·05 %.
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DISCUSSION |
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In this study, vaccine efficacy was tested by intrarectal challenge with the highly pathogenic SIVmac251. This model is thought to be highly relevant to HIV infection in humans because: (i) most humans get infected via mucosal exposure; (ii) as with HIV-1 in humans, it typically leads to a gradual depletion of CD4+ T cells, followed by the onset of opportunistic infections and development of AIDS; and (iii) as with HIV, it is difficult to raise neutralizing antibodies to the virus. All six vaccinated animals became infected, as well as two of the four vector control animals and both the untreated, naïve-control animals, as determined by quantitative RT-PCR performed on plasma samples. These results were confirmed by DNA PCR on PBMCs, as well as a by co-culture of PBMCs with C8166 indicator cells. In order to confirm the virus-negative status of the two PCR-negative animals, DNA PCR and co-culture assays were also performed on peripheral lymph nodes, mesenteric lymph nodes and spleen mononuclear cells that were obtained at autopsy. In addition, no cellular or humoral responses were found in these animals (Table 3 and Fig. 4
). With mucosal challenge, the virus needs to cross several barriers in order to reach the immune cells that are susceptible to infection. In addition, the innate immune system may provide ways to neutralize the virus via soluble factors as well, e.g. the activity of natural killer cells. These factors, which are still far from understood, may be the reason that mucosal challenges are generally far less reliable than an intravenously applied virus challenge. The virus stock used in this study has been tested in several other studies, where 25 of 31 control animals became infected, underscoring the fact that failed challenges do occur.
If the two virus-negative animals were considered to represent failed challenges' and only the infected animals were compared, then the group of vaccinated animals showed a somewhat lower peak virus level in plasma 2 weeks after challenge than the control group. However, at later time points when steady-state virus levels were reached, plasma virus levels in vaccinated, as well as control, animals varied between 500 and 300 000 copies ml1 and no clear differences were seen. Strikingly, the three animals with an IL4 response were less effective at controlling virus load (Table 1, Fig. 2
). The two animals with a typical Th1-type response were among the three animals that had a relatively low steady-state plasma virus load. This would be in agreement with experiments in mice, where both the IFN-
produced by Th cells and the Th cell-mediated induction of CTL responses were shown to be important in suppression of virus replication. Importantly, the numbers of CD4+ T cells were well-maintained in the vaccinated animals (Fig. 3
), whereas all four control animals that became infected showed a gradual loss of CD4+ T cells. In contrast to previous reports (Veazey et al., 2000
; Koopman et al., 2001
), we did not observe an increase in the number of CD45RA+ CD62L+ naïve CD4+ T cells in peripheral blood after SIVmac infection (Fig. 3
). It is possible that differences in the viral isolate being used may play a role. During the course of the study, none of the animals developed AIDS symptoms and CD4+ T-cell counts remained relatively stable from weeks 16 to 32 after infection. However, in several animals, a small further decrease in CD4+ T-cell count was seen at the last time point, i.e. week 36, and disease progression may be envisaged on a longer time-course. Also, one could speculate about possible disease progression in two macaques, Ri420 and Ri418 from the vaccine group, which had a relatively low CD4+ T-cell count. However, macaque Ri408 from this vaccine group also started with a relatively low CD4+ T-cell count, but at later time points, its CD4+ cell numbers increased (Fig. 3
).
After challenge, there was a strong reduction in IL2 and IL4 production (Table 3). This may indicate that, as a result of acute infection, responses were shifted to IFN-
production. However, the high background of this assay made it difficult to confirm this supposition. A similarly high background after challenge was observed in the intracellular cytokine assay (data not shown). Surprisingly, high lymphoproliferative responses were seen after challenge in the non-vaccinated control animals that became infected. As it was shown recently that HIV-specific CD4+ T cells are particularly prone to HIV infection (Douek et al., 2002
), one could speculate that these high proliferative responses may have contributed to the decline of CD4+ T cells.
As expected, the vaccines used in this study did not induce antibodies against the virus. However, all infected animals did develop high antibody titres that were able to block SIV infection in vitro (Fig. 4). Apparently, these neutralizing antibodies are not effective in the animals as high virus loads were seen, despite the presence of these antibodies. Lack of correlation between the post-challenge levels of neutralizing antibodies in plasma and suppression of virus load is not unusual. In a study by Vogel et al. (2002a)
, animals with the lowest neutralizing antibody levels during chronic infection also had the lowest plasma virus loads. This may reflect the artificial nature of the in vitro assays that were used to measure such antibodies (T cell-adapted SIVmac and human T-cell indicator cells) or may reflect a genuine failure of neutralizing antibodies to significantly influence virus levels in infected animals.
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ACKNOWLEDGEMENTS |
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Received 26 April 2004;
accepted 23 June 2004.