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* Institute of Hygiene and Social Medicine, Innsbruck Medical University, Ludwig Boltzmann Institute for AIDS Research, and
Institute of Physiology and Balneology, Innsbruck Medical University, Innsbruck, Austria
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Abstract |
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(CCL19). Preincubation of DCs with a p38 MAPK inhibitor blocked CCR7-dependent DC migration. Migrating DCs were able to induce infection of autologous unstimulated PBLs in the Transwell system. These data indicate that HIV-1 triggers a cell-specific signaling machinery, thereby manipulating DCs to migrate along a chemokine gradient, which results in productive infection of nonstimulated CD4+ cells. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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(CCL19) induces homing of maturing DCs to LT (1).
Because HIV transmission occurs at mucosal surfaces, the virus has to be transported from the mucosal sites of infection to lymphoid tissues. There the virus is transmitted to its primary targets, CD4+ T lymphocytes. This process is thought to be mediated by DCs. To use these cells as a shuttle, HIV has to modulate DCs and induce at least their partial maturation. This process is reflected not only by altered expression of cell surface proteins, but also by a switch in the phosphorylation pattern of MAPK (2, 3). Recent studies in DCs revealed that p38 MAPK and ERK1/2 play distinct roles after stimulation with various agents, e.g., LPS, TNF-
, ribotoxic stress, or contact sensitizers (4, 5, 6, 7, 8, 9, 10). Although p38 MAPK was associated with augmentation of costimulatory molecules upon DC stimulation (6, 9), ERK1/2 were linked to induction of cytokine production and DC survival (4, 5). Also, the up-regulation of CCR7 and increased migration of maturing DCs in response to MIP-3 and SLC is mediated by the p38 MAPK pathway (11).
Recently, it was shown by Popik and Pitha (12) that binding of HIV-1 to T cells triggers a broad array of signaling pathways. HIV-1-induced signals can be modulated by concomitant engagement of other cell surface molecules, e.g., chemokine receptors or proteoglycans (12, 13). However, the specific effects of the single MAPK isoforms on HIV-1 stimulation of DCs are poorly characterized to date. In this study we correlate HIV-1-induced extracellular alterations of cell phenotype with distinct intracellular changes in DCs to establish a causal relationship between the state of DC differentiation and a defined activation pattern of both the ERK and the p38 MAPK pathways as a consequence of contact with the virus.
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Materials and Methods |
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Purified recombinant human IL-4 was obtained from PromoCell (Heidelberg, Germany), GM-CSF was purchased from Schering-Plough (Dardilly, France), and LPS (Escherichia coli serotype O26:B6) was purchased from Sigma-Aldrich (St. Louis, MO). mAbs for FACS analyses directed against cell surface markers were obtained from BD Pharmingen (San Diego, CA; CD83 and CD86), DakoCytomation (Glostrup, Denmark; HLA-DP, DQ, DR), R&D Systems (Minneapolis, MN; DC-SIGN), or eBiosciences (San Diego, CA; CCR7). The FITC-conjugated goat anti-mouse Ig polyclonal Ab was obtained from DakoCytomation.
The bicinchoninic acid protein detection kit was purchased from Pierce (Rockford, IL). The p44/42 MAPK Ab, phospho-p44/42 MAPK (Thr202/Tyr204) E10 mAb, the phospho-p38 MAPK (Thr180/Tyr182) 28B10 mAb, the anti-mouse IgG/HRP-linked Ab, the anti-rabbit IgG/HRP-linked Ab, and the MEK1/2 inhibitor U0126 were obtained from Cell Signaling Technology (Beverley, MA). The p38 MAPK assay kit (nonradioactive IP-kinase assays) was obtained from Cell Signaling Technology. PD98059 and SB202190 were from Calbiochem (San Diego, CA). SB203580 was from Sigma-Aldrich. The 3-µm pore size Transwells for the migration assays were obtained from Costar (Cambridge, MA).
Cells
PBMCs were isolated from blood of normal healthy donors (obtained from the local blood bank, Innsbruck, Austria) by centrifugation on a Ficoll-Hypaque density gradient (Pharmacia Biotech, Uppsala, Sweden). Monocytes were separated from PBMCs by adhesion to gelatin-coated petri dishes as described previously (14). Adherent cells were detached using RPMI 1640 medium (Life Technologies, Rockville, MD) supplemented with 5 mM EDTA. Monocytes were washed and cultivated in RPMI 1640/10% FCS/2 mM L-glutamine (RPMIc) containing 1500 U/ml IL-4 and 1600 U/ml GM-CSF at a density of 1 x 106 cells/ml medium in six-well plates (Costar) to generate monocyte-derived DCs. IL-4 (1000 U/ml) and GM-CSF (1600 U/ml) were added to the medium after 2 days in culture. Day 5 cells were spun, medium was replaced by fresh RPMIc containing 1000 U/ml IL-4, and 1600 U/ml GM-CSF without any further stimulation to generate iDCs with either 0.1 ng/ml (LPS low) or 100 ng/ml LPS (mDCs) or with different strains of HIV-1. For FACS analyses cells were cultivated for an additional 2 days to investigate the effects of HIV-1 on maturation of DCs. For Western blot analyses, iDCs and mDCs were incubated for the time periods indicated in Results.
Virus propagation
Virus stocks of the HIV-1 laboratory strains IIIB and BaL (obtained from the Medical Research Council AIDS Reagent Project, Herts, U.K.), infectious viral clones MC1 (X4-tropism) and MC3 (R5-tropism; donated to the Medical Research Council AIDS Reagent Project by H. Schuitemaker) and the primary isolates 93BR029 (subtype F, X4-tropism), 92UG029 (A/A, X4), 92BR030 (B/B, R5), or 92UG037 (A/A, R5) were prepared in PHA- and IL-2-prestimulated PBMCs. To prevent carryover of IL-2, PHA or proteins from heat-inactivated FCS, which may function as putative modulators of DCs, virus supernatants (SNs) were filtered through a 0.22-µm pore size filter and centrifuged at 20,000 rpm for 90 min at 4°C, and the SN was discarded to remove all putative contaminations. Pelleted intact virus particles were resuspended in RPMI 1640 without any supplement, aliquoted, stored at –80°C, and thawed only once for each experiment to ensure constant infectivity. From all preparations an aliquot was taken to determine the concentration of the ultracentrifuged virus by p24 ELISA (14) and the tissue culture 50% infective dose of the viral stock. The endotoxin levels of the virus preparations used in the experiments were below the detection limit of 0.005 U/ml (0.0005 ng/ml), and the threshold concentration for an LPS-induced effect on DCs was 0.01 ng/ml in our experiments (not shown). The activation of MAPK by putative contaminations, which may interact with TLRs, was excluded by testing the virus SN after ultracentrifugation of the virus. The weak ERK signal obtained by the SN compared with the resuspended virus pellet was probably due to HIV-1 proteins shed in the SN (not shown). To exclude that cellular vesicles or other cell-derived contaminations induce DC activation, uninfected PBMCs were included as an additional control. No effect of the mock preparations on DCs was observed (not shown).
Flow cytometry
Unstimulated and LPS- or HIV-1-stimulated day 7 DCs (1 x 106/ml) were washed with cold PBS/1% BSA/0.1% NaN3. Isotype controls (IgG1, IgG2b isotype controls; American Type Culture Collection, Manassas, VA) or cell surface-specific Abs (CD83, CD86, and DC-SIGN) were added at a concentration of 5 µg/ml for 1 h at 4°C. The CCR7 mAb was used for FACS analysis at a concentration of 25 µg/ml as recommended by the manufacturer. After washing, the FITC-conjugated, goat anti-mouse Ig Ab was added for 30 min at 4°C in the dark. Thereafter, cells were washed, fixed in PBS/4% paraformaldehyde, and incubated overnight at 4°C. The analyses were performed using a FACScan flow cytometer (BD Biosciences, Mountain View, CA) and CellQuest software (BD Biosciences).
Real-time quantitative RT-PCR
CCR7 levels of DCs after incubation with either LPS or HIV-1 were determined by quantitative real-time RT-PCR using the SYBR Green Mastermix (Stratagene, La Jolla, CA). Primers for CCR7 (gi:23243433) were designed using Primer 3 software and were as follows: CCR7 forward, 5'-cag atg caa tga ctc agg ac-3'; and CCR7 reverse, 5'-ctg ttt ccc agt gtt gtc tg-3'. A GAPDH RT-PCR (GAPDH forward, 5'-CTC ATg ACC ACA gTC CAT gC-3', GAPDH reverse, 5'-CAC gCC ACA gTT TCC Cg-3'; GAPDH probe, 5'-FAM-CAg AAg ACT gTg gAT ggC CCC-TAMRA-3') served as the control for identical RNA amounts in the preparations. Total RNA from iDCs, LPS-, and HIV-1-stimulated cells was prepared 0 h (not shown), 2 h (not shown), and 48 h postincubation (Fig. 2) using TRIzol reagent (Invitrogen Life Technologies, Gaithersburg, MD). Assay mixtures for the CCR7-specific RT-PCR contained 20 µl of 2x SYBR Green Mastermix, 200 nM of each of the specific primers, 1.25 U of StrataScript reverse transcriptase, and 5 µl of isolated RNA in a total volume of 40 µl. The GAPDH RT-PCR was performed using the single-step RT-PCR kit from Stratagene. Four microliters of 10x RT-PCR buffer, 5 mM MgCl2, 200 nM of each of the primers, 125 nM of the fluorescently labeled probe, 1.25 U of StrataScript reverse transcriptase, and 5 µl of the isolated RNA were added in a total volume of 40 µl. Thermal cycling conditions consisted of 30 min at 45°C for RT, 10 min at 95°C, 50 cycles of 15 s at 95°C and 30 s at 60°C for PCR, and melt curve analyses for specification of PCR products with SYBR Green. Real-time quantitative PCR was performed on the iCycler (Bio-Rad, Hercules, CA).
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After HIV-1 stimulation, cells were washed twice with cold PBS and lysed in ice-cold 1% Triton X-100 lysis buffer containing protease and phosphatase inhibitors for 25 min at 4°C. Insoluble material was removed by centrifugation at 12,000 x rpm for 15 min at 4°C. The protein content of the resuspended lysates was determined using a bicinchoninic acid assay as described by the manufacturer with BSA as standard. Cell lysates were matched for protein, separated on 10% SDS-PAGE, and transferred to a nitrocellulose membrane. Membranes were incubated with ERK1/2 (p44/42 MAPK) Ab to verify protein loadings, with anti-ERK1/2 Ab, or with anti-p38 MAPK Ab (Cell Signaling Technology). Binding of the primary Ab was visualized by HRP-conjugated anti-mouse or anti-rabbit IgG Ab and ECL (Pierce).
Migration and infection assays
For migration assays, day 7 DCs were incubated overnight with either LPS or HIV-1 in the absence or the presence of MAPK inhibitors U0126 and SB203580. Cells were washed to remove unbound virus or inhibitors, resuspended in 500 µl of fresh RPMIc, and transferred into the upper chamber of 3-µm pore size polycarbonate filters in 12-well Transwell chambers (Corning; Costar). The lower chamber contained 1 ml of RPMIc supplemented with 100 ng/ml MIP-3 and 100 ng/ml SLC (PeproTech EC, London, U.K.). Lower chambers with medium only served as a control for spontaneous migration. The migrated cells from the bottom chamber were spun, then half of them were fixed in PBS/4% formaldehyde and counted by flow cytometry in a FACScan, acquiring events for a time period of 60 s using CellQuest software (BD Biosciences). Experiments were performed four times to verify the HIV-1-mediated migration due to CCR7 up-regulation. The total number of cells migrated due to MIP-3/SLC was divided by the number of spontaneously migrated cells, and values are given as fold migration. The other half of the migrated cells from the bottom chamber was resuspended in RPMIc and mixed with nonstimulated, autologous PBLs at a ratio 1:5. Infection assays were performed as described below.
Infection experiments
Immature DCs and mDCs were spun, washed, and resuspended at a density of 0.5–1 x 106 cells/ml in RPMIc supplemented with IL-4 and GM-CSF. Where indicated, DCs were preincubated with the MAPK inhibitors, U0126 (5 µM), PD98059 (40 µM), SB202190 (10 µM), or SB203580 (10 µM) for 1 h, then incubated for 3 h at 37°C with 10 ng p24/ml of the various HIV-1 strains. After the incubation period, cells were washed and resuspended in RPMIc, and primary, unstimulated, autologous PBLs were added at a ratio of 5:1. Cells were cultivated at 37°C in 5% CO2, and SNs were taken up to 13 days postinfection as indicated in Figs. 4B and 5. Virus propagation was measured by p24 ELISA.
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To test whether viral replication is necessary for the signaling and maturing events in DCs, day 5 iDCs were preincubated for 2 h with 10 nM/ml nucleoside reverse transcriptase inhibitor AZT, or not, before HIV-1-addition for another 2 days (FACS) or the times indicated for Western blot experiments. Thereafter, the cells were washed and treated as described above to perform FACS or Western blot analyses. AZT (10 nM/ml) was present during the entire period of the infection assays.
Statistical analysis
Differences between surface expressions of DC activation markers were analyzed using PRISM software (GraphPad, San Diego, CA). A value of p < 0.05 in unpaired Student’s t test was considered significant.
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Results |
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Given the central role of DCs regarding initiation and regulation of immune responses (15, 16) and the fact that recombinant gp120 of HIV-1 LAV/IIIB modulates the expression pattern of several surface proteins on iDCs and mDCs (17), we first studied whether cell surface molecules associated with DC activation are modulated when iDCs are exposed to intact HIV-1 preparations.
Thus, day 5 iDCs were incubated 2 days in the presence of either X4- or R5-tropic HIV-1 primary isolates (93BR029, 92UG029, 92BR030, and92UG037), molecular clones (MC1 and MC3) or the laboratory strain HIV-1-BaL, all at a concentration of 10 ng of p24/ml. The expression pattern of characteristic DC surface markers (CD83, CD86, and DC-SIGN) on HIV-1-exposed DCs was studied by FACS analyses and compared with that of LPS-treated DCs (100 ng/ml; mDCs) or untreated iDCs. As expected, incubation of iDCs with LPS for an additional 2 days induced a statistically significant up-regulation of the maturation and differentiation markers CD83 (Fig. 1A) and CD86 (not shown). Similarly, exposure to HIV-1 or low amounts of LPS (0.1 ng/ml) induced a significant up-regulation of CD83 (Fig. 1A) or CD86 (not shown). The effect varied considerably dependent on the donor (Table I). In parallel, the Ag-capturing molecule DC-SIGN was down-regulated compared with untreated iDCs (Fig. 1B). Treatment of iDCs with both R5- and X4-tropic HIV-1 preparations resulted in a decrease in DC-SIGN expression, although this effect was not statistically significant compared with iDCs. To investigate whether viral replication is necessary for the HIV-1-mediated DC surface alterations, cells were preincubated with AZT. No major effects were observable due to treatment of iDCs with AZT before HIV-1 addition compared with DCs treated with HIV-1 only (not shown). The HIV-1-induced surface changes on iDCs were strongly donor-dependent. In 14 of 44 experiments performed, only a weak or no up-regulation of CD83 or CD86 and no down-regulation of DC-SIGN upon exposure to HIV-1 were observed.
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Next, we investigated whether HIV-1-loaded DCs up-regulate CCR7. This is of interest, because triggering surface-expressed CCR7 by its natural ligands, SLC and MIP-3, induces homing of DCs to the LT (18). Therefore, we performed a CCR7-specific real-time RT-PCR. Untreated iDCs served as controls for baseline CCR7 mRNA expression, and LPS-stimulated DCs were used as positive controls. When compared with iDCs, a significant increase in CCR7 mRNA was observable in HIV-1- and LPS-stimulated DCs 48 h after incubation (Fig. 2A). HIV-1-incubated DCs and LPS-stimulated DCs showed an increase in CCR7 mRNA expression of a 
Ct of 3.9 (Ct 21.8 in HIV-1-incubated DCs compared with a Ct of 25.7 in untreated iDCs) and of 8.8 (Ct of 16.9 in mDCs vs 25.7 in untreated iDCs), respectively (Fig. 2A). This corresponds to a >1 log increase in CCR7 mRNA expression for HIV-1- and an 
2.7 log increase in CCR7 mRNA up-regulation for LPS-exposed DCs. To control the specificity of the primers, a melt-curve analysis was performed (Fig. 2B). Comparable mRNA inputs were verified by a GAPDH RT-PCR (not shown). Ct values of 19.5–19.8 were measured for the mRNA products of the housekeeping gene of all samples (not shown). LPS- and HIV-mediated CCR7 up-regulation was furthermore confirmed by FACS analyses (not shown).
Involvement of ERK1/2 and p38 MAPK in HIV-1-mediated changes of DC cell surface markers
Because protein kinases of the MAPK family, in particular ERK1/2 and p38 MAPK, were shown to be involved in LPS- and TNF--induced DC differentiation (11, 12, 16, 19, 20), we next examined HIV-1-induced alterations of ERK1/2 and p38 MAPK activation patterns in iDCs upon short- and long-term exposure to the virus.
ERK1/2 phosphorylation in DCs
iDCs were incubated with the X4-tropic primary isolate HIV-1–93BR029 for 5 min to 4 h (short-term exposure) to investigate whether MAPKs are activated upon HIV-1 exposure. Compared with untreated iDCs, stimulation of iDCs with HIV-1–93BR029 initiated a weak, but reproducible, increase in ERK1/2 phosphorylation after 5 min (Fig. 3A). The ERK1/2 phosphorylation was most prominent 15 min after HIV-1 incubation and remained slightly elevated up to 4 h (Fig. 3A). Although the HIV-1-induced ERK1/2 phosphorylation signal was completely blocked after preincubation of iDCs with the specific MEK1/2 inhibitors U0126 (Fig. 3A) or PD98059 (not shown), a significantly enhanced ERK2 phosphorylation was observed, when the p38 MAPK pathway was blocked with SB203580 before short-term stimulation with HIV-1 (Fig. 3A). A similar ERK2 activation pattern was also obtained after stimulation of iDCs with other X4- or R5-tropic HIV-1 strains (not shown). In additional experiments, the phosphorylation signals obtained in iDCs and mDCs upon HIV-1 stimulation were compared. Basal ERK2 phosphorylation in the absence of the virus was higher in iDCs, whereas in unstimulated mDCs no ERK2 phosphorylation was detected (Fig. 3B). Both R5- and X4-tropic (not shown) HIV-1 strains initiated a strong induction of ERK2 phosphorylation in iDCs, but no ERK1/2 signal was obtained in mDCs after short-term exposure to the virus.
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p38 MAPK phosphorylation in DCs
Basal p38 MAPK phosphorylation was increased in iDCs and mDCs after short-term incubation with X4-tropic (Fig. 3D) and R5-tropic (not shown) HIV-1, respectively. Preincubation of iDCs and mDCs with the specific MEK1/2 inhibitor U0126 before HIV-1 treatment did not alter the HIV-1-induced p38 MAPK signal (Fig. 3D). The p38 MAPK pathway inhibitors, SB203580 and SB202190, blocked p38 MAPK activity in iDCs and mDCs at concentrations of 50–100 µM as analyzed by nonradioactive p38 MAPK enzymatic activity assay (not shown).
In long-term exposure experiments, the p38 MAPK phosphorylation signal obtained upon incubation of iDCs with X4- and R5-tropic HIV-1 was similar to that observed after LPS activation, but increased, compared with that in unstimulated iDC (Fig. 3E).
Inhibition of p38 MAPK activity and simultaneous induction of ERK1/2 is associated with suppression of HIV-1-induced up-regulation of activation markers
In the next step, we elucidated the roles of ERK1/2- and p38 MAPK-signaling cascades for the differentiation process of iDCs. Thus, iDCs were treated with the p38 MAPK inhibitors, SB203580 and SB202190, or the MEK1/2 inhibitors, U0126 and PD98059, before incubation of cells with HIV-1, and expression of cell surface markers was again monitored by FACS analyses. Preincubation of iDCs with SB203580 or SB202190 before HIV-1 exposure kept DCs in an iDC state. Under these circumstances neither up-regulation of CD83 (Fig. 4) or CD86 (not shown) nor alterations in DC-SIGN surface expression (not shown) were observed. In contrast, preincubation of iDCs with U0126 or PD98059 before HIV-1 treatment produced no major differences in the phenotypic changes compared with the pattern obtained after incubation with HIV-1 alone (Fig. 4). The inhibitors U0126 and PD98059 by themselves did not stimulate CD83 or CD86 up-regulation in iDCs (not shown). Additional FACS analyses with X4- and R5-tropic primary isolates, molecular clones, as well as laboratory strains further confirmed these results (not shown).
HIV-1 promotes migration of DCs
Given the fact that HIV-1 triggers iDCs to express more CD83 and CD86, to influence ERK1/2 and p38 MAPK signaling, and to up-regulate CCR7, we next examined whether these alterations are associated with the ability of the cells to migrate. For this, we performed chemotaxis assays in a Transwell chamber with DCs matured with LPS as a positive control or treated with R5- and X4-tropic primary HIV-1 isolates in the absence and the presence of MAPK inhibitors (SB203580 and SB202190 for p38 MAPK inhibition, and U0126 and PD98059 for MEK1/2 inhibition). As controls for spontaneous migration, iDCs were included in the studies. MIP-3 and SLC induced a 9.5-fold migration of LPS-stimulated DCs (Fig. 5A), a 2.7-fold migration of R5-tropic, HIV-1-treated (Fig. 5A), and a 4.7-fold migration of X4-tropic HIV-1-loaded DCs (Fig. 5A). Spontaneous, but not MIP-3- and SLC-dependent, migration was observed when iDCs were either left untreated or preincubated with SB203580 before HIV-1 addition (Fig. 5A). In contrast, U0126 preincubation only slightly reduced the MIP-3/SLC-induced migration of HIV-1-loaded DCs (Fig. 5A). This demonstrated that p38 MAPK activation is a prerequisite for HIV-1-induced, CCR7-driven migration of DCs, whereas ERK1/2 do not have any impact on HIV-1-initiated DC migration.
Migrated HIV-1-loaded DCs infect CD4+ cells in cocultures
We furthermore investigated whether HIV-1-triggered DCs were able to promote infection of nonstimulated PBLs in cocultures after migration. HIV-1-loaded DCs that migrated to the lower chamber of the Transwell without addition of a chemokine induced a weaker and delayed infection of unstimulated CD4+ cells in the cocultures (Fig. 5B, lower panel) compared with chemokine-migrated DC/PBL cocultures (Fig. 5B, upper and lower panels). This delayed infection in the absence of chemokines was due to spontaneously migrated cells. As shown in Fig. 5B, infection of unstimulated CD4+ cells was observable in both X4-tropic (upper panel) and R5-tropic (lower panel) HIV-1-loaded DCs that migrated to the lower Transwell chamber upon MIP-3/SLC stimulation.
As mentioned above, SB203580-pretreated, HIV-1-exposed DCs showed MIP-3/SLC-independent spontaneous migration. The same amounts of cells were counted in the case of no addition of chemokines and in the presence of chemokines in the lower chambers of SB203580-preincubated DCs, thus resulting in similar infection rates in DC/PBL coculture experiments in both cases (not shown).
p38 MAPK inhibition increases infectivity of DCs in coculture experiments
To further investigate the roles of ERK1/2 and p38 MAPK, iDCs and mDCs were incubated with HIV-1 primary isolates and HIV-1 MC1 and MC3 for 3 h at 37°C in the presence and the absence of the MAPK inhibitors, SB202190, SB203580, U0126, and PD98059. In addition, experiments were performed in the presence of AZT. After the incubation period, HIV-1-loaded DCs were washed twice to remove any inhibitors or unbound virus, and nonstimulated, autologous PBLs were added at a ratio of 5:1. As shown in Fig. 6, strong p24 production was measured in SNs of DC/PBL cocultures on day 4 (Fig. 6). Infection was statistically significantly enhanced (p < 0.001) when DCs were preincubated with the p38 MAPK inhibitors SB203580 (Fig. 6) and SB202190 (not shown) before HIV-1 stimulation. At later time points, the differences were not as pronounced as on day 4. This indicates that arresting DCs in an immature state due to p38 MAPK inhibition and simultaneous induction of enhanced ERK1/2 phosphorylation promotes a strong augmentation of the potential of HIV-1-loaded DCs to infect CD4+ cells in coculture. This effect was observed in 14 independent experiments performed in triplicate. Blocking p38 MAPK in mDCs also resulted in a stronger productive infection in cocultures compared with cocultures in which no p38 MAPK inhibitors were added before HIV-1 addition (not shown). In contrast, pretreatment of iDCs with the MEK1/2 inhibitor U0126 before HIV-1 addition did not result in a stronger infection potential of the APCs in coculture experiments. In all experiments performed, the p24 amounts in cocultures of DCs, preincubated with U0126, were either comparable to those infected with HIV-1 alone (not shown) or weaker (Fig. 6). When DCs were preincubated for 2 h with AZT before HIV-1 addition, a very weak infection was observed 10 days postinfection (not shown). AZT was present during the course of the infection experiment. X4-loaded as well as R5-loaded (not shown) DCs were able to infect nonstimulated CD4+ cells in coculture experiments. Productive infection of CD4+ cells was more pronounced in iDC-PBL cocultures than in mDC-PBL cocultures (not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and SLC. The migrated DCs were able to transmit HIV-1 to autologous PBL. These phenomena were observed in two-thirds of 30 individuals tested. Although a recent report suggests that HIV-1 cannot induce maturation of DCs (21), the interpretation of the data described in that paper is hampered by the fact that the positive control (LPS) was unable to trigger the expression of DC maturation markers. By contrast, our findings indicate that the effects described above were HIV-1 specific, because the endotoxin level of the virus preparations used in the experiments was below the detection limit of 0.005 U/ml (0.0005 ng/ml), and the threshold concentration for an LPS-induced effect on DCs was 0.01 ng/ml in our experiments (not shown). Investigations by Fonteneau et al. (13) suggest that the HIV-1-induced maturation of myeloid DCs is not due to a direct interaction of HIV-1 with these cells, but rather a bystander effect of plasmacytoid DCs, which mature after contact with HIV-1. However, the myeloid DCs used in these experiments were already partially matured, because 25–75% of the cells expressed CD83. Thus, the activation of myeloid DCs by HIV-1 was probably not detectable in this study. This interpretation is supported by the finding that in iDCs that showed partial CD83 up-regulation before stimulation, no additional increase in CD83 expression upon exposure to HIV-1 was detectable in our system (not shown). In our assays <10% of untreated iDCs expressed detectable amounts of CD83. In a recent publication, Granelli-Piperno et al. (22) used DCs, which were comparable with our cells with regard to the baseline CD83 expression of iDCs. Upon stimulation with HIV-1, no or only minimal up-regulation of CD83 was found. The discrepancy with our findings is presently unclear and may reflect differences in the donors (nonresponders?) or the viral strains used. The lack of DC-LAMP expression, a marker for full maturation of DCs (22), also reflects the fact that HIV induces only partial activation of the cells. This is additionally supported by an HIV-1-dependent, moderate up-regulation of CCR7 mRNA and protein expression in FACS analyses (not shown) compared with LPS-stimulated DCs and indicates that HIV-1 slightly activates iDCs, but does not induce their full maturation. The activation of DCs was independent of infection, because the up-regulation of different surface markers was also observed in the presence of AZT or X4-tropic virus (not shown), which is unable to infect iDCs (23). Thus, exposure to HIV seemed to be sufficient to activate DCs. This may provide one explanation of why relatively high amounts of DCs become activated, but only a few cells are infected (22).
In general, viral infections induce cellular phosphorylation events to initiate effective replication. In recent studies several viruses were shown to activate MAPKs, which act as a central pathway in the signaling network of the host cell (12, 19, 20, 24, 25). The involvement of MAPKs in DC maturation and differentiation upon stimulation with various agents (LPS, TNF-, contact sensitizers, and ribotoxic stress) (4, 5, 8, 9) was recently described. As shown in these studies, p38 MAPK inhibition blocked the DC maturation induced by these agents, indicating that p38 MAPK phosphorylation is a prerequisite for DC activation. In contrast, an involvement of ERKs seemed to be important for DC survival, but not for DC maturation, in the mouse system (4). Our study tried to elucidate the correlation of HIV-1-mediated induction of intracellular signaling pathways with an altered phenotype and function of responding human DCs. In initial short-term experiments, R5- and X4-tropic HIV-1 isolates activated ERK1/2 in iDCs, but not in mDCs. The phosphorylation of these ERK isoforms was most prominent early after addition of the virus preparation. Also, short-term HIV-1 exposure induced an enhancement of p38 MAPK phosphorylation in both iDCs and mDCs. Because phenotypical alterations and migration induced by either HIV-1 or LPS were measured after extended exposure times, we investigated the MAPK phosphorylation pattern in long-term experiments. Untreated iDC control cells showed a relatively high basal ERK1/2 phosphorylation, whereas in mDCs no basal ERK1/2 activation was observable. In contrast to p38 MAPK phosphorylation, ERK1/2 phosphorylation was decreased in long-term HIV-1-exposed DCs, causing an intermediate signaling pattern between iDCs and mDCs. This corresponded to the up-regulation of CCR7 mRNA. In contrast, basal p38 MAPK phosphorylation was detectable in both iDC and mDC control cells. Activated p38 MAPK, which is associated with DC activation, was slightly higher in LPS-matured mDCs compared with iDCs and was also inducible by long-term exposure to HIV-1. Pretreatment of iDCs with the specific MEK1/2 inhibitors, U0126 and PD98059, blocked HIV-1-mediated phosphorylation of ERK1/2 in short as well as long-term experiments, but had no major inhibitory effects on the up-regulation of costimulatory molecule expression with regard to HIV-1-induced DC activation. On the contrary, the p38 MAPK inhibitor SB203580 significantly enhanced HIV-1-initiated ERK1/2 phosphorylation in short- and long-term experiments. Stimulation, however, was not seen vice versa when the ERK pathway was blocked, and the p38 MAPK signal was measured. FACS analyses revealed that blocking p38 MAPK by SB203580 or SB202190 before HIV-1-addition to DCs completely suppressed HIV-1-mediated DC activation, reinforcing the idea that p38 MAPK was involved in this step. p38 MAPK was recently associated with TNF--, contact sensitizer-, or LPS-induced up-regulation of DC maturation and DC costimulation and migration markers such as CCR7 (9). In our migration assay, a moderate CCR7 up-regulation was sufficient to induce MIP-3/SLC-dependent migration. Its inhibition by SB203580, but not by MEK1/2 blockers, clearly indicated that p38 was involved in this process, whereas ERK inhibitors had no effect. HIV-1-mediated migration of DCs may mimic the in vivo situation, because HIV-1 is thought to be transported by DCs to the LT, where it promotes infection of T cells. In contrast to the combined migration-infection experiments, which were dependent on p38 MAPK activation, infection of PBLs by HIV-1-loaded DCs in classical coculture experiments was enhanced when p38 MAPK was inhibited. Thus, keeping DCs in an immature state clearly favors the productive infection of DCs. The enhanced activation of the ERK pathway by SB203580 may additionally favor DC proliferation and inhibition of apoptosis and thereby additionally contribute to virus replication. This is in line with previous reports that iDCs, but not mDCs, can be productively infected with HIV (23, 26, 27).
It is interesting to note that only in two-thirds of 30 individuals tested, the p38 signal was triggered in DCs. This was not due to differences in viral strains or tropism, because with the same viral preparation in parallel experiments, DC activation was observed in one donor, whereas the second donor failed to respond to exposure to HIV-1. This may also exclude the involvement of TLRs, which were recently described to bind ssRNAs (28, 29) and other putative contaminations, such as LPS. As reported by others, prestimulated DCs could not be additionally triggered by HIV-1 (22) (our unpublished observations). Thus, nonresponders may have already undergone prior stimulations that are not detectable by FACS analyses. Alternatively, responders or nonresponders may be defined by putative polymorphisms of extracellular DC molecules, which trigger p38 MAPK upon exposure to HIV-1. Such surface proteins were previously described for T cells or macrophages (12, 13). Activation of the MAPK pathway is likely to be dependent on interaction of the viral envelope protein(s) with cellular receptors, because neither the enhanced HIV-1-induced p38 MAPK phosphorylation nor the migration of DCs was affected by the presence of AZT. In addition, it was shown recently that recombinant gp120 is sufficient to modulate the expression pattern of DCs (17)
In conclusion, this study shows that HIV-1 initiates differential MAPK signaling in DCs, which is associated with partial and moderate up-regulation of DC activation markers and the homing receptor CCR7 in two-thirds of the donors. In responders, an enhanced migratory capacity of HIV-1-loaded DCs upon MIP-3/SLC stimulation as well as infection of unstimulated autologous CD4+cells by migrated HIV-1-exposed DCs were observed. The ERK1/2:p38 MAPK ratios changed after incubation of iDCs with the virus, indicating HIV-1-mediated modulations of DCs. Stimulation of p38 MAPK and down-regulation of ERK1/2 associated with the moderate up-regulation of CCR7 may manipulate DCs to migrate to the LT. Because full maturation and, thus, Ag presentation of HIV-1-exposed DCs seem to be impaired (22), DCs are used by HIV-1 as a shuttle to the LT, where the virus may be efficiently transmitted to T cells.
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Acknowledgments |
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1 This work was supported by the Austrian Research Fond (FWF; Grants P14661 and P15379), the fifth and sixth frameworks of the European Union (MUVADEN QLK-CT-2002-00882 and TIP-VAC-012116), the Ludwig Boltzmann Institute for AIDS Research, and the federal government of Tyrol. 
2 Address correspondence and reprint requests to Dr. Heribert Stoiber, Institute of Hygiene and Social Medicine, Innsbruck Medical University, Fritz Pregl Strasse 3, 6020 Innsbruck, Austria. E-mail address: heribert.stoiber{at}uibk.ac.at
3 Abbreviations used in this paper: iDC, immature dendritic cell; AZT, zidovudine; LT, lymphoid tissue; SLC, secondary lymphoid tissue chemokine; mDC, mature dendritic cell; RPMIc, RPMI 1640/10% FCS/2 mM L-glutamine; SN, supernatant.
Received for publication June 18, 2004. Accepted for publication October 14, 2004.
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induced activation and maturation of human peripheral blood monocyte-derived DCs. J. Immunol. 162:3865.
B signal transduction pathways are involved in the survival and maturation of lipopolysaccharide-stimulated human monocyte-derived dendritic cells. Blood 96:1039., and contact sensitizers. J. Immunol. 166:3837. and SLC is mediated by the p38 stress-activated protein kinase pathway. Br. J. Hematol. 119:826.[Medline]
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