Dendritic cells (DC) are the most potent cells involved in the generation of primary and secondary immune responses. To assess the feasibility of using autologous DC as immunotherapy for HIV disease, we analyzed a variety of immune parameters using DC isolated from HIV-infected (HIV+) individuals, as well as DC obtained from HIV-uninfected (HIV) individuals infected in vitro with HIV. After stimulation with recombinant CD40 ligand (CD40LT), cytokine and β-chemokine production were similar by DC from HIV donors infected in vitro with the CCR5-using HIV Ba-L strain (n = 8) compared with uninfected DC from the same donors. Production of β-chemokines, but not of cytokines, was increased by a CXCR4-using IIIB strain-infected DC (n = 7). Stimulation of HIV-infected DC with CD40LT decreased infection in Ba-L-infected DC, but had no effect on IIIB-infected DC. Consistent with this finding, CD40LT down-regulated CCR5 and up-regulated CXCR4 expression on DC. Monocyte-derived DC were also propagated from 15 HIV+ and 13 HIV donors. They exhibited similar expression of costimulatory molecules and produced similar amounts of IL-12, IL-10, and β-chemokines, following stimulation. By contrast, stimulated PBMC from HIV+ patients exhibited decreased IL-12 and increased IL-10 production. In summary, phenotype, cytokine secretion, and β-chemokine production by DC from HIV+ individuals were normal. These cells may prove useful in boosting cellular immune responses in HIV+ individuals.

Dendritic cells (DC)2 (2) are potent immunostimulating leukocytes for both primary and secondary T cell responses (1, 2). These stimulating functions have been related to several properties: 1) immature DC can capture and process proteins and nonproteic molecules using different pathways; 2) upon activation, they migrate to the lymphoid tissues; 3) they express high levels of MHC class I and II molecules and, after further maturation, high levels of the costimulatory molecules CD40, CD80, and CD86; and 4) they produce cytokines that are essential for T cell differentiation. Moreover, DC produce high levels of chemokines that attract memory and naive T cells (3, 4, 5, 6).

Although the role of DC in the transmission of HIV to T cells has been demonstrated in vitro by several groups (7, 8, 9, 10, 11), the extent to which DC remain phenotypically and functionally unaltered during HIV infection is not clear. Given the efficiency of DC for stimulating T cell-dependent immune responses, alterations in DC number and/or function could be an important factor in the development of T cell functional impairment in HIV disease, as previously suggested (12). To address this question, our laboratories and others have studied the capacity of Langerhans cells or blood-derived DC to present recall Ag to CD4+ or CD8+ T cells and have detected no clear defect (13, 14, 15, 16, 17). Similarly, no decrease in levels of HLA-DR or costimulatory molecule expression were reported either on DC obtained from HIV-infected (HIV+) patients or after in vitro infection (14, 18, 19).

In vitro techniques are now available that permit the differentiation and expansion of DC from blood nonproliferating precursors using GM-CSF and IL-4. These monocyte-derived DC have been suggested to be good candidates for clinical use directed at enhancing immune responses in humans (2, 20). Although this approach is efficient and convenient in that it allows for experimentation on relatively large numbers of purified DC, caution must be exercised in drawing parallels between these in vitro, cytokine-driven DC and natural DC, found in nonlymphoid as well as lymphoid tissues. Mindful of this caveat, we studied cytokine and chemokine secretion of both DC from HIV donors following in vitro infection with HIV-1 and DC obtained from HIV+ donors that were at different clinical stages of HIV disease.

Production of several cytokines by monocyte/macrophages (M/M) obtained from HIV+ donors or after in vitro infection is severely dysregulated (21, 22, 23, 24, 25, 26, 27, 28). Of particular interest is the finding that IL-12 production by activated PBMC obtained from HIV+ patients or after in vitro infection of PBMC from HIV-uninfected (HIV) donors was decreased (25, 26, 27). In contrast, IL-10 production was reported to be increased in some studies (26, 28), but not in other (25). IL-12/IL-10 imbalance was also described in the plasma of HIV+ patients compared with HIV donors (29). Furthermore, an elevated IL-10/IL-12 ratio predicted the loss of type 1 cellular immunity in the feline immunodeficiency virus model (30). Therefore, we also compared cytokine and chemokine production between DC and PBMC from which they were derived to investigate whether DC obtained from HIV+ donors would exhibit the same cytokine imbalance that was reported for PBMC and M/M.

Blood samples were obtained from 15 HIV+ donors from Rush-Presbyterian-St. Luke’s Medical Center (Chicago, IL). They were shipped by overnight express to our National Cancer Institute laboratory in Bethesda, MD, where they were processed within 24 h of collection. These patients were receiving stable antiretroviral therapy, and their CD4 counts ranged from 0–1,297 cells/mm3 (four patients, <300; nine patients, 300–1,000; two patients, >1,000). Their viral loads ranged from undetectable (<500) to 273,100 copies/ml. Blood from 24 healthy adult HIV donors was obtained from the Department of Transfusion Medicine, National Institutes of Health (Bethesda, MD), and processed using the same conditions as the blood from HIV+ donors. The protocols were approved by the institutional review boards of all participating centers.

Soluble trimeric recombinant human CD40L (CD40LT) was provided by Immunex (Seattle, WA) (31). Recombinant human IL-4 (rhIL-4) and rhGM-CSF were obtained from PeproTech (Rocky Hill, NJ). All reagents were screened for low endotoxin levels using the Limulus amebocyte lysate assay (E-Toxate, Sigma, St. Louis, MO). Staphylococcus aureus Cowan (SAC) was purchased from Calbiochem-Behring (Pansorbin, La Jolla, CA).

PBMC.

PBMC were separated on lymphocyte separation medium (Lymphoprep, Organon Teknika, Rockville, MD) and resuspended at 3 × 106/ml in complete medium (RPMI 1640 containing 100 U/ml penicillin, 100 μg/ml streptomycin, 5 mM HEPES, and 2 mM glutamine).

Blood-derived DC.

DC were propagated from adult peripheral blood using a previously described protocol (11). Briefly, PBMC were suspended in DC medium at 5–8 × 106 cells/ml in 35-mm tissue culture plates (Costar, Cambridge, MA) for 2 h at 37°C. DC medium is composed of complete medium supplemented with 10% heat-inactivated FCS (HyClone, Logan, UT) and 5 × 10−5 M 2-ME (Life Technologies, Gaithersburg, MD). After this incubation, nonadherent cells were removed, and fresh DC medium was added to culture wells, supplemented with 1000 U/ml rhGM-CSF and 1000 U/ml rhIL-4. Cells were cultured for 7 days at 37°C at 7% C02. Half the volume of the medium was removed on alternate days and replaced with fresh medium supplemented with the two above mentioned cytokines.

On day 7 nonadherent cells were harvested, washed, resuspended at 108 cells/ml in washing medium (PBS containing 10% heat-inactivated FCS), and purified by negative selection. Cells were incubated with a mixture containing the following mouse anti-human Ab (each at 5 μg/ml): anti-CD3, -CD19, -CD16, and -CD14 (PharMingen, San Diego, CA), for 30 min on ice with gentle agitation. After this incubation, cells were washed three times in washing medium and incubated with magnetic beads coated with sheep anti-mouse IgG Abs (10 beads/cell; Dynal, Great Neck, NY), for 30 min on ice with gentle agitation. Unbound cells were separated through a series of washes using a magnetic particle concentrator (MCP-6, Dynal).

DC were washed twice in FACS buffer (balanced salt solution, 0.1% BSA, and 0.01% sodium azide), incubated with human IgG (20 μg/ml) for 10 min at 4°C to block Fc receptors, and stained with Ab that recognize the following cell surface markers: anti-CD1a-FITC, -CD40-FITC, -CD80-PE, -CD86-PE, -CD95-PE, -CXCR4-PE (all from PharMingen), and anti-CCR5 (Coulter/Immunotech, Miami, FL) or with isotype-matched Ab (γ1-FITC and γ2-PE, PharMingen), for 30 min at 4°C. The purity of the preparation was checked by staining with T cell, B cell, or M/M markers (CD3-PE, CD19-PE, and CD14-PE, PharMingen). The cells were then washed twice and resuspended in FACS buffer, and surface expression was determined by FACS analysis, using a Becton Dickinson FACScan and CellQuest software (Becton Dickinson, Mountain View, CA). Results are presented as the percentage of cells expressing a given marker compared with the isotype staining or as the mean fluorescence intensity (MFI).

Stocks of HIV-1 BaL, a CCR5-using strain, and HIV-1 IIIB, a CXCR4-using strain (Advanced Biotechnologies, Columbia, MD), were used in these experiments. DC were resuspended in DC medium supplemented with rhIL-4 and rhGM-CSF, at 106 cells/ml, and HIV-1 was added to cultures for overnight incubation. On day 0 (after overnight coincubation with HIV), DC were harvested, washed three times in washing medium, resuspended in DC medium supplemented with cytokines at the same cellular concentration, and cultured for 10–14 additional days (time point when maximum p24 production is measured). In addition, DC obtained from three different donors were infected and cultured for only 5 days, when p24 Ag could be first detected. These different time points were chosen according to previous kinetics experiments (11). On alternate days, half the medium was removed and replaced by fresh medium and cytokines. To assess DC infection by HIV, aliquots of these supernatants were inactivated with Triton X-100 (1% final concentration; Sigma, St. Louis, MO) and kept frozen for measurement of HIV-1 p24 protein levels by ELISA (Coulter, Miami, FL; detection limit, 8 pg/ml).

PBMC were stimulated with SAC (0.01% final concentration) at 37°C for 48 h at 1.5 × 106/ml in 48-well plates (Costar). DC were stimulated with SAC (0.01% final concentration) or CD40LT (10 μg/ml) at 37°C for 48 h at 0.4 × 106/ml in 48-well plates, in DC medium supplemented with rhIL-4 and rhGM-CSF. In vitro HIV-infected DC on days 10–14 postinfection were stimulated with CD40LT using the above mentioned conditions. After 48 h supernatants were harvested and frozen at −80°C before cytokine and chemokine analysis. IL-12 p70 and β-chemokines (RANTES, MIP-1α, and MIP-1β) were measured by ELISA, using R&D Systems kits (Minneapolis, MN). IL-10 was measured using PharMingen reagents. The detection limits of these ELISA are 4, 31.2, and 20 pg/ml for IL-12, chemokines, and IL-10, respectively. For relevant statistical processing of the data, all values below the detection limits were assigned an arbitrary value of half the detection limit.

To assess immunostimulatory functions, DC were tested for their ability to stimulate proliferation of purified allogeneic T cells. T cells were prepared from PBMC obtained from a healthy blood donor by negative selection, using the same protocol as that described for DC with a different mixture of mouse anti-human Ab: anti-CD14, -CD16, -CD19, and -CD1a (PharMingen), and were >90% CD3+. T cells were resuspended in freezing medium (FCS containing 10% DMSO; Sigma) and frozen in liquid nitrogen. At the time of testing DC for allogeneic stimulating activity, T cells were thawed and washed twice in DC medium. T cells (2 × 105) were resuspended in DC medium and cocultured with varying numbers of irradiated DC (3000 cGy, Cs137 source). Cultures were performed in triplicate in flat-bottom 96-well plates (Costar) and incubated at 37°C for 5 days. Cultures were pulsed with 1 μCi of [3H]thymidine overnight, and thymidine incorporation was detected using a beta counter. Results are expressed as the stimulation index = cpm (cultures containing DC + T cells)/cpm (cultures containing T cells alone).

Data were compared using nonparametric tests (Mann-Whitney test for unpaired samples or Wilcoxon signed rank test for paired samples). A p value <0.05 was considered significant.

DC from HIV donors were infected in vitro with a CCR5-using (R5-using) strain of HIV-1 (Ba-L; n = 8) or a CXCR4-using (X4-using) strain (IIIB; n = 7) (32). For these infection experiments, DC were >95% CD1a+ (results not shown). As previously reported, secreted HIV-1 p24 protein could be detected in the supernatants of infected cells, starting 4–5 days after infection, with increasing amounts occurring between days 10 and 14 postinfection (11). According to previous experiments, p24 Ag could be detected in 2–3% of these DC (11). Four Ba-L and two IIIB infections induced detectable, but relatively low, levels of p24 (<10 ng/ml). The remaining nine infections (four Ba-L and five IIIB) induced high levels of p24 (>10 ng/ml; cf., Fig. 1). As controls, uninfected cells from the same donors were cultured under the same conditions.

FIGURE 1.

Production of p24 after in vitro infection. DC from HIV donors were infected in vitro with Ba-L or IIIB viruses and cultured for 10–14 days postinfection. Every other day, p24 production was measured in the supernatants of infected cells by ELISA. Four Ba-L and 2 IIIB infections induced detectable, but relatively low, levels of p24 (<10 ng/ml). The remaining nine infections (four Ba-L and five IIIB) induced high levels of p24 (>10 ng/ml). One representative experiment of each situation is shown: open symbols represent Ba-L infections (one high and one low); filled symbols represent IIIB infections (one high and one low).

FIGURE 1.

Production of p24 after in vitro infection. DC from HIV donors were infected in vitro with Ba-L or IIIB viruses and cultured for 10–14 days postinfection. Every other day, p24 production was measured in the supernatants of infected cells by ELISA. Four Ba-L and 2 IIIB infections induced detectable, but relatively low, levels of p24 (<10 ng/ml). The remaining nine infections (four Ba-L and five IIIB) induced high levels of p24 (>10 ng/ml). One representative experiment of each situation is shown: open symbols represent Ba-L infections (one high and one low); filled symbols represent IIIB infections (one high and one low).

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At days 10–14, production of cytokine and chemokine was measured after stimulation with CD40LT. DC infected by either HIV-1 strain produced similar amounts of IL-12 and IL-10 as the uninfected control DC (all p > 0.17, by Wilcoxon signed rank test; cf., Table I). To exclude that the lack of differences observed between uninfected and infected cells was due to low levels of HIV infection achieved in some experiments, we performed the same analysis taking into account only infections inducing p24 production >10 ng/ml. Results were identical for both IL-12 and IL-10 and for both Ba-L and IIIB (all p > 0.07, by Wilcoxon signed rank test). Ba-L-infected DC also produced amounts of chemokines similar to uninfected DC (all p > 0.06; cf., Table I). By contrast, after CD40LT stimulation, production of MIP-1α and MIP-1β by IIIB-infected DC was significantly higher compared with that by uninfected DC (both p < 0.03; cf., Table I). Unstimulated IIIB-infected DC also tended to produce more MIP-1α and MIP-1β than unstimulated uninfected controls (MIP-1α, 0.6 vs 0.3 ng/ml; MIP-1β, 2.4 vs 0.4 ng/ml), although differences were not significant (both p = 0.14).

Table I.

CD40LT-stimulated production of cytokines and chemokines by DC infected in vitro with HIV-1 Ba-L or IIIBa

Median Production, ng/ml (range)
Uninfected (n = 9)Ba-L infected (n = 8)IIIB infected (n = 7)
IL-12 2.5 (0.6–5.8) 2.4 (1.6–4.6) 2.9 (1.5–3.7) 
IL-10 4.4 (0.5–14.7) 4.1 (0.3–18.5) 6.5 (1.8–9.8) 
RANTES 23.6b (1.5–46.3) 20.2c (1.0–49.7) 33.1 (7.2–50.4) 
MIP-1α 25.6b (1.0–164.8) 27.3c (0.8–92.0) 44.1d (27.2–190.5) 
MIP-1β 13.5b (2.3–40.4) 9.6c (2.0–21.8) 24.7d (4.3–51.9) 
Median Production, ng/ml (range)
Uninfected (n = 9)Ba-L infected (n = 8)IIIB infected (n = 7)
IL-12 2.5 (0.6–5.8) 2.4 (1.6–4.6) 2.9 (1.5–3.7) 
IL-10 4.4 (0.5–14.7) 4.1 (0.3–18.5) 6.5 (1.8–9.8) 
RANTES 23.6b (1.5–46.3) 20.2c (1.0–49.7) 33.1 (7.2–50.4) 
MIP-1α 25.6b (1.0–164.8) 27.3c (0.8–92.0) 44.1d (27.2–190.5) 
MIP-1β 13.5b (2.3–40.4) 9.6c (2.0–21.8) 24.7d (4.3–51.9) 
a

DC obtained from HIV donors (>95% CD1a+) were infected in vitro with HIV-1 Ba-L or IIIB. At days 10–14 postinfection, DC were stimulated with CD40LT for 48 h, and production of cytokine and chemokine was measured in the supernatants. As controls, uninfected cells from the same donors were cultured under the same conditions.

b

n = 7.

c

n = 6.

d

A significant difference between uninfected and HIV-infected cells (p < 0.05, Wilcoxon signed rank test).

We then addressed the question of whether production of cytokines and chemokines by CD40LT-stimulated DC was affected at an earlier time point postinfection. At day 5 postinfection, DC uninfected or infected by either HIV-1 strain produced similar amounts of IL-12 and IL-10 (median IL-12 production: 4.2, 4.0, and 6.1 ng/ml for uninfected, Ba-l-, and IIIB-infected, respectively; IL-10 production: 3.0, 3.8, and 2.0 ng/ml; all p > 0.29, by Wilcoxon signed rank test). Similar to that observed at the time of maximum p24 production, production of β-chemokines was not increased with BaL infection compared with that in uninfected controls (median MIP-1α production for uninfected and Ba-l-infected, respectively: 17.5 and 11.5 ng/ml; MIP-1β production: 10.2 and 9.9 ng/ml; RANTES production: 7.7 and 10.9 ng/ml; all p > 0.6). By contrast, the increased production of chemokines by IIIB-infected DC at days 10–14 was not observed at an earlier time point (median MIP-1α production for uninfected and IIIB-infected, respectively: 17.5 and 15.2 ng/ml; MIP-1β production: 10.2 and 9.2 ng/ml; RANTES production: 7.7 and 5.8 ng/ml; all p > 0.6).

DC infected for 10–14 days were stimulated with CD40LT for 48 h or were left unstimulated, and p24 production in supernatants from DC infected with Ba-L (n = 7) or IIIB (n = 7) was measured and compared with that in unstimulated HIV-infected DC. Production of p24 after CD40LT stimulation was significantly decreased in Ba-L-infected DC compared with unstimulated cells (cf., Fig. 2; p = 0.018, by Wilcoxon test). Interestingly, stimulation with CD40LT did not induce the same decrease in IIIB-infected DC (cf., Fig. 2; p > 0.99, by Wilcoxon test). A possible explanation for the differential effect of CD40LT stimulation on p24 release by DC infected with Ba-L or IIIB virus is that stimulation through the CD40-CD154 pathway differentially regulates chemokine receptor expression at the surface of DC. Therefore, we stimulated DC obtained from HIV donors for 48 h with CD40LT and analyzed CCR5 and CXCR4 expression as well as CD86 expression as a marker of CD40LT-induced maturation of DC. As shown in Fig. 3, expression of CCR5 was down-regulated on CD40LT-stimulated DC (∼8-fold decrease), whereas expression of CXCR4 was up-regulated (∼14-fold increase). As expected, expression of CD86 was strongly up-regulated on CD40LT-stimulated DC (∼28-fold increase).

FIGURE 2.

Effect of CD40LT stimulation on p24 production by HIV-infected DC. DC from HIV donors were infected in vitro with Ba-L or IIIB viruses and cultured for 10–14 days postinfection. On days 10–14 cells were stimulated for 48 h with CD40LT (10 μg/ml) or were left unstimulated, and p24 production was measured in those supernatants. The asterisk indicates a significant difference between stimulated and unstimulated cultures (p < 0.05, by Wilcoxon test).

FIGURE 2.

Effect of CD40LT stimulation on p24 production by HIV-infected DC. DC from HIV donors were infected in vitro with Ba-L or IIIB viruses and cultured for 10–14 days postinfection. On days 10–14 cells were stimulated for 48 h with CD40LT (10 μg/ml) or were left unstimulated, and p24 production was measured in those supernatants. The asterisk indicates a significant difference between stimulated and unstimulated cultures (p < 0.05, by Wilcoxon test).

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FIGURE 3.

Modulation of expression of CCR5 and CXCR4 on the surface of DC by CD40LT stimulation. DC from an HIV donor were stimulated for 48 h with CD40LT (10 μg/ml). Cells were harvested and stained with anti-CD1a Ab (FITC labeled). They were also stained with isotype Ab (fine line, open histogram), anti-CD86 or anti-CXCR4 Ab, or anti-CCR5 Ab (all PE labeled; filled histograms). CD1a+ cells were gated, and expressions of CD86, CXCR4, and CCR5 were analyzed in the gated population. Numbers represent the corrected mean fluorescence intensity (ΔMFI), defined as ΔMFI = MFI Ab − MFI isotype). Data are representative of three donors.

FIGURE 3.

Modulation of expression of CCR5 and CXCR4 on the surface of DC by CD40LT stimulation. DC from an HIV donor were stimulated for 48 h with CD40LT (10 μg/ml). Cells were harvested and stained with anti-CD1a Ab (FITC labeled). They were also stained with isotype Ab (fine line, open histogram), anti-CD86 or anti-CXCR4 Ab, or anti-CCR5 Ab (all PE labeled; filled histograms). CD1a+ cells were gated, and expressions of CD86, CXCR4, and CCR5 were analyzed in the gated population. Numbers represent the corrected mean fluorescence intensity (ΔMFI), defined as ΔMFI = MFI Ab − MFI isotype). Data are representative of three donors.

Close modal

Because in vitro HIV infection of DC did not alter their capacity to produce cytokines upon stimulation (our results), their phenotypic characteristics (18), or their capacity to induce an allogeneic response (11), we investigated whether DC derived from PBMC from HIV+ patients would also have similar characteristics to DC derived from PBMC from HIV+ donors. Therefore, DC were obtained by culture of plastic-adherent PBMC from HIV+ and HIV donors for 7 days in the presence of rhGM-CSF and rhIL-4. In the case of HIV+ donors, samples of 50–100 ml of peripheral blood were drawn, and 0.2–3.8 × 106 DC were obtained after culture and negative selection. The median yield of DC per unit of PBMC was not significantly different between HIV+ and HIV donors (2 × 106 and 1.8 × 106 DC/100 × 106 PBMC for HIV+ and HIV donors, respectively; p = 0.89, by Mann-Whitney test). Cells were >75% CD1a+ bright (results not shown), and, importantly, the rest of the cells did not express T, B, or M/M lineage markers (<3% CD3+ and CD19+ cells, and <1% CD14+), and were CD1a+ dull. In five different experiments, DC were derived in parallel from PBMC obtained from one HIV+ and one HIV donor, and expression of several surface markers was analyzed. DC presented with a low level of CD80 and high levels of CD86, CD40, and CD95 expression as previously described (11). No differences in phenotype were observed between cells derived from HIV+ and HIV donors (cf., Fig. 4; all p > 0.27, by Wilcoxon signed rank test). To test for Ag-presenting function, we measured the capacity of DC to induce an allogeneic response. DC from both HIV+ and HIV donors induced a strong dose-dependent stimulation of allogeneic T cells (cf., Fig. 5); this result is in agreement with the recent study of APC function of monocyte-derived DC obtained from HIV+ patients (17).

FIGURE 4.

Phenotypic profile of DC from HIV+ and HIV patients. Monocyte-derived DC were stained with Ab that recognize the following cell markers: anti-CD1a-FITC, -CD40-FITC, -CD80-PE, -CD86-PE, and -CD95-PE, or with isotype-matched Ab (γ1-FITC and γ2-PE), for 30 min at 4°C. Surface expression was determined by FACS analysis. Each experiment consisted of one HIV+ donor and one HIV donor, whose DC were derived and stained in parallel. Data are representative of five experiments. No differences in phenotype were observed between cells derived from HIV+ and HIV donors (all p > 0.05, by Wilcoxon signed rank test).

FIGURE 4.

Phenotypic profile of DC from HIV+ and HIV patients. Monocyte-derived DC were stained with Ab that recognize the following cell markers: anti-CD1a-FITC, -CD40-FITC, -CD80-PE, -CD86-PE, and -CD95-PE, or with isotype-matched Ab (γ1-FITC and γ2-PE), for 30 min at 4°C. Surface expression was determined by FACS analysis. Each experiment consisted of one HIV+ donor and one HIV donor, whose DC were derived and stained in parallel. Data are representative of five experiments. No differences in phenotype were observed between cells derived from HIV+ and HIV donors (all p > 0.05, by Wilcoxon signed rank test).

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FIGURE 5.

Capacity of DC to induce an allogeneic response. Purified T cells (2 × 105) from one HIV donor (>90% CD3+) were resuspended in DC medium and cocultured with varying numbers of irradiated DC obtained from HIV+ and HIV donors. Two HIV+ donors (both donors had CD4 counts between 300 and 1000, were receiving combination antiretroviral therapy, and had viral loads ∼4000 copies/ml) and two HIV donors were studied. After 5 days cultures were pulsed with [3H]thymidine, and incorporated radioactivity was measured using a beta counter. Results are expressed as the stimulation index [SI = cpm (cultures containing DC + T cells)/cpm (cultures containing T cells alone)]. Incorporated radioactivity in unstimulated cultures was <1200 cpm.

FIGURE 5.

Capacity of DC to induce an allogeneic response. Purified T cells (2 × 105) from one HIV donor (>90% CD3+) were resuspended in DC medium and cocultured with varying numbers of irradiated DC obtained from HIV+ and HIV donors. Two HIV+ donors (both donors had CD4 counts between 300 and 1000, were receiving combination antiretroviral therapy, and had viral loads ∼4000 copies/ml) and two HIV donors were studied. After 5 days cultures were pulsed with [3H]thymidine, and incorporated radioactivity was measured using a beta counter. Results are expressed as the stimulation index [SI = cpm (cultures containing DC + T cells)/cpm (cultures containing T cells alone)]. Incorporated radioactivity in unstimulated cultures was <1200 cpm.

Close modal

Monocyte-derived DC produce IL-12 and IL-10 following stimulation through the CD40-CD154 pathway or by bacterial products (33, 34). Therefore, we investigated whether cytokine production by DC from 12 HIV+ patients would be defective compared with that by cells obtained from 13 HIV donors. DC were cultured without stimulation, with CD40LT, or with SAC. Unstimulated DC from HIV+ and HIV donors did not produce detectable amounts of IL-12 and produced low levels of IL-10. After stimulation, DC from all HIV+ donors produced large amounts of IL-12 (cf., Fig. 6). This production was similar to that by cells from HIV donors (both p > 0.33, by Mann-Whitney test). Production of IL-10 by DC from the HIV+ and HIV donors was also similar after CD40LT or SAC stimulation (cf., Fig. 6; both p > 0.21).

FIGURE 6.

Cytokine production by DC and PBMC from HIV+ and HIV patients. Box plot analysis of IL-12 and IL-10 production by DC from HIV+ patients (n = 13) or HIV patients (n = 12). Production of cytokines by SAC-stimulated PBMC from nine of these HIV+ donors and eight of these HIV donors was analyzed. The horizontal bars correspond to the median (50th percentile of the variable), the box limits correspond to the 25th and 75th percentiles of the variable, and the vertical bars correspond to the 10th and 90th percentiles. Values above the 90th and below the 10th percentile are plotted as points. ∗, Significant difference between HIV+ and HIV donors (p < 0.05, by Mann-Whitney test).

FIGURE 6.

Cytokine production by DC and PBMC from HIV+ and HIV patients. Box plot analysis of IL-12 and IL-10 production by DC from HIV+ patients (n = 13) or HIV patients (n = 12). Production of cytokines by SAC-stimulated PBMC from nine of these HIV+ donors and eight of these HIV donors was analyzed. The horizontal bars correspond to the median (50th percentile of the variable), the box limits correspond to the 25th and 75th percentiles of the variable, and the vertical bars correspond to the 10th and 90th percentiles. Values above the 90th and below the 10th percentile are plotted as points. ∗, Significant difference between HIV+ and HIV donors (p < 0.05, by Mann-Whitney test).

Close modal

Chemokine production by DC from 12 HIV+ and 10 HIV donors was also measured. Unstimulated DC from HIV+ and HIV donors produced low levels of RANTES, MIP-1α, and MIP-1β. After stimulation, DC from all HIV+ donors produced large amounts of all three chemokines (cf., Fig. 7). Production of chemokines was similar to that by cells from HIV donors after CD40LT or SAC stimulation (cf., Fig. 7; all p > 0.1).

FIGURE 7.

Chemokine production by DC and PBMC from HIV+ and HIV patients. Box plot analysis of RANTES, MIP-1α, and MIP-1β production by DC from HIV+ patients (n = 12) or HIV patients (n = 10). Production of cytokines by SAC-stimulated PBMC from nine of these HIV+ donors and eight of these HIV donors was analyzed. No significant difference between HIV+ and HIV donors was observed (all p > 0.05, by Mann-Whitney test).

FIGURE 7.

Chemokine production by DC and PBMC from HIV+ and HIV patients. Box plot analysis of RANTES, MIP-1α, and MIP-1β production by DC from HIV+ patients (n = 12) or HIV patients (n = 10). Production of cytokines by SAC-stimulated PBMC from nine of these HIV+ donors and eight of these HIV donors was analyzed. No significant difference between HIV+ and HIV donors was observed (all p > 0.05, by Mann-Whitney test).

Close modal

p24 HIV protein production was also measured in the supernatants of DC derived from 10 of these HIV+ patients. Interestingly, no p24 was detectable (<8 pg/ml) in any of the unstimulated or CD40LT- or SAC-stimulated cultures (results not shown), even in the case of two patients who had high viral loads (>250,000 copies/ml).

Production of cytokines by SAC-stimulated PBMC from nine HIV+ donors and eight HIV donors, whose cells were used to derive DC, was analyzed in parallel. In contrast to the results obtained with DC, IL-12 production by PBMC was decreased in HIV+ donors compared with HIV donors (cf., Fig. 6; p = 0.006, by Mann-Whitney test). IL-10 production by PBMC was also significantly increased compared with that in HIV donors (cf., Fig. 6; p = 0.05). Interestingly, SAC-stimulated IL-12 and IL-10 production was greater by DC than by PBMC of both HIV+ and HIVdonors. β-Chemokine production by PBMC after SAC stimulation was also measured. In contrast to what was observed for cytokine production, chemokine production was similar between HIV+ and HIV donors (cf., Fig. 7; all p > 0.14, by Mann-Whitney test).

The primary objective of the present study was to analyze the cytokine and chemokine secretion profiles of monocyte-derived DC from HIV+ patients and from HIV individuals after in vitro infection with HIV. HIV+ patients whose cells were analyzed were receiving stable antiretroviral therapy, comprising two to four drugs, and their CD4 counts ranged from 0–1,297 cells/mm3 (four patients, <300; nine patients, 300-1000; two patients, >1000). Viral loads detected in their plasma ranged from undetectable (<500) to 273,100 copies/ml. Our results show no defect in the ability of these DC to produce IL-12, IL-10, and β-chemokines, even in the case of DC obtained from AIDS patients. To our knowledge this is the first study investigating the cytokine and chemokine secretion profile of DC obtained from HIV+ patients or DC infected in vitro with different HIV-1 strains. In addition, the cells from HIV+ patients were morphologically and phenotypically comparable to DC from healthy donors, expressing similar high levels of costimulatory molecules. The DC obtained from HIV+ donors also had potent allogeneic stimulating capacity, a defining characteristic of DC (2). Furthermore, DC obtained from healthy donors and infected in vitro with HIV-1 had characteristics similar to those of uninfected DC obtained from the same donors.

The extent to which DC remain phenotypically and functionally intact during HIV infection is still controversial. The various methods of isolation and maturation of DC may result in the study of different cell populations, and this variability may lead to discrepancies in the reports of their numbers, infection levels, and functional capacities. No decrease in numbers of HLA-DR+ cells or in levels of surface expression of HLA-DR or costimulatory molecules was reported on Langerhans cells or spleen DC obtained from HIV+ patients or after in vitro infection of blood-derived DC (14, 18, 19). In addition, using samples from monozygotic twins discordant for HIV infection or sequential samples from the same patients at different stages of disease progression, no defect was observed in the ability of Langerhans cells or blood-derived DC to present recall Ag to CD4+ or CD8+ T cells (15, 16). In contrast, a reduction in the number and function of blood DC, which was particularly severe in AIDS patients, has been reported (35). These deficient parameters were observed to improve in zidovudine-treated patients (36). Thus, differences in duration and efficiency of antiviral drug therapy of patients whose DC functions are analyzed could also contribute to discrepant results.

Monocyte-derived DC generated from cells of HIV+ patients may serve as vehicles for enhancing the reactivity of secondary memory T cells to HIV-1 (16). Therefore, our results showing that those cells have a normal cytokine profile and, in particular that they can produce large amounts of IL-12 have significant potential implications. In parallel to our DC studies, PBMC were obtained from the same HIV+ and HIV donors whose cells were used to derive DC. In contrast to DC, PBMC from HIV+ patients demonstrated cytokine imbalance that we and others have described. Specifically, decreased production of IL-12 and increased production of IL-10 were observed after stimulation (26, 27, 28, 37, 38, 39). Several studies have shown that IL-12 production by PBMC can be restored in vitro in most HIV+ donors to a level similar to that in HIV donors (27, 39). Of particular interest, was the observation that pretreatment of PBMC from HIV+ patients with IL-4 or IL-13 primed these cells for enhanced production of IL-12 in response to SAC (40). Thus, the phase of differentiation of monocytes into DC, by culturing cells in the presence of high doses of hIL-4 and hGM-CSF could have primed DC from HIV+ donors to produce normal IL-12 levels. In vitro infection of M/M with HIV leads to decreased IL-12 production (25, 26), increased IL-10 production (41), and increased expression of Fas ligand molecules (42). In contrast, in vitro infection of DC does not induce any of these changes (our results, (42)), highlighting the differential effects of HIV infection on DC and M/M. A similar discrepancy between DC and M/M has been reported in the murine model of Leishmania major infection; infection of M/M with amastigotes inhibited IL-12 production (43, 44, 45). In contrast, amastigotes, in the presence of IFN-γ, induced fetal skin-derived DC to release IL-12 (44).

In addition, no changes in the capacity of DC obtained from HIV+ donors to produce β-chemokines (RANTES, MIP-1α, or MIP-1β) was detected. Interestingly, the amount of β-chemokines produced by stimulated DC (10–100 ng/ml, depending on the chemokine) was similar to that produced by activated M/M. These β-chemokines have been reported to inhibit T cell infection by R5-using HIV-1 (46). Those results suggest that in addition to their potential role to induce or enhance T cell responses, activated DC might represent a major source of the HIV-1-suppressive β-chemokines. Ba-L-infected DC produced the same amount of chemokines as uninfected controls. In contrast, IIIB-infected DC produced significantly more MIP-1α or MIP-1β than uninfected controls. Similar results have been reported after ex vivo HIV infection of tonsils; endogenous MIP-1α, MIP-1β, and RANTES production is up-regulated following infection with X4-using, but not with R5-using, HIV (47). However, the cellular sources of these chemokines were not characterized in the tonsil model.

Interestingly, CD40LT stimulation of infected DC had a differential effect on p24 release by these cells depending on the type of virus. It significantly decreased p24 release from Ba-L-infected DC, while it had no effect on IIIB-infected DC. One explanation of this differential effect of CD40LT is that stimulation through the CD40-CD154 pathway differentially regulates chemokine receptor expression at the surface of DC. Accordingly, our results showed that expression of CCR5 was down-regulated on CD40LT-stimulated DC, whereas expression of CXCR4 was up-regulated. These results are in agreement with recent studies of chemokine receptor expression on DC and M/M (48, 49). An alternative, but nonexclusive, explanation is that chemokines produced following CD40LT stimulation were able to block DC infection with an R5-using virus, whereas they had no effect on an X4-using virus, similar to stimulated M/M (46, 50).

These results suggested that stimulated APC could limit transmission of R5-using virus to CD4+ T cells, in contrast to earlier work in which CD40/CD154 stimulation enhanced replication of X4-using virus in DC cocultures with CD4+ T cells (8). However, it was recently reported that addition of CD40LT to cocultures of naive CD4+ T cells and autologous DCs infected with an R5-using isolate caused a reduction in RT activity, whereas its addition to cocultures of T cells and DC infected with an X4-using virus caused the inverse result, i.e., an increase in RT activity (51). Therefore, activation of APC could create a more favorable environment for X4-using strains and a less favorable one for R5-using viruses, and thus accelerate the transition from R5- to X4-using virus. However, caution should be exercised in patients with advanced disease, who might harbor predominantly X4-using viruses. No detectable p24 production could be measured in the DC preparations we analyzed, and a recent study showed the absence of HIV-1 DNA in cytokine-derived DC obtained from 13 asymptomatic patients (17). Nevertheless, the existence of a small number of DC infected with X4-using virus cannot be completely ruled out in the monocyte-derived DC, and activation of those DC could lead to an enhanced transmission of virus to T cells.

Monocyte-derived DC could constitute a source of potent APC and be used to stimulate naive CD4 and CD8 T cell-mediated responses in HIV+ patients or to enhance the reactivity of secondary memory T cells to HIV and non-HIV Ag (16, 52, 53). Moreover, recent studies conducted in the mouse showed that CD40L-stimulated DC can prime CTL, even in the absence of CD4 lymphocytes (54, 55, 56). Procedures recently introduced permit the efficient generation of DC from a limited amount of peripheral blood (20). Our results show that this approach can be applied to obtain DC from HIV+ patients who have a wide range of CD4+ T cell counts, including AIDS patients. These monocyte-derived patient DC have a normal cytokine and chemokine profile, express normal and high levels of costimulatory molecules, and do not produce p24 viral Ag. Because a recent study has shown that cytokine-derived DC obtained from HIV+ donors have strong APC function (17), our findings of normal cytokine and chemokine production by these cells raise the possibility of ex vivo differentiation and expansion of DC from precursors present in the blood of HIV+ patients receiving effective anti-retroviral therapy. These autologous DC might then be pulsed with specific Ag and introduced back into the patients as effective adjuvants for active immunotherapy.

We thank PeproTech, Inc. (Rocky Hill, NJ) for the recombinant human GM-CSF and IL-4. We also thank Drs. Geneviève Milon, David Sacks, and Dinah Singer for their critical review of this manuscript.

2

Abbreviations used in this paper: DC, dendritic cells; M/M, monocytes/macrophages; CD40LT, recombinant trimeric human CD40 ligand; rhIL-4, recombinant human IL-4; SAC, Staphylococcus aureus Cowan; MFI, mean fluorescence index; MIP-1β, macrophage inflammatory protein-1β.

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